- Email: [email protected]
Production of pectin-whey protein nano-complexes as carriers of orange peel oil
Carbohydrate Polymers 177 (2017) 369–377
Contents lists available at ScienceDirect
Carbohydrate Polymers journal homepage: www.elsevier.com/locate/carbpol
Production of pectin-whey protein nano-complexes as carriers of orange peel oil
T
⁎
Sanaz Ghasemi, Seid Mahdi Jafari , Elham Assadpour, Morteza Khomeiri Faculty of Food Science and Technology, Gorgan University of Agricultural Sciences and Natural Resources, Gorgan, Iran
A R T I C L E I N F O
A B S T R A C T
Keywords: Nanocomplex Orange peel oil Encapsulation Whey protein Pectin
Orange peel oil is one of the most common flavorings used in the food industry which is volatile under environmental conditions. Encapsulation is the best way to protect it and control its release. One of the nanoencapsulation systems for food bioactive ingredients is complexation method, which entraps the core materials in a complex of two different biopolymers. In this study, orange peel oil was nanoencapsulated by pectinwhey protein nanocomplexes. After determining the optimum nanocomplex suspensions containing orange peel oil based on the stability, viscosity, and color, they were formulated in three different pH values (3, 6 and 9) and converted into powdered forms by freeze drying. The analysis of size and zeta potential of nanocomplexes revealed that the smallest particles formed in pH = 6. The encapsulation efficiency of the powders at pH = 3, 6 and 9 were 88, 84, and 70%, respectively and there was a reverse linear correlation between encapsulation efficiency and the color index (b*). The microstructure and the morphology of the nanocomplex powders was investigated by SEM and AFM and the results showed that more spherical particles are formed in pH = 3. FTIR analysis determined that there was a chemical reaction and bond formation between whey proteins and pectin as a sharp band was appeared in 991 cm−1.
1. Introduction Flavor is known as the essence of foods and plays an important role in consumer satisfaction and consumption of foods. Most liquid food flavorings are volatile substances and chemically unstable when exposed to the air, light, moisture and high temperatures. Orange peel oil is one of the most widely used flavorings in the food industry. It is a mixture of various chemical components and is mainly composed of Dlimonene (> 90%). D-Limonene is the base sensory character of the citrus oils and due to its instability under the process and environmental conditions, it is necessary to encapsulate it prior to use in foods or beverages to limit aroma degradation or losses during processing and storage (Jun-xia, Hai-yan, & Jian, 2011; Madene, Jacquot, Scher, & Desobry, 2006). Encapsulation is a quickly expanding technology which has been generally used for many food applications such as taste and odor covering in the form of micro- and nanoparticles. This technology can control the release of bioactive ingredients under specific conditions, so causing a long shelf-life. One of the nanoencapsulation systems for protection of flavors is complexation method which entraps the flavor oils in a complex of two different biopolymers. In general, these biopolymers are proteins and polysaccharides (Yeo, Bellas, Firestone,
⁎
Corresponding author. E-mail address: [email protected] (S.M. Jafari).
http://dx.doi.org/10.1016/j.carbpol.2017.09.009 Received 4 July 2017; Received in revised form 16 August 2017; Accepted 5 September 2017 Available online 06 September 2017 0144-8617/ © 2017 Elsevier Ltd. All rights reserved.
Langer, & Kohane, 2005) which can join together through electrostatic attractions. Polysaccharide-protein nanoparticles are taken more into consideration than pure single biopolymer nanoparticles, because of their higher chemical and colloidal protection. Proteins often have a negative charge in the pH values above their isoelectric points (pI ≈ 5) and positive charge below this pH value. So, when proteins and polysaccharides are mixed together in a liquid medium, two phenomena can occur: 1) Higher rate of the attractive interactions resulting in formation of soluble and insoluble complexes; 2) Higher rate of the repulsive interactions causing separation of the two biopolymers from each other (Weinbreck, Nieuwenhuijse, Robijn, & de Kruif, 2004). In practice, one of these two phenomena will occur depending on the charge of both biopolymers and effective factors such as pH and ionic strength. The absorption interactions, further occurs through the creation of electrostatic bonds between oppositely charged biopolymers such as proteins with the positive charges (in pH < pI) and anionic polysaccharides containing carboxyl groups, phosphates and sulfates (at a pH > pKa), or negatively charged proteins (pH > pI) and cationic polysaccharides such as chitosan (Matalanis, Jones, & McClements, 2011). This complex formation is guided by biopolymer parameters including charge density of the biopolymer, its molecular weight, chemical nature; environmental conditions including ionic strength, ion
Carbohydrate Polymers 177 (2017) 369–377
S. Ghasemi et al.
2. Materials and methods
type, biopolymer concentration, biopolymer ratio, pH; and processing factors, including temperature, pressure, shearing rate and time which may affect the formation and stability of the complexes (Ye, 2008). The complexation of proteins and polysaccharides such as β-lactoglobulin and pectin can be applied to encapsulate hydrophilic and hydrophobic nutraceuticals including different vitamins, flavors, essential oils, fatty acids, etc. (Zimet & Livney, 2009). When formulating these biopolymer complexes by high energy densities, it results in nano sized complexes which are optically transparent with higher surface-to-volume ratio and could protect the encapsulated nutraceuticals from chemical degradation more efficiently. Ultrasonication is one of these high energy devices which can result in nano-sized droplets and particles through cavitation and it has been successfully applied in preparation of nanoemulsions and nanodispersions loaded with different bioactive ingredients and drugs (Tang, Shridharan, & Sivakumar, 2013; Tang, Sivakumar, & Nashiru, 2013; Tan Khang, Tang Siah, Thomas, Vasanthakumari, & Manickam, 2016). Milk proteins, mostly whey proteins, have been extensively used in the studies of protein– polysaccharide complex systems (Livney, 2010). On the other hand, pectin is an anionic hetero polysaccharide associated with the cell wall in plants and its main chain has been formed by galacturonic acid units with α-D connector (1–4) and esterified partially with methanol. Commercial low methoxyl pectin (LMP) is an anionic polysaccharide obtained from de-esterification of high methoxyl pectin (Mendanha et al., 2009). Bédié et al. (2008) investigated the influence of protein to pectin ratios, pH and pre or post-blending acidification on the formation of thiamin-loaded complexes. They blended LMP and whey protein isolate (WPI) solutions to achieve protein to pectin ratios of 2:1, 1:1 and 1:2 at a total biopolymer concentration of 0.4% (w/w) with pH adjustment in the range of 2.5–4.0 either before or after blending. Their results showed that at a ratio of 2:1 (protein: pectin), the size of complex was constant and lower at post-blending acidification compared to preblending. But at a ratio of 2:1 (WPI:pectin), the optimum pH for maximum turbidity and complex yield was around 3.0 with both pre- and post-blending acidifications. Anal et al. (2008) studied inter-macromolecular complexation between sodium caseinate and chitosan in aqueous solutions as a function of pH (3.0–6.5). The chitosan–caseinate complexes formed in the pH range 4.8–6.0 (with opposite charges) were stable and soluble. The soluble complexes associated to form larger particles at higher concentrations of chitosan (0.15 wt%). It was revealed that the particles formed by the chitosan- caseinate complexes at pH 4.8–6.0, had sizes between 250 and 350 nm. At pH values above 6.0, the nano-complex particles attached together and formed larger particles, due to phase separation. In the last couple of years, some other studies have been conducted about production and application of nano complexes of different proteins with polysaccharides such as WPI – beet pectin (Arroyo-Maya & McClements, 2015), carboxy methyl chitosan – soy protein (Teng, Luo, & Wang, 2013), WPC – pectin (Mohammadi, Jafari, Assadpour, & Esfanjani, 2016), WPI – gum Arabic (Bosnea, Moschakis, & Biliaderis, 2014) and WPI – high-methoxyl pectin (Wagoner & Foegeding, 2017). Our main goal in this study was producing orange peel oil-loaded nanocapsules based on complexed biopolymers of whey protein concentrate and low methoxyl pectin to encapsulate this popular flavor and introduce a new formulation of orange peel oil for applications in the food industry, particularly in beverages. We decided to answer this question whether is it possible to produce pectin-WPC nanocomplex carriers for successful loading of orange peel oil or not? Also, we worked on the influence of various factors including the concentration of pectin and whey protein concentrate and pH values on the complex formation, in order to find the optimal conditions for producing best nanocomplexes. The final goal was to evaluate the microstructure and physicochemical properties of freeze-dried powder particles containing orange peel oil-loaded nanocomplexes.
2.1. Materials The applied biopolymers were an aqueous solution of whey protein concentrate (WPC; 80 w/w%) in a combination with maltodextrin (MD; DE 16–20) and pectin (LMP). Whey protein concentrate and maltodextrin were obtained from Arla (Denmark) and Qinhuangdao Starch Co. (China), respectively. Low methoxyl pectin from citrus peel (galacturonic acid, ≥74.0% (dried basis), DE = 25%) was purchased from Sigma Chemicals Company (USA). Orange peel oil was purchased from Ramsar Citrus Concentrate Co. (Iran). A non-ionic surfactant, Tween 80 (Sigma) was used as the emulsifying agent during emulsion preparation stages of this work. Deionized water was used to prepare all the solutions. 2.2. Biopolymer solution preparation Firstly, aqueous solutions of WPC were prepared by dispersing 4, 6 and 8 g of WPC powder into deionized water to obtain 100 ml solutions. Also, aqueous solutions of pectin were prepared by dispersing 0.5, 0.75, and 1 g LMP powder into hot deionized water (70 °C) to obtain 100 ml solutions. Then, 50% w/w maltodextrin solutions were prepared as a filler to increase the total solids for preparation of powders. Solutions were stirred for 30 min on a magnetic stirrer (IKA, Germany) then they were stored overnight at 4 °C for complete hydration of biopolymers (Assadpour, Maghsoudlou, Jafari, Ghorbani, & Aalami, 2016a, 2016b). 2.3. Preparation of WPC-Pectin nano-complexes containing orange peel oil WPC and pectin solutions were mixed together along with maltodextrin at constant ratio of 50%w/w with the same volume of each solution for 30 min on a magnetic stirrer. Then, Tween 80 at a ratio of 10% of the total solids was added into the solution and mixed again to dissolve completely. Finally, orange peel oil was added into these prepared solutions gradually during homogenization by an ultrasonic probe (Iranian Ultrasonic Technology Company, 400 W, 20kHZ, 12 mm probe diameter) at 25 °C and power of 350 W for 10 min. pH of the solutions was adjusted to 3, 6, 9 using HCl and NaOH (1 M) (Jafari, He, & Bhandari, 2007a; Lutz, Aserin, Wicker, & Garti, 2009). 2.4. Physicochemical properties of nano-complex solutions After preparing the complexes, the effect of composition and preparation conditions of biopolymer solutions on viscosity, color (L*) and stability were investigated. Viscosity was measured by using a Brookfield viscometer (LVDV Pro II, Brook-field Engineering Laboratories, spindle S00, USA), L* was measured by image analysis using Image J software and also for measuring the stability of the nanocomplexes, accelerated method by a centrifuge (Sigma Co., 3K30 model) in 20,000 g and 25 °C for 30 min was applied (Hosseini, EmamDjomeh, Sabatino, & Van der Meeren, 2015; Zimet & Livney, 2009). 2.5. Particle size and zeta potential measurement Particle size and zeta potential (surface electrical charge) of prepared nano-complexes were measured using a dynamic light scattering (DLS) method (Zeta Sizer, Malvern Instruments, Malvern, UK). To avoid multiple scattering, the samples were diluted with distilled water and at a temperature of 25 °C, the average particle size (based on volume, number, and intensity mean diameter), and zeta potential were determined. 2.6. Stability of nano-complex solutions For performing the stability test, 50 ml of nanocomplex solutions 370
Carbohydrate Polymers 177 (2017) 369–377
S. Ghasemi et al.
(primary nanocomplex volume) was centrifuged and then, the volume of remained stable nanocomplex was determined by a graduated cylinder. Then, the percentage of stability was calculated with the following equation.
Stability (%) =
The volume of nanocomplex remained stable × 100 The volume of primary nanocomplex
ρ ε = 1 − ⎜⎛ b ⎞⎟ ⎝ ρa ⎠ ρa = absolute density, ρb = bulk density.
2.8.6. Solubility For measuring the solubility, 1 g of powder was added to 100 ml of distilled water under stirring with a magnetic stirrer at 385 rpm for 5 min. The resulting dispersion was centrifuged at 3000g for 5 min. 25 ml of the upper solution was transferred to a Petri dish that has already been weighed and dried in an oven at temperature of 105° C for 5 h. The percentage of solubility was determined based on the amount of powder dissolved in water.
(1)
2.7. Freeze-drying of the orange peel oil-loaded nano-complexes The optimum prepared nano-complexes were frozen in flasks at −20 °C for 48 h. Frozen samples were dried using a freeze dryer (FDB5503 Freeze-dryer; Operon, South Korea). The freeze-dried samples were stored in hermetic containers at 4 °C. For further experiments, the dried nano-complexes were grinded into a suitable powder by a pestle and mortar.
2.8.7. Surface color analysis The surface color of powders, which is relevant to the surface component content (orange peel oil), was evaluated by the procedure of Khazaei et al. (2014) with some modifications. The yellow (b*) index at the powder surface was measured because of yellow–orange color of orange peel oil. For all samples, color was measured using image analysis software (Image j). For evaluating the surface color of the powders, 10 g of the powders was transferred into a Petri dish and was placed at the scanner (HP, Scanjet G3110, UK) and images were taken. Subsequently, powders color was analyzed by using Image J software and b* factor of the samples was measured.
2.8. Physicochemical properties of the prepared powders containing nanocomplexes The physicochemical properties of powders including solubility, water activity, moisture content, bulk density, absolute density, porosity, and surface color were measured. 2.8.1. Moisture About 2 g of the powder was placed in a Petri dish for 2–3 h in an oven (UFB 400, Memert, Germany) at temperature of 105 °C, then cooled in a desiccator, and weighed. The moisture content was calculated using Eq. (2).
M% =
W2 − W3 × 100 W2 − W1
2.9. FTIR analysis The structure analysis of the WPC–pectin nanocomplex samples were investigated by Fourier Transform Infrared Spectroscopy (FTIR). In addition, the IR spectra of the samples were registered by FTIR spectrophotometer (Spectrum RX1, PerkinElmer, US) using attenuated total reflection technique. For structure analysis, at first the powder samples were mixed with KBr powder in 1–10 ratios and pressed into a disk before spectrum acquisition. The spectrum was scanned in transmission mode from 400 to 4500−1 cm range.
(2)
M = moisture (%), W1 = weight of petri dish, W2 = the total weight of petri dish and powder, W3 = the total weight of dried powder and petri dish after processing (oven). 2.8.2. Water activity (aw) For determination of water activity, we applied instrumental analysis by Novasina LabMaster Std Labmaster Standard Water Activity Instrument (UK).
2.10. Morphological characterization of nano-complexes The morphology of the powders loaded with orange peel oil was observed by scanning electron microscopy (SEM) (Vega II, Tescan, Czech Republic). The powder was spread on a clean glass plate and then coated with a thin layer of gold metal under high vacuum and observed. Representative SEM images were described (Hosseini, Zandi, Rezaei, & Farahmandghavi, 2013). Atomic force microscope (AFM) (DualScopeTM DS95-50, DME, Denmark) was used for morphological characterization and particle size and size distribution of WPC-pectin nano-complexes. Samples for AFM imaging were prepared by applying a drop of diluted nano-complexes suspension (0.05 mg/mL) on a clean glass surface, spread and dried at room temperature for 30 min. Then the image analysis by AFM was performed (Peinado, Lesmes, Andrés, & McClements, 2010).
2.8.3. Bulk density The bulk density of the powders was measured by weighing 2 g of powder and placing it in a 25 ml graduated cylinder. Then the cylinder was tapped 100 times by hand and the bulk density was calculated from the ratio between the mass of powder (2 g) and the volume occupied in the cylinder (Assadpour and Jafari, 2017). 2.8.4. Absolute density To calculate absolute density of powders, we applied displacement method through pycnometer. Toluene volume (Vp) was obtained Using Eq. (3) and the absolute density was calculated using Eq. (4) (Chegini & Ghobadian, 2005).
Vp =
2.11. Encapsulation efficiency of orange peel oil
(Mpt − Mp) − (Mpst − Mps ) ρt
It was necessary to analyze encapsulated nanocomplex powders in terms of total content and surface content of orange peel oil in final powders. For surface (unencapsulated) content, 0.5 g of each sample was dispersed in 20 ml hexane, vortexed for 2 min and filtered using Whatman paper no. 41 and the orange peel oil content was determined spectrophotometrically at 231 nm (Matias, Ribeiro, Sarraguça, & Lopes, 2014). For total content, 0.5 g of powders was dissolved in 20 ml distilled water and vortexed until completely dissolved. Then 20 ml hexane was added into the prepared solution and vortexed for extra 5 min. This solution was centrifuged at 3500 rpm for 20 min. The supernatant was collected for estimation of orange peel oil nano-
(3)
M = weight, s = sample, t = toluene, p = pycnometer, ρ t = toluene density
ρa =
Mps − Mp Vp
(5)
(4)
2.8.5. Porosity The porosity was calculated using Eq. (5). 371
Carbohydrate Polymers 177 (2017) 369–377
S. Ghasemi et al.
on surface charges, therefore they attract each other and by reaching to the zero charge, they precipitate. Because of this phenomenon, orange peel oil is entrapped into the complex and thus L* index increases. The highest L* value (75.14) was related to treatment No. 6 (WPC = 4%, pectin = 1% and pH = 3), which in this case the prepared complex was stronger and could entrap and encapsulate more orange peel oil, so the sample was brighter. The lowest L* (60) was related to treatment No. 8 (WPC = 4%, pectin = 0.5% and pH = 9) in which, the stability was very high and there was no precipitation. As can be seen in Table 1, maximum (98.5%) and minimum (83%) stability was obtained for treatment No. 10 (WPC = 4%, pectin = 1% and pH = 9) and No. 6 (WPC = 4%, pectin=1% and pH = 3), respectively. Increase in pH values results in higher negative surface charges on the applied protein and then, an increase in the repulsive forces between pectin and WPC. In other words, the zeta potential of the complex increases by negative values and prevents formation of the aggregates and the large complexes between these two biopolymers. As a result, smaller nanocomplexes with a high stability are created. To determine the optimal concentrations for WPC and pectin, and suitable pH values for complex formation, response surface methodology (RSM) was used (Table 1). Our results indicated that 4% WPC and 1% pectin prepared in pH = 3 resulted in the best complex solution in terms of color, viscosity and stability. In fact, a ratio of 4–1 (whey protein to pectin) was the best ratio for complex formation. After determining the optimal biopolymer concentrations, the nanocomplex formation was investigated in three pH values of 3, 6 and 9 for encapsulation of orange peel oil. Firstly, the size and zeta potential of these samples was evaluated and then, they were converted into powders by freeze drying and their qualitative properties including physical, microstructural, and encapsulation efficiency was analyzed which are discussed in the following sections.
encapsulated into biopolymer complexes. The encapsulation efficiency (EE%) of orange peel oil was calculated using Eq. (6) (Jafari, He, & Bhandari, 2007b).
EE % =
(Total amount of loaded oil − surface content of oil) × 100 Total amount of loaded oil (6)
2.12. Statistical analysis Treatments were designed and analyzed by Central Composite Design pattern of Response Surface Methodology through Design Expert Software (version 10). In the 1st phase, effect of three independent variables of WPC content (4, 6, 8% w/w), pectin content (0.5, 0.75, 1% w/w) and pH value (3, 6, 9) on the viscosity, stability and color of liquid samples was evaluated and the optimum samples were selected and freeze-dried. Then, physicochemical and microstructural properties of orange peel oil-loaded powders were determined. 3. Results and discussion 3.1. Physicochemical properties of nano-complex solutions containing orange peel oil Our results revealed that the lowest viscosity value was related to acidic pH values with the exception of treatment No. 6 (WPC = 4%, pectin = 1% and pH = 3) which had the highest viscosity since separation of WPC-pectin complexes was highest in this treatment showing the best attractions between these biopolymers together which could react at the highest electrostatic rate. The Lowest viscosity was related to the treatment No. 16 (WPC = 6%, pectin = 0.75% and pH = 3) because WPC and pectin concentrations and the ratio between these biopolymers was not balanced and their electrostatic interaction was not suitable so the complex didn’t form and by reducing the pH, WPC was precipitated and separated resulting in a decrease in viscosity. For evaluating the effect of concentration and pH on the color or brightness of nano-complex solutions, L* index was reported, because it corresponds well with the transparent appearance of nanocomplex solutions. According to Table 1, samples prepared in pH = 3 had the highest L* since at this pH, pectin is negative and WPC is positive based
3.2. Particle size and zeta potential measurement The particle size and zeta-potential of the optimum WPC-pectin complexes loaded with orange peel oil at three different pH values are summarized in Fig. 1. The particle size for nanocomplexes at pH = 3, 6 and 9 were 360, 182 and 185 nm, respectively; therefore, pH value had a significant influence on the particle size of complexes. Similarly, in a study by Arroyo-Maya and McClement’s (2015), it was shown that the particle size of WPI-pectin complexes varies with pH changes. They encapsulated anthocyanins by formation of electrostatic WPI-Pectin complexes and obtained an average particle size of approximately 200 nm at pH = 4. However, they confirmed that the average particle size was increased at higher or lower pH values. DLS measurements revealed that the smallest complexes between WPC and pectin were formed at pH = 6 (Fig. 1B) which had a high stability to aggregation, similar to the results of Assadpour et al. (2016a) and Lutz et al. (2009a, 2009b). We found that the particle size of orange peel oil-loaded nanocomplexes is increasing at higher pH value of 9, compared to pH = 6. It could be due to the high repulsion between two different applied biopolymers and detachment of pectin biopolymer from the whey protein surfaces, since by increasing the pH to 9, both biopolymers had a net surface negative charge as shown in their z-potential in Fig. 1F. Therefore, they cannot attract each other and by formation of bigger complexes, the particle size is increased. On the other hand, at pH = 3, which is well below the isoelectric point of the whey proteins, both biopolymers have opposite charges (the positive charge on the WPC chain and the negative charge on the pectin chain) which causes formation of strong and comprehensive complex particles with larger sizes (Fig. 1A) and finally, maximum complex coacervation is occurred, resulting in relatively large precipitating aggregates (flocs). Bédié et al. (2008), indicated that the particle size in WPI-pectin complex solutions increased from 6.96 to 21.80 nm by decreasing pH from 4.0 to 2.5 because in pH = 2.5, maximum sedimentable-complexes were formed.
Table 1 RSM treatments and the results of physicochemical properties for prepared WPC-pectin nanocomplexes loaded with orange peel oil.
*
The treatment selected as the suitable one for formation of nanocomplexes for the next stage.
372
Carbohydrate Polymers 177 (2017) 369–377
S. Ghasemi et al.
Fig. 1. The results of DLS (left) and Z-potential (right) analysis for prepared nano-complexes containing orange peel oil at three pH values: (A and D) pH = 3, (B and E) pH = 6, (C and F) pH = 9.
3.3. Influence of pH values on the physicochemical properties of the complex powders
The Z-potential results for complexes formed in pH values of 3, 6 and 9 were +1.7, −19, and −30.3 mV respectively (Fig. 1). The surface charge of complexed biopolymers (zeta potential) determined from electrophoretic mobility measurements was changed from highly negative at pH = 9 (−30.3 mV) to a little positive at pH = 3 (+1.7 mV). At pH = 3 (lower than isoelectric point of whey proteins), two biopolymers neutralize each other which results in a low positive zeta potential approaching zero (+1.7 mV). While, by increasing pH to 6 and 9 (above the isoelectric point), zeta potential of the formed complexes increases by negative values from −19 to −30.3 mV. Similarly, in a study by Lutz et al. (2009a), they indicated that at pH = 2, Z-potential of double emulsions stabilized by WPI-pectin complexes was positive (+18 mV), because the protein was below its isoelectric point (IEP). Near the IEP of the biopolymer mixture (pH = 4.0), the Z-potential was negative, approaching zero (−7 mV). Also, above the IEP (pH = 6), Zpotential was negative (−28 mV).
It was found that pH value is affecting the physicochemical parameters of the freeze-dried powders obtained from complex solutions loaded with orange peel oil. By increasing pH from 3 to 9, water activity and moisture of powder samples was decreased. Moisture content is a crucial powder characteristic, which is related to the stickiness, storage stability, drying efficiency, and powder flowability. Also, high moisture content can cause off-flavors through higher rates of lipid oxidation (Li, Hu, & McClements, 2011). On the other hand, lower moisture content limits the ability of water to act as a plasticizer and reduces the glass transition temperature (Rader, Weaver, & Angyal, 2000). The water activity of all three prepared powders from WPC-pectin complexes was under 0.2 which could be an acceptable level for microbiological stability of the powders. This variation in water activity could be attributed to the strength of the complex between pectin and WPC. Because at pH = 3, formation of stronger complexes results in an increase of water binding within the complex and prevents water movement through capillary tubes during freeze-drying, so the water activity in
Table 2 Physicochemical properties of the WPC-pectin nanocomplex powders containing orange peel oil. Samples
Moisture (%)
Water activity
Solubility (%)
Bulk density (kg/m3)
Absolute density(kg/m3)
Porosity
Surface color (b*)
1 2 3
3.96 3.7 2.47
0.125 0.056 0.041
23 32 88
333 286 222
1197.6 1117.3 1066
0.72 0.74 0.79
40.51 47.86 49.65
373
Carbohydrate Polymers 177 (2017) 369–377
S. Ghasemi et al.
samples with pH = 3 was at higher level. At pH = 9, a weaker complex is formed so the water at capillary tubes removes more easily during drying and the water activity is decreased. The bulk and absolute density of the obtained powders were increased at lower pH values (Table 2). Powders with higher moisture content (sample 1) tend to have a higher weight, because of the existence of water, which is greatly denser than the dry solids (Chegini & Ghobadian, 2005). At pH = 3, a stronger complex is formed between WPC and pectin due to higher attractive forces (electrostatic charges) in this pH situation and therefore, more condensed particles are created resulting in higher absolute and bulk densities in pH = 3. While at pH = 9, because of weaker electrostatic attractive forces between applied biopolymers, hollow particles with lower densities are formed. The porosity of the nanocomplex powders depends on the bulk and absolute density according to Eq. (4), so that by increasing bulk density and decreasing absolute density, the porosity decreases. Therefore, with the increase of pH value from 3 to 9, the bulk density decreased and resulted in a higher porosity. As it is shown in Table 2, by increasing pH, the solubility is increased, which means that the complex powders prepared in pH = 9, can be solved easier and faster than the samples prepared in pH = 3 in water. Since WPC and pectin form stronger bonds with each other in pH = 3 resulting in stronger complexes, probably it is more difficult to break these bonds for dissolving complex powders in water compared with those complex powders prepared at the higher pH values (6 and 9). At pH = 9, a weaker complex is formed with an easier solubility in water. On the other hand, at pH = 3 bigger particles have been formed which need more time to dissolve in water. To assess the effect of pH on the color or brightness of samples, index b* was used because it is correlated well with the effect of pH on powder color. When, orange peel oil is entrapped into the nanocomplex of WPC and pectin, the yellowness (b*) is decreased. According to Table 2, the sample prepared in pH = 3 had the lowest b* value (and the highest encapsulation efficiency) because at this pH, pectin has the most negative charges and WPC has the highest positive charges; so they attracted each other and at the same time, the surface charges of the complexes are neutralizing resulting in precipitation (complex coacervation) of the formed particles containing orange peel oil entrapped within their structure and as a result, b* index is decreased. b* index is indirectly associated with EE% so that an increase in EE% due to entrapment of higher content of orange peel oil through nanocomplexes is correlated with decreasing in b* index (Khazaei, Jafari, Ghorbani, & Kakhki, 2014). This relationship has been described in Section 3.4
Fig. 2. Encapsulation Efficiency (EE%) and b* index of orange peel oil-loaded nanocomplexes of WPC-pectin at three different pH values (A), and the correlation between EE % and b* index (B).
have been entrapped within nanocomplexes; resulted in an increase in the encapsulation efficiency. Conversely, with the increase of the b* index, i.e., the higher yellowness of the powders, it reveals a higher content of orange peel oil has not been encapsulated successfully and it has been deposited on the surface of powder particles, so the encapsulation efficiency is reduced. Therefore, these two responses have an inverse linear correlation together as can be seen in Fig. 2A and B. 3.5. FTIR results FTIR studies have been shown as an accompanying technique to discover any interaction between applied biopolymers. Owing to the interaction of carboxyl groups existing in pectin with amino groups in whey proteins, it could be predicted presence of amide groups in charged complexes of pectin-WPC (Esfanjani, Jafari, Assadpoor, & Mohammadi, 2015). Thus, FTIR analysis was performed to confirm the formation of amide in WPC- pectin complexes loaded with orange peel oil (Fig. 3). The FTIR spectra of pure WPC powders exposed the existence of characteristic functional groups at 1628, 1512, 1421, and 1042 cm−1 corresponding to the stretching of C]O, and the bending of NeH, CeH, and CeN bonds, respectively (Assadpour, Jafari, & Maghsoudlou, 2017; Gujska, Michalak, & Klepacka, 2009). In addition, the FTIR spectra of pure pectin powders showed the presence of a peak at 1023 cm−1 related to the carboxyl group. On the other hand, the spectrum of orange peel oil-loaded WPC-pectin powders demonstrated a sharp band at 991 cm−1, which was related to the C]O stretching; i.e., formation of WPC-pectin complex. Also the peak of 1241 cm−1 in pectin corresponds to sulfate groups that have been moved into 1216 cm−1 in complexes. This cross-linking between two applied biopolymers would led to the formation of a WPC-pectin nanocomplex structure via establishment of net charge between the carboxylate ions of pectin on the amino groups of WPC (Lutz et al., 2009a). Furthermore, all the charged complexes indicated a sharp peak at 1629 cm−1, which could be the amide bond formed between the carboxyl groups of pectin and the amino groups of WPC that shifted from 1628 to 1629 cm−1. In agreement, Assadpour et al. (2017) in their study on spray dried powder particles of pectin-whey protein nano-capsules also achieved similar results and showed a peak at 1629 cm−1, which was related to
3.4. Encapsulation efficiency of WPC-pectin nanocomplexes A weight ratio of 1:4 for pectin:WPC resulted in the best nanocomplex formation and a high encapsulation efficiency similar to the results of Assadpour et al. (2016a). We compared the encapsulation efficiency (EE%) of nanocomplexes prepared by this ratio at three pH values (3, 6 and 9). The amount of orange peel oil on the surface of the nano-complex powders was determined as the non-encapsulated fraction using UV–vis spectrophotometry at 231 nm. The EE% was then calculated using Eq. (1). In addition, it was revealed that EE% of prepared nano-complexes in pH values of 3, 6, 9 was 88%, 84% and 70%, respectively. Maximum EE value was obtained in pH = 3 as the complex coacervate was formed with higher and stronger entrapment of orange peel oil within nanocomplex particles. It was found that by increasing the pH from 6 to 9, EE% of orange peel oil within prepared nano complexes was decreased which can be attributed to weaker complex formation at higher pH values. EE% had a very well correlation with b* index. As it is shown in Fig. 2A, by reduction in the yellowness of the powders, b* index is decreased which means the amount of orange peel oil on the surface of powder particles (non-encapsulated fraction) is reduced and higher bioactive ingredients (orange peel oil) 374
Carbohydrate Polymers 177 (2017) 369–377
S. Ghasemi et al.
Fig. 3. FTIR spectra of WPC-pectin nanocomplex powders in three pH values: red line: (pH = 6), green line: pH = 9, brown line: pH = 3. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
Fig. 4. SEM images of powders formulated with WPC-Pectin nanocomplexes loaded with orange peel oil and prepared using an initial weight ratio of WPC to Pectin of 4:1 at pH = 3.
which is an effective technique to provide the surface morphology and size distribution of nano scale samples. AFM images confirmed the spheroid like appearance and nano size structure of the WPC-pectin complexes (Fig. 5). It was revealed that most of WPC-pectin nanocomplexes loaded with orange peel oil were distributed between the 50 and 200 nm in terms of particle size. However, the size of the WPCpectin nanoparticles in pH = 6 was smaller compared to the nanoparticles in others pH values. It is noticeable that AFM imaging was performed on dehydrated samples, and therefore, the size of particles was not comparable to DLS measurements. In addition, these images (Fig. 5) provide the surface morphology with a more accurate size and size distribution. AFM images also confirmed the spherical morpholgy and nanosize structure of WPC-pectin complexes. Comparison between AFM images of the prepared samples showed that the nano-complex particles prepared in pH = 3 (Fig. 5, A-1 and A-2) appeared to be somewhat larger but more spherical than the particles formulated in pH = 6 (Fig. 5, B-1 and B-2) and pH = 9 (Fig. 5, C-1 and C-2). It may be a result of complex coacervation and aggregation of the WPC-pectin complex particles in pH = 3. The smallest particles were observed at pH = 6. In samples prepared at pH = 9, both biopolymers had a net surface negative charge so, the repulsion between two different applied biopolymers is increased and detachment of pectin biopolymer from the whey protein surfaces is occurring which results in bigger particle sizes as it is
the amide bond formed between the amino groups of WPC and carboxyl groups of pectin. What initially seen was the simultaneous presence of peaks related to the WPC and pectin in the final prepared nanocomplexes, which had been moved somehow. One of the main peaks was related to hydrogen bonding that occurred in the range of 3000 to 3500 cm−1. As can be seen in Fig. 3, this peak was observed in WPC and pectin at 3323 and 3421 cm−1 respectively which was shifted to another position in 3388 cm−1 in the complexes. It was sharper in complexes at 3300 cm−1 and associated with hydrogen bonds. These shifts clearly indicated the occurrence of interaction between the biopolymers. Luo et al. (2011) investigated the electrostatic interaction between alpha-tocopherol, zein and chitosan carriers and indicated that the 1664 and 1550 cm−1peaks (in the zein) were shifted to the 1657 and 1542 cm−1 that was related to the electrostatic bonding between the applied ingredients. 3.6. Shape and morphology of nano-complexes The morphology of orange peel oil-loaded nanocomplex powders was observed by SEM (Fig. 4). A regular distribution and representative sample of freeze-dried powders was observed. Also, the morphological characteristics of the particles formed by WPC-pectin complex at three pH values were determined using AFM, 375
Carbohydrate Polymers 177 (2017) 369–377
S. Ghasemi et al.
Fig. 5. AFM images (left) and size distribution (right) of nanocomplexes of WPC-pectin in three pH values: A: pH = 3 B: pH = 6, C: pH = 9.
but at pH = 9, the repulsion between WPC and pectin was increased and weak complexes were formed. Furthermore, the smallest particles with high stability were formed in pH = 6. The encapsulation efficiency of the powders at pH = 3, 6 and 9 were 88, 84, and 70%, respectively and a well reverse linear correlation was obtained between encapsulation efficiency and the color index (b*). The morphology of orange peel oil-loaded nanocomplexes was analyzed by AFM, SEM and confirmed the spherical shape of complexes especially at pH = 3.
obvious in Fig. 5C. 4. Conclusion This study showed that complex biopolymer nanoparticles can be fabricated by electrostatic complexation of whey protein–pectin mixtures. It was found that formation of nanocomplexes between WPC and pectin as wall materials was capable of encapsulating high amounts of orange peel oil. The pH value was a significant factor influencing formation of the nanocomplexs in aqueous solution. Our results showed that strongest and weakest nanocomplexes containing orange peel oil were formed in pH = 3 and pH = 9, respectively; because at pH = 3, WPC and pectin had opposite charges and attracted each other strongly
Acknowledgement It is necessary to appreciate Iran Nanotechnology Initiative Council (INIC) for the financial support. 376
Carbohydrate Polymers 177 (2017) 369–377
S. Ghasemi et al.
characterization of zein/chitosan complex for encapsulation of α-tocopherol, and its in vitro controlled release study. Colloids and Surfaces B: Biointerfaces, 85, 145–152. Lutz, R., Aserin, A., Wicker, L., & Garti, N. (2009a). Double emulsions stabilized by a charged complex of modified pectin and whey protein isolate. Colloids and Surfaces B: Biointerfaces, 72, 121–127. Lutz, R., Aserin, A., Wicker, L., & Garti, N. (2009b). Release of electrolytes from W/O/W double emulsions stabilized by a soluble complex of modified pectin and whey protein isolate. Colloids and Surfaces B: Biointerfaces, 74, 178–185. Madene, A., Jacquot, M., Scher, J., & Desobry, S. (2006). Flavour encapsulation and controlled release – A review. International Journal of Food Science & Technology, 41, 1–21. Matalanis, A., Jones, O. G., & McClements, D. J. (2011). Structured biopolymer-based delivery systems for encapsulation, protection, and release of lipophilic compounds. Food Hydrocolloids, 25, 1865–1880. Matias, R., Ribeiro, P., Sarraguça, M., & Lopes, J. (2014). A UV spectrophotometric method for the determination of folic acid in pharmaceutical tablets and dissolution tests. Analytical Methods, 6, 3065–3071. Mendanha, D. V., Ortiz, S. E. M., Favaro-Trindade, C. S., Mauri, A., Monterrey-Quintero, E. S., & Thomazini, M. (2009). Microencapsulation of casein hydrolysate by complex coacervation with SPI/pectin. Food Research International, 42, 1099–1104. Mohammadi, A., Jafari, S. M., Assadpour, E., & Esfanjani, A. F. (2016). Nano-encapsulation of olive leaf phenolic compounds through WPC–pectin complexes and evaluating their release rate. International Journal of Biological Macromolecules, 82, 816–822. Peinado, I., Lesmes, U., Andrés, A., & McClements, J. D. (2010). Fabrication and morphological characterization of biopolymer particles formed by electrostatic complexation of heat treated lactoferrin and anionic polysaccharides. Langmuir, 26, 9827–9834. Rader, J. I., Weaver, C. M., & Angyal, G. (2000). Total folate in enriched cereal-grain products in the United States following fortification. Food Chemistry, 70, 275–289. Tan Khang, W., Tang Siah, Y., Thomas, R., Vasanthakumari, N., & Manickam, S. (2016). Curcumin-loaded sterically stabilized nanodispersion based on non-ionic colloidal system induced by ultrasound and solvent diffusion-evaporation. Pure and Applied Chemistry, 88, 37–43. Tang, S. Y., Shridharan, P., & Sivakumar, M. (2013a). Impact of process parameters in the generation of novel aspirin nanoemulsions – Comparative studies between ultrasound cavitation and microfluidizer. Ultrasonics Sonochemistry, 20(1), 485–497. Tang, S. Y., Sivakumar, M., & Nashiru, B. (2013b). Impact of osmotic pressure and gelling in the generation of highly stable single core water-in-oil-in-water (W/O/W) nano multiple emulsions of aspirin assisted by two-stage ultrasonic cavitational emulsification. Colloids and Surfaces B: Biointerfaces, 102, 653–658. Teng, Z., Luo, Y., & Wang, Q. (2013). Carboxymethyl chitosan–soy protein complex nanoparticles for the encapsulation and controlled release of vitamin D3. Food Chemistry, 141, 524–532. Wagoner, T. B., & Foegeding, E. A. (2017). Whey protein–pectin soluble complexes for beverage applications. Food Hydrocolloids, 63, 130–138. Weinbreck, F., Nieuwenhuijse, H., Robijn, G. W., & de Kruif, C. G. (2004). Complexation of whey proteins with carrageenan. Journal of Agricultural and Food Chemistry, 52, 3550–3555. Ye, A. (2008). Complexation between milk proteins and polysaccharides via electrostatic interaction: Principles and applications – A review. International Journal of Food Science & Technology, 43, 406–415. Yeo, Y., Bellas, E., Firestone, W., Langer, R., & Kohane, D. S. (2005). Complex coacervates for thermally sensitive controlled release of flavor compounds. Journal of Agricultural and Food Chemistry, 53, 7518–7525. Zimet, P., & Livney, Y. D. (2009). Beta-lactoglobulin and its nanocomplexes with pectin as vehicles for ω-3 polyunsaturated fatty acids. Food Hydrocolloids, 23, 1120–1126.
References Anal, A. K., Tobiassen, A., Flanagan, J., & Singh, H. (2008). Preparation and characterization of nanoparticles formed by chitosan–caseinate interactions. Colloids and Surfaces B: Biointerfaces, 64, 104–110. Arroyo-Maya, I. J., & McClements, D. J. (2015). Biopolymer nanoparticles as potential delivery systems for anthocyanins: Fabrication and properties. Food Research International, 69, 1–8. Assadpour, E., & Jafari, S.-M. (2017). Spray drying of folic acid within nano-emulsions: optimization by taguchi approach. Drying Technology, 35(9), 1152–1160. Assadpour, E., Maghsoudlou, Y., Jafari, S.-M., Ghorbani, M., & Aalami, M. (2016a). Evaluation of folic acid nano-encapsulation by double emulsions. Food and Bioprocess Technology, 9, 2024–2032. Assadpour, E., Maghsoudlou, Y., Jafari, S.-M., Ghorbani, M., & Aalami, M. (2016b). Optimization of folic acid nano-emulsification and encapsulation by maltodextrinwhey protein double emulsions. International Journal of Biological Macromolecules, 86, 197–207. Assadpour, E., Jafari, S.-M., & Maghsoudlou, Y. (2017). Evaluation of folic acid release from spray dried powder particles of pectin-whey protein nano-capsules. International Journal of Biological Macromolecules, 95, 238–247. Bédié, G. K., Turgeon, S. L., & Makhlouf, J. (2008). Formation of native whey protein isolate–low methoxyl pectin complexes as a matrix for hydro-soluble food ingredient entrapment in acidic foods. Food Hydrocolloids, 22, 836–844. Bosnea, L. A., Moschakis, T., & Biliaderis, C. G. (2014). Complex coacervation as a novel microencapsulation technique to improve viability of probiotics under different stresses. Food and Bioprocess Technology, 7, 2767–2781. Chegini, G., & Ghobadian, B. (2005). Effect of spray-drying conditions on physical properties of orange juice powder. Drying Technology, 23, 657–668. Esfanjani, A. F., Jafari, S. M., Assadpoor, E., & Mohammadi, A. (2015). Nano-encapsulation of saffron extract through double-layered multiple emulsions of pectin and whey protein concentrate. Journal of Food Engineering, 165, 149–155. Gujska, E., Michalak, J., & Klepacka, J. (2009). Folates stability in two types of rye breads during processing and frozen storage. Plant Foods for Human Nutrition, 64, 129–134. Hosseini, S. F., Zandi, M., Rezaei, M., & Farahmandghavi, F. (2013). Two-step method for encapsulation of oregano essential oil in chitosan nanoparticles: preparation, characterization and in vitro release study. Carbohydrate Polymers, 95, 50–56. Hosseini, S. M. H., Emam-Djomeh, Z., Sabatino, P., & Van der Meeren, P. (2015). Nanocomplexes arising from protein-polysaccharide electrostatic interaction as a promising carrier for nutraceutical compounds. Food Hydrocolloids, 50, 16–26. Jafari, S. M., He, Y., & Bhandari, B. (2007a). Encapsulation of nanoparticles of d-limonene by spray drying: Role of emulsifiers and emulsifying techniques. Drying Technology, 25, 1069–1079. Jafari, S. M., He, Y., & Bhandari, B. (2007b). Production of sub-micron emulsions by ultrasound and microfluidization techniques. Journal of Food Engineering, 82, 478–488. Jun-xia, X., Hai-yan, Y., & Jian, Y. (2011). Microencapsulation of sweet orange oil by complex coacervation with soybean protein isolate/gum Arabic. Food Chemistry, 125, 1267–1272. Khazaei, K. M., Jafari, S., Ghorbani, M., & Kakhki, A. H. (2014). Application of maltodextrin and gum Arabic in microencapsulation of saffron petal's anthocyanins and evaluating their storage stability and color. Carbohydrate Polymers, 105, 57–62. Li, Y., Hu, M., & McClements, D. J. (2011). Factors affecting lipase digestibility of emulsified lipids using an in vitro digestion model: Proposal for a standardised pHstat method. Food Chemistry, 126, 498–505. Livney, Y. D. (2010). Milk proteins as vehicles for bioactives. Current Opinion in Colloid & Interface Science, 15, 73–83. Luo, Y., Zhang, B., Whent, M., Yu, L. L., & Wang, Q. (2011). Preparation and
377
Contents lists available at ScienceDirect
Carbohydrate Polymers journal homepage: www.elsevier.com/locate/carbpol
Production of pectin-whey protein nano-complexes as carriers of orange peel oil
T
⁎
Sanaz Ghasemi, Seid Mahdi Jafari , Elham Assadpour, Morteza Khomeiri Faculty of Food Science and Technology, Gorgan University of Agricultural Sciences and Natural Resources, Gorgan, Iran
A R T I C L E I N F O
A B S T R A C T
Keywords: Nanocomplex Orange peel oil Encapsulation Whey protein Pectin
Orange peel oil is one of the most common flavorings used in the food industry which is volatile under environmental conditions. Encapsulation is the best way to protect it and control its release. One of the nanoencapsulation systems for food bioactive ingredients is complexation method, which entraps the core materials in a complex of two different biopolymers. In this study, orange peel oil was nanoencapsulated by pectinwhey protein nanocomplexes. After determining the optimum nanocomplex suspensions containing orange peel oil based on the stability, viscosity, and color, they were formulated in three different pH values (3, 6 and 9) and converted into powdered forms by freeze drying. The analysis of size and zeta potential of nanocomplexes revealed that the smallest particles formed in pH = 6. The encapsulation efficiency of the powders at pH = 3, 6 and 9 were 88, 84, and 70%, respectively and there was a reverse linear correlation between encapsulation efficiency and the color index (b*). The microstructure and the morphology of the nanocomplex powders was investigated by SEM and AFM and the results showed that more spherical particles are formed in pH = 3. FTIR analysis determined that there was a chemical reaction and bond formation between whey proteins and pectin as a sharp band was appeared in 991 cm−1.
1. Introduction Flavor is known as the essence of foods and plays an important role in consumer satisfaction and consumption of foods. Most liquid food flavorings are volatile substances and chemically unstable when exposed to the air, light, moisture and high temperatures. Orange peel oil is one of the most widely used flavorings in the food industry. It is a mixture of various chemical components and is mainly composed of Dlimonene (> 90%). D-Limonene is the base sensory character of the citrus oils and due to its instability under the process and environmental conditions, it is necessary to encapsulate it prior to use in foods or beverages to limit aroma degradation or losses during processing and storage (Jun-xia, Hai-yan, & Jian, 2011; Madene, Jacquot, Scher, & Desobry, 2006). Encapsulation is a quickly expanding technology which has been generally used for many food applications such as taste and odor covering in the form of micro- and nanoparticles. This technology can control the release of bioactive ingredients under specific conditions, so causing a long shelf-life. One of the nanoencapsulation systems for protection of flavors is complexation method which entraps the flavor oils in a complex of two different biopolymers. In general, these biopolymers are proteins and polysaccharides (Yeo, Bellas, Firestone,
⁎
Corresponding author. E-mail address: [email protected] (S.M. Jafari).
http://dx.doi.org/10.1016/j.carbpol.2017.09.009 Received 4 July 2017; Received in revised form 16 August 2017; Accepted 5 September 2017 Available online 06 September 2017 0144-8617/ © 2017 Elsevier Ltd. All rights reserved.
Langer, & Kohane, 2005) which can join together through electrostatic attractions. Polysaccharide-protein nanoparticles are taken more into consideration than pure single biopolymer nanoparticles, because of their higher chemical and colloidal protection. Proteins often have a negative charge in the pH values above their isoelectric points (pI ≈ 5) and positive charge below this pH value. So, when proteins and polysaccharides are mixed together in a liquid medium, two phenomena can occur: 1) Higher rate of the attractive interactions resulting in formation of soluble and insoluble complexes; 2) Higher rate of the repulsive interactions causing separation of the two biopolymers from each other (Weinbreck, Nieuwenhuijse, Robijn, & de Kruif, 2004). In practice, one of these two phenomena will occur depending on the charge of both biopolymers and effective factors such as pH and ionic strength. The absorption interactions, further occurs through the creation of electrostatic bonds between oppositely charged biopolymers such as proteins with the positive charges (in pH < pI) and anionic polysaccharides containing carboxyl groups, phosphates and sulfates (at a pH > pKa), or negatively charged proteins (pH > pI) and cationic polysaccharides such as chitosan (Matalanis, Jones, & McClements, 2011). This complex formation is guided by biopolymer parameters including charge density of the biopolymer, its molecular weight, chemical nature; environmental conditions including ionic strength, ion
Carbohydrate Polymers 177 (2017) 369–377
S. Ghasemi et al.
2. Materials and methods
type, biopolymer concentration, biopolymer ratio, pH; and processing factors, including temperature, pressure, shearing rate and time which may affect the formation and stability of the complexes (Ye, 2008). The complexation of proteins and polysaccharides such as β-lactoglobulin and pectin can be applied to encapsulate hydrophilic and hydrophobic nutraceuticals including different vitamins, flavors, essential oils, fatty acids, etc. (Zimet & Livney, 2009). When formulating these biopolymer complexes by high energy densities, it results in nano sized complexes which are optically transparent with higher surface-to-volume ratio and could protect the encapsulated nutraceuticals from chemical degradation more efficiently. Ultrasonication is one of these high energy devices which can result in nano-sized droplets and particles through cavitation and it has been successfully applied in preparation of nanoemulsions and nanodispersions loaded with different bioactive ingredients and drugs (Tang, Shridharan, & Sivakumar, 2013; Tang, Sivakumar, & Nashiru, 2013; Tan Khang, Tang Siah, Thomas, Vasanthakumari, & Manickam, 2016). Milk proteins, mostly whey proteins, have been extensively used in the studies of protein– polysaccharide complex systems (Livney, 2010). On the other hand, pectin is an anionic hetero polysaccharide associated with the cell wall in plants and its main chain has been formed by galacturonic acid units with α-D connector (1–4) and esterified partially with methanol. Commercial low methoxyl pectin (LMP) is an anionic polysaccharide obtained from de-esterification of high methoxyl pectin (Mendanha et al., 2009). Bédié et al. (2008) investigated the influence of protein to pectin ratios, pH and pre or post-blending acidification on the formation of thiamin-loaded complexes. They blended LMP and whey protein isolate (WPI) solutions to achieve protein to pectin ratios of 2:1, 1:1 and 1:2 at a total biopolymer concentration of 0.4% (w/w) with pH adjustment in the range of 2.5–4.0 either before or after blending. Their results showed that at a ratio of 2:1 (protein: pectin), the size of complex was constant and lower at post-blending acidification compared to preblending. But at a ratio of 2:1 (WPI:pectin), the optimum pH for maximum turbidity and complex yield was around 3.0 with both pre- and post-blending acidifications. Anal et al. (2008) studied inter-macromolecular complexation between sodium caseinate and chitosan in aqueous solutions as a function of pH (3.0–6.5). The chitosan–caseinate complexes formed in the pH range 4.8–6.0 (with opposite charges) were stable and soluble. The soluble complexes associated to form larger particles at higher concentrations of chitosan (0.15 wt%). It was revealed that the particles formed by the chitosan- caseinate complexes at pH 4.8–6.0, had sizes between 250 and 350 nm. At pH values above 6.0, the nano-complex particles attached together and formed larger particles, due to phase separation. In the last couple of years, some other studies have been conducted about production and application of nano complexes of different proteins with polysaccharides such as WPI – beet pectin (Arroyo-Maya & McClements, 2015), carboxy methyl chitosan – soy protein (Teng, Luo, & Wang, 2013), WPC – pectin (Mohammadi, Jafari, Assadpour, & Esfanjani, 2016), WPI – gum Arabic (Bosnea, Moschakis, & Biliaderis, 2014) and WPI – high-methoxyl pectin (Wagoner & Foegeding, 2017). Our main goal in this study was producing orange peel oil-loaded nanocapsules based on complexed biopolymers of whey protein concentrate and low methoxyl pectin to encapsulate this popular flavor and introduce a new formulation of orange peel oil for applications in the food industry, particularly in beverages. We decided to answer this question whether is it possible to produce pectin-WPC nanocomplex carriers for successful loading of orange peel oil or not? Also, we worked on the influence of various factors including the concentration of pectin and whey protein concentrate and pH values on the complex formation, in order to find the optimal conditions for producing best nanocomplexes. The final goal was to evaluate the microstructure and physicochemical properties of freeze-dried powder particles containing orange peel oil-loaded nanocomplexes.
2.1. Materials The applied biopolymers were an aqueous solution of whey protein concentrate (WPC; 80 w/w%) in a combination with maltodextrin (MD; DE 16–20) and pectin (LMP). Whey protein concentrate and maltodextrin were obtained from Arla (Denmark) and Qinhuangdao Starch Co. (China), respectively. Low methoxyl pectin from citrus peel (galacturonic acid, ≥74.0% (dried basis), DE = 25%) was purchased from Sigma Chemicals Company (USA). Orange peel oil was purchased from Ramsar Citrus Concentrate Co. (Iran). A non-ionic surfactant, Tween 80 (Sigma) was used as the emulsifying agent during emulsion preparation stages of this work. Deionized water was used to prepare all the solutions. 2.2. Biopolymer solution preparation Firstly, aqueous solutions of WPC were prepared by dispersing 4, 6 and 8 g of WPC powder into deionized water to obtain 100 ml solutions. Also, aqueous solutions of pectin were prepared by dispersing 0.5, 0.75, and 1 g LMP powder into hot deionized water (70 °C) to obtain 100 ml solutions. Then, 50% w/w maltodextrin solutions were prepared as a filler to increase the total solids for preparation of powders. Solutions were stirred for 30 min on a magnetic stirrer (IKA, Germany) then they were stored overnight at 4 °C for complete hydration of biopolymers (Assadpour, Maghsoudlou, Jafari, Ghorbani, & Aalami, 2016a, 2016b). 2.3. Preparation of WPC-Pectin nano-complexes containing orange peel oil WPC and pectin solutions were mixed together along with maltodextrin at constant ratio of 50%w/w with the same volume of each solution for 30 min on a magnetic stirrer. Then, Tween 80 at a ratio of 10% of the total solids was added into the solution and mixed again to dissolve completely. Finally, orange peel oil was added into these prepared solutions gradually during homogenization by an ultrasonic probe (Iranian Ultrasonic Technology Company, 400 W, 20kHZ, 12 mm probe diameter) at 25 °C and power of 350 W for 10 min. pH of the solutions was adjusted to 3, 6, 9 using HCl and NaOH (1 M) (Jafari, He, & Bhandari, 2007a; Lutz, Aserin, Wicker, & Garti, 2009). 2.4. Physicochemical properties of nano-complex solutions After preparing the complexes, the effect of composition and preparation conditions of biopolymer solutions on viscosity, color (L*) and stability were investigated. Viscosity was measured by using a Brookfield viscometer (LVDV Pro II, Brook-field Engineering Laboratories, spindle S00, USA), L* was measured by image analysis using Image J software and also for measuring the stability of the nanocomplexes, accelerated method by a centrifuge (Sigma Co., 3K30 model) in 20,000 g and 25 °C for 30 min was applied (Hosseini, EmamDjomeh, Sabatino, & Van der Meeren, 2015; Zimet & Livney, 2009). 2.5. Particle size and zeta potential measurement Particle size and zeta potential (surface electrical charge) of prepared nano-complexes were measured using a dynamic light scattering (DLS) method (Zeta Sizer, Malvern Instruments, Malvern, UK). To avoid multiple scattering, the samples were diluted with distilled water and at a temperature of 25 °C, the average particle size (based on volume, number, and intensity mean diameter), and zeta potential were determined. 2.6. Stability of nano-complex solutions For performing the stability test, 50 ml of nanocomplex solutions 370
Carbohydrate Polymers 177 (2017) 369–377
S. Ghasemi et al.
(primary nanocomplex volume) was centrifuged and then, the volume of remained stable nanocomplex was determined by a graduated cylinder. Then, the percentage of stability was calculated with the following equation.
Stability (%) =
The volume of nanocomplex remained stable × 100 The volume of primary nanocomplex
ρ ε = 1 − ⎜⎛ b ⎞⎟ ⎝ ρa ⎠ ρa = absolute density, ρb = bulk density.
2.8.6. Solubility For measuring the solubility, 1 g of powder was added to 100 ml of distilled water under stirring with a magnetic stirrer at 385 rpm for 5 min. The resulting dispersion was centrifuged at 3000g for 5 min. 25 ml of the upper solution was transferred to a Petri dish that has already been weighed and dried in an oven at temperature of 105° C for 5 h. The percentage of solubility was determined based on the amount of powder dissolved in water.
(1)
2.7. Freeze-drying of the orange peel oil-loaded nano-complexes The optimum prepared nano-complexes were frozen in flasks at −20 °C for 48 h. Frozen samples were dried using a freeze dryer (FDB5503 Freeze-dryer; Operon, South Korea). The freeze-dried samples were stored in hermetic containers at 4 °C. For further experiments, the dried nano-complexes were grinded into a suitable powder by a pestle and mortar.
2.8.7. Surface color analysis The surface color of powders, which is relevant to the surface component content (orange peel oil), was evaluated by the procedure of Khazaei et al. (2014) with some modifications. The yellow (b*) index at the powder surface was measured because of yellow–orange color of orange peel oil. For all samples, color was measured using image analysis software (Image j). For evaluating the surface color of the powders, 10 g of the powders was transferred into a Petri dish and was placed at the scanner (HP, Scanjet G3110, UK) and images were taken. Subsequently, powders color was analyzed by using Image J software and b* factor of the samples was measured.
2.8. Physicochemical properties of the prepared powders containing nanocomplexes The physicochemical properties of powders including solubility, water activity, moisture content, bulk density, absolute density, porosity, and surface color were measured. 2.8.1. Moisture About 2 g of the powder was placed in a Petri dish for 2–3 h in an oven (UFB 400, Memert, Germany) at temperature of 105 °C, then cooled in a desiccator, and weighed. The moisture content was calculated using Eq. (2).
M% =
W2 − W3 × 100 W2 − W1
2.9. FTIR analysis The structure analysis of the WPC–pectin nanocomplex samples were investigated by Fourier Transform Infrared Spectroscopy (FTIR). In addition, the IR spectra of the samples were registered by FTIR spectrophotometer (Spectrum RX1, PerkinElmer, US) using attenuated total reflection technique. For structure analysis, at first the powder samples were mixed with KBr powder in 1–10 ratios and pressed into a disk before spectrum acquisition. The spectrum was scanned in transmission mode from 400 to 4500−1 cm range.
(2)
M = moisture (%), W1 = weight of petri dish, W2 = the total weight of petri dish and powder, W3 = the total weight of dried powder and petri dish after processing (oven). 2.8.2. Water activity (aw) For determination of water activity, we applied instrumental analysis by Novasina LabMaster Std Labmaster Standard Water Activity Instrument (UK).
2.10. Morphological characterization of nano-complexes The morphology of the powders loaded with orange peel oil was observed by scanning electron microscopy (SEM) (Vega II, Tescan, Czech Republic). The powder was spread on a clean glass plate and then coated with a thin layer of gold metal under high vacuum and observed. Representative SEM images were described (Hosseini, Zandi, Rezaei, & Farahmandghavi, 2013). Atomic force microscope (AFM) (DualScopeTM DS95-50, DME, Denmark) was used for morphological characterization and particle size and size distribution of WPC-pectin nano-complexes. Samples for AFM imaging were prepared by applying a drop of diluted nano-complexes suspension (0.05 mg/mL) on a clean glass surface, spread and dried at room temperature for 30 min. Then the image analysis by AFM was performed (Peinado, Lesmes, Andrés, & McClements, 2010).
2.8.3. Bulk density The bulk density of the powders was measured by weighing 2 g of powder and placing it in a 25 ml graduated cylinder. Then the cylinder was tapped 100 times by hand and the bulk density was calculated from the ratio between the mass of powder (2 g) and the volume occupied in the cylinder (Assadpour and Jafari, 2017). 2.8.4. Absolute density To calculate absolute density of powders, we applied displacement method through pycnometer. Toluene volume (Vp) was obtained Using Eq. (3) and the absolute density was calculated using Eq. (4) (Chegini & Ghobadian, 2005).
Vp =
2.11. Encapsulation efficiency of orange peel oil
(Mpt − Mp) − (Mpst − Mps ) ρt
It was necessary to analyze encapsulated nanocomplex powders in terms of total content and surface content of orange peel oil in final powders. For surface (unencapsulated) content, 0.5 g of each sample was dispersed in 20 ml hexane, vortexed for 2 min and filtered using Whatman paper no. 41 and the orange peel oil content was determined spectrophotometrically at 231 nm (Matias, Ribeiro, Sarraguça, & Lopes, 2014). For total content, 0.5 g of powders was dissolved in 20 ml distilled water and vortexed until completely dissolved. Then 20 ml hexane was added into the prepared solution and vortexed for extra 5 min. This solution was centrifuged at 3500 rpm for 20 min. The supernatant was collected for estimation of orange peel oil nano-
(3)
M = weight, s = sample, t = toluene, p = pycnometer, ρ t = toluene density
ρa =
Mps − Mp Vp
(5)
(4)
2.8.5. Porosity The porosity was calculated using Eq. (5). 371
Carbohydrate Polymers 177 (2017) 369–377
S. Ghasemi et al.
on surface charges, therefore they attract each other and by reaching to the zero charge, they precipitate. Because of this phenomenon, orange peel oil is entrapped into the complex and thus L* index increases. The highest L* value (75.14) was related to treatment No. 6 (WPC = 4%, pectin = 1% and pH = 3), which in this case the prepared complex was stronger and could entrap and encapsulate more orange peel oil, so the sample was brighter. The lowest L* (60) was related to treatment No. 8 (WPC = 4%, pectin = 0.5% and pH = 9) in which, the stability was very high and there was no precipitation. As can be seen in Table 1, maximum (98.5%) and minimum (83%) stability was obtained for treatment No. 10 (WPC = 4%, pectin = 1% and pH = 9) and No. 6 (WPC = 4%, pectin=1% and pH = 3), respectively. Increase in pH values results in higher negative surface charges on the applied protein and then, an increase in the repulsive forces between pectin and WPC. In other words, the zeta potential of the complex increases by negative values and prevents formation of the aggregates and the large complexes between these two biopolymers. As a result, smaller nanocomplexes with a high stability are created. To determine the optimal concentrations for WPC and pectin, and suitable pH values for complex formation, response surface methodology (RSM) was used (Table 1). Our results indicated that 4% WPC and 1% pectin prepared in pH = 3 resulted in the best complex solution in terms of color, viscosity and stability. In fact, a ratio of 4–1 (whey protein to pectin) was the best ratio for complex formation. After determining the optimal biopolymer concentrations, the nanocomplex formation was investigated in three pH values of 3, 6 and 9 for encapsulation of orange peel oil. Firstly, the size and zeta potential of these samples was evaluated and then, they were converted into powders by freeze drying and their qualitative properties including physical, microstructural, and encapsulation efficiency was analyzed which are discussed in the following sections.
encapsulated into biopolymer complexes. The encapsulation efficiency (EE%) of orange peel oil was calculated using Eq. (6) (Jafari, He, & Bhandari, 2007b).
EE % =
(Total amount of loaded oil − surface content of oil) × 100 Total amount of loaded oil (6)
2.12. Statistical analysis Treatments were designed and analyzed by Central Composite Design pattern of Response Surface Methodology through Design Expert Software (version 10). In the 1st phase, effect of three independent variables of WPC content (4, 6, 8% w/w), pectin content (0.5, 0.75, 1% w/w) and pH value (3, 6, 9) on the viscosity, stability and color of liquid samples was evaluated and the optimum samples were selected and freeze-dried. Then, physicochemical and microstructural properties of orange peel oil-loaded powders were determined. 3. Results and discussion 3.1. Physicochemical properties of nano-complex solutions containing orange peel oil Our results revealed that the lowest viscosity value was related to acidic pH values with the exception of treatment No. 6 (WPC = 4%, pectin = 1% and pH = 3) which had the highest viscosity since separation of WPC-pectin complexes was highest in this treatment showing the best attractions between these biopolymers together which could react at the highest electrostatic rate. The Lowest viscosity was related to the treatment No. 16 (WPC = 6%, pectin = 0.75% and pH = 3) because WPC and pectin concentrations and the ratio between these biopolymers was not balanced and their electrostatic interaction was not suitable so the complex didn’t form and by reducing the pH, WPC was precipitated and separated resulting in a decrease in viscosity. For evaluating the effect of concentration and pH on the color or brightness of nano-complex solutions, L* index was reported, because it corresponds well with the transparent appearance of nanocomplex solutions. According to Table 1, samples prepared in pH = 3 had the highest L* since at this pH, pectin is negative and WPC is positive based
3.2. Particle size and zeta potential measurement The particle size and zeta-potential of the optimum WPC-pectin complexes loaded with orange peel oil at three different pH values are summarized in Fig. 1. The particle size for nanocomplexes at pH = 3, 6 and 9 were 360, 182 and 185 nm, respectively; therefore, pH value had a significant influence on the particle size of complexes. Similarly, in a study by Arroyo-Maya and McClement’s (2015), it was shown that the particle size of WPI-pectin complexes varies with pH changes. They encapsulated anthocyanins by formation of electrostatic WPI-Pectin complexes and obtained an average particle size of approximately 200 nm at pH = 4. However, they confirmed that the average particle size was increased at higher or lower pH values. DLS measurements revealed that the smallest complexes between WPC and pectin were formed at pH = 6 (Fig. 1B) which had a high stability to aggregation, similar to the results of Assadpour et al. (2016a) and Lutz et al. (2009a, 2009b). We found that the particle size of orange peel oil-loaded nanocomplexes is increasing at higher pH value of 9, compared to pH = 6. It could be due to the high repulsion between two different applied biopolymers and detachment of pectin biopolymer from the whey protein surfaces, since by increasing the pH to 9, both biopolymers had a net surface negative charge as shown in their z-potential in Fig. 1F. Therefore, they cannot attract each other and by formation of bigger complexes, the particle size is increased. On the other hand, at pH = 3, which is well below the isoelectric point of the whey proteins, both biopolymers have opposite charges (the positive charge on the WPC chain and the negative charge on the pectin chain) which causes formation of strong and comprehensive complex particles with larger sizes (Fig. 1A) and finally, maximum complex coacervation is occurred, resulting in relatively large precipitating aggregates (flocs). Bédié et al. (2008), indicated that the particle size in WPI-pectin complex solutions increased from 6.96 to 21.80 nm by decreasing pH from 4.0 to 2.5 because in pH = 2.5, maximum sedimentable-complexes were formed.
Table 1 RSM treatments and the results of physicochemical properties for prepared WPC-pectin nanocomplexes loaded with orange peel oil.
*
The treatment selected as the suitable one for formation of nanocomplexes for the next stage.
372
Carbohydrate Polymers 177 (2017) 369–377
S. Ghasemi et al.
Fig. 1. The results of DLS (left) and Z-potential (right) analysis for prepared nano-complexes containing orange peel oil at three pH values: (A and D) pH = 3, (B and E) pH = 6, (C and F) pH = 9.
3.3. Influence of pH values on the physicochemical properties of the complex powders
The Z-potential results for complexes formed in pH values of 3, 6 and 9 were +1.7, −19, and −30.3 mV respectively (Fig. 1). The surface charge of complexed biopolymers (zeta potential) determined from electrophoretic mobility measurements was changed from highly negative at pH = 9 (−30.3 mV) to a little positive at pH = 3 (+1.7 mV). At pH = 3 (lower than isoelectric point of whey proteins), two biopolymers neutralize each other which results in a low positive zeta potential approaching zero (+1.7 mV). While, by increasing pH to 6 and 9 (above the isoelectric point), zeta potential of the formed complexes increases by negative values from −19 to −30.3 mV. Similarly, in a study by Lutz et al. (2009a), they indicated that at pH = 2, Z-potential of double emulsions stabilized by WPI-pectin complexes was positive (+18 mV), because the protein was below its isoelectric point (IEP). Near the IEP of the biopolymer mixture (pH = 4.0), the Z-potential was negative, approaching zero (−7 mV). Also, above the IEP (pH = 6), Zpotential was negative (−28 mV).
It was found that pH value is affecting the physicochemical parameters of the freeze-dried powders obtained from complex solutions loaded with orange peel oil. By increasing pH from 3 to 9, water activity and moisture of powder samples was decreased. Moisture content is a crucial powder characteristic, which is related to the stickiness, storage stability, drying efficiency, and powder flowability. Also, high moisture content can cause off-flavors through higher rates of lipid oxidation (Li, Hu, & McClements, 2011). On the other hand, lower moisture content limits the ability of water to act as a plasticizer and reduces the glass transition temperature (Rader, Weaver, & Angyal, 2000). The water activity of all three prepared powders from WPC-pectin complexes was under 0.2 which could be an acceptable level for microbiological stability of the powders. This variation in water activity could be attributed to the strength of the complex between pectin and WPC. Because at pH = 3, formation of stronger complexes results in an increase of water binding within the complex and prevents water movement through capillary tubes during freeze-drying, so the water activity in
Table 2 Physicochemical properties of the WPC-pectin nanocomplex powders containing orange peel oil. Samples
Moisture (%)
Water activity
Solubility (%)
Bulk density (kg/m3)
Absolute density(kg/m3)
Porosity
Surface color (b*)
1 2 3
3.96 3.7 2.47
0.125 0.056 0.041
23 32 88
333 286 222
1197.6 1117.3 1066
0.72 0.74 0.79
40.51 47.86 49.65
373
Carbohydrate Polymers 177 (2017) 369–377
S. Ghasemi et al.
samples with pH = 3 was at higher level. At pH = 9, a weaker complex is formed so the water at capillary tubes removes more easily during drying and the water activity is decreased. The bulk and absolute density of the obtained powders were increased at lower pH values (Table 2). Powders with higher moisture content (sample 1) tend to have a higher weight, because of the existence of water, which is greatly denser than the dry solids (Chegini & Ghobadian, 2005). At pH = 3, a stronger complex is formed between WPC and pectin due to higher attractive forces (electrostatic charges) in this pH situation and therefore, more condensed particles are created resulting in higher absolute and bulk densities in pH = 3. While at pH = 9, because of weaker electrostatic attractive forces between applied biopolymers, hollow particles with lower densities are formed. The porosity of the nanocomplex powders depends on the bulk and absolute density according to Eq. (4), so that by increasing bulk density and decreasing absolute density, the porosity decreases. Therefore, with the increase of pH value from 3 to 9, the bulk density decreased and resulted in a higher porosity. As it is shown in Table 2, by increasing pH, the solubility is increased, which means that the complex powders prepared in pH = 9, can be solved easier and faster than the samples prepared in pH = 3 in water. Since WPC and pectin form stronger bonds with each other in pH = 3 resulting in stronger complexes, probably it is more difficult to break these bonds for dissolving complex powders in water compared with those complex powders prepared at the higher pH values (6 and 9). At pH = 9, a weaker complex is formed with an easier solubility in water. On the other hand, at pH = 3 bigger particles have been formed which need more time to dissolve in water. To assess the effect of pH on the color or brightness of samples, index b* was used because it is correlated well with the effect of pH on powder color. When, orange peel oil is entrapped into the nanocomplex of WPC and pectin, the yellowness (b*) is decreased. According to Table 2, the sample prepared in pH = 3 had the lowest b* value (and the highest encapsulation efficiency) because at this pH, pectin has the most negative charges and WPC has the highest positive charges; so they attracted each other and at the same time, the surface charges of the complexes are neutralizing resulting in precipitation (complex coacervation) of the formed particles containing orange peel oil entrapped within their structure and as a result, b* index is decreased. b* index is indirectly associated with EE% so that an increase in EE% due to entrapment of higher content of orange peel oil through nanocomplexes is correlated with decreasing in b* index (Khazaei, Jafari, Ghorbani, & Kakhki, 2014). This relationship has been described in Section 3.4
Fig. 2. Encapsulation Efficiency (EE%) and b* index of orange peel oil-loaded nanocomplexes of WPC-pectin at three different pH values (A), and the correlation between EE % and b* index (B).
have been entrapped within nanocomplexes; resulted in an increase in the encapsulation efficiency. Conversely, with the increase of the b* index, i.e., the higher yellowness of the powders, it reveals a higher content of orange peel oil has not been encapsulated successfully and it has been deposited on the surface of powder particles, so the encapsulation efficiency is reduced. Therefore, these two responses have an inverse linear correlation together as can be seen in Fig. 2A and B. 3.5. FTIR results FTIR studies have been shown as an accompanying technique to discover any interaction between applied biopolymers. Owing to the interaction of carboxyl groups existing in pectin with amino groups in whey proteins, it could be predicted presence of amide groups in charged complexes of pectin-WPC (Esfanjani, Jafari, Assadpoor, & Mohammadi, 2015). Thus, FTIR analysis was performed to confirm the formation of amide in WPC- pectin complexes loaded with orange peel oil (Fig. 3). The FTIR spectra of pure WPC powders exposed the existence of characteristic functional groups at 1628, 1512, 1421, and 1042 cm−1 corresponding to the stretching of C]O, and the bending of NeH, CeH, and CeN bonds, respectively (Assadpour, Jafari, & Maghsoudlou, 2017; Gujska, Michalak, & Klepacka, 2009). In addition, the FTIR spectra of pure pectin powders showed the presence of a peak at 1023 cm−1 related to the carboxyl group. On the other hand, the spectrum of orange peel oil-loaded WPC-pectin powders demonstrated a sharp band at 991 cm−1, which was related to the C]O stretching; i.e., formation of WPC-pectin complex. Also the peak of 1241 cm−1 in pectin corresponds to sulfate groups that have been moved into 1216 cm−1 in complexes. This cross-linking between two applied biopolymers would led to the formation of a WPC-pectin nanocomplex structure via establishment of net charge between the carboxylate ions of pectin on the amino groups of WPC (Lutz et al., 2009a). Furthermore, all the charged complexes indicated a sharp peak at 1629 cm−1, which could be the amide bond formed between the carboxyl groups of pectin and the amino groups of WPC that shifted from 1628 to 1629 cm−1. In agreement, Assadpour et al. (2017) in their study on spray dried powder particles of pectin-whey protein nano-capsules also achieved similar results and showed a peak at 1629 cm−1, which was related to
3.4. Encapsulation efficiency of WPC-pectin nanocomplexes A weight ratio of 1:4 for pectin:WPC resulted in the best nanocomplex formation and a high encapsulation efficiency similar to the results of Assadpour et al. (2016a). We compared the encapsulation efficiency (EE%) of nanocomplexes prepared by this ratio at three pH values (3, 6 and 9). The amount of orange peel oil on the surface of the nano-complex powders was determined as the non-encapsulated fraction using UV–vis spectrophotometry at 231 nm. The EE% was then calculated using Eq. (1). In addition, it was revealed that EE% of prepared nano-complexes in pH values of 3, 6, 9 was 88%, 84% and 70%, respectively. Maximum EE value was obtained in pH = 3 as the complex coacervate was formed with higher and stronger entrapment of orange peel oil within nanocomplex particles. It was found that by increasing the pH from 6 to 9, EE% of orange peel oil within prepared nano complexes was decreased which can be attributed to weaker complex formation at higher pH values. EE% had a very well correlation with b* index. As it is shown in Fig. 2A, by reduction in the yellowness of the powders, b* index is decreased which means the amount of orange peel oil on the surface of powder particles (non-encapsulated fraction) is reduced and higher bioactive ingredients (orange peel oil) 374
Carbohydrate Polymers 177 (2017) 369–377
S. Ghasemi et al.
Fig. 3. FTIR spectra of WPC-pectin nanocomplex powders in three pH values: red line: (pH = 6), green line: pH = 9, brown line: pH = 3. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
Fig. 4. SEM images of powders formulated with WPC-Pectin nanocomplexes loaded with orange peel oil and prepared using an initial weight ratio of WPC to Pectin of 4:1 at pH = 3.
which is an effective technique to provide the surface morphology and size distribution of nano scale samples. AFM images confirmed the spheroid like appearance and nano size structure of the WPC-pectin complexes (Fig. 5). It was revealed that most of WPC-pectin nanocomplexes loaded with orange peel oil were distributed between the 50 and 200 nm in terms of particle size. However, the size of the WPCpectin nanoparticles in pH = 6 was smaller compared to the nanoparticles in others pH values. It is noticeable that AFM imaging was performed on dehydrated samples, and therefore, the size of particles was not comparable to DLS measurements. In addition, these images (Fig. 5) provide the surface morphology with a more accurate size and size distribution. AFM images also confirmed the spherical morpholgy and nanosize structure of WPC-pectin complexes. Comparison between AFM images of the prepared samples showed that the nano-complex particles prepared in pH = 3 (Fig. 5, A-1 and A-2) appeared to be somewhat larger but more spherical than the particles formulated in pH = 6 (Fig. 5, B-1 and B-2) and pH = 9 (Fig. 5, C-1 and C-2). It may be a result of complex coacervation and aggregation of the WPC-pectin complex particles in pH = 3. The smallest particles were observed at pH = 6. In samples prepared at pH = 9, both biopolymers had a net surface negative charge so, the repulsion between two different applied biopolymers is increased and detachment of pectin biopolymer from the whey protein surfaces is occurring which results in bigger particle sizes as it is
the amide bond formed between the amino groups of WPC and carboxyl groups of pectin. What initially seen was the simultaneous presence of peaks related to the WPC and pectin in the final prepared nanocomplexes, which had been moved somehow. One of the main peaks was related to hydrogen bonding that occurred in the range of 3000 to 3500 cm−1. As can be seen in Fig. 3, this peak was observed in WPC and pectin at 3323 and 3421 cm−1 respectively which was shifted to another position in 3388 cm−1 in the complexes. It was sharper in complexes at 3300 cm−1 and associated with hydrogen bonds. These shifts clearly indicated the occurrence of interaction between the biopolymers. Luo et al. (2011) investigated the electrostatic interaction between alpha-tocopherol, zein and chitosan carriers and indicated that the 1664 and 1550 cm−1peaks (in the zein) were shifted to the 1657 and 1542 cm−1 that was related to the electrostatic bonding between the applied ingredients. 3.6. Shape and morphology of nano-complexes The morphology of orange peel oil-loaded nanocomplex powders was observed by SEM (Fig. 4). A regular distribution and representative sample of freeze-dried powders was observed. Also, the morphological characteristics of the particles formed by WPC-pectin complex at three pH values were determined using AFM, 375
Carbohydrate Polymers 177 (2017) 369–377
S. Ghasemi et al.
Fig. 5. AFM images (left) and size distribution (right) of nanocomplexes of WPC-pectin in three pH values: A: pH = 3 B: pH = 6, C: pH = 9.
but at pH = 9, the repulsion between WPC and pectin was increased and weak complexes were formed. Furthermore, the smallest particles with high stability were formed in pH = 6. The encapsulation efficiency of the powders at pH = 3, 6 and 9 were 88, 84, and 70%, respectively and a well reverse linear correlation was obtained between encapsulation efficiency and the color index (b*). The morphology of orange peel oil-loaded nanocomplexes was analyzed by AFM, SEM and confirmed the spherical shape of complexes especially at pH = 3.
obvious in Fig. 5C. 4. Conclusion This study showed that complex biopolymer nanoparticles can be fabricated by electrostatic complexation of whey protein–pectin mixtures. It was found that formation of nanocomplexes between WPC and pectin as wall materials was capable of encapsulating high amounts of orange peel oil. The pH value was a significant factor influencing formation of the nanocomplexs in aqueous solution. Our results showed that strongest and weakest nanocomplexes containing orange peel oil were formed in pH = 3 and pH = 9, respectively; because at pH = 3, WPC and pectin had opposite charges and attracted each other strongly
Acknowledgement It is necessary to appreciate Iran Nanotechnology Initiative Council (INIC) for the financial support. 376
Carbohydrate Polymers 177 (2017) 369–377
S. Ghasemi et al.
characterization of zein/chitosan complex for encapsulation of α-tocopherol, and its in vitro controlled release study. Colloids and Surfaces B: Biointerfaces, 85, 145–152. Lutz, R., Aserin, A., Wicker, L., & Garti, N. (2009a). Double emulsions stabilized by a charged complex of modified pectin and whey protein isolate. Colloids and Surfaces B: Biointerfaces, 72, 121–127. Lutz, R., Aserin, A., Wicker, L., & Garti, N. (2009b). Release of electrolytes from W/O/W double emulsions stabilized by a soluble complex of modified pectin and whey protein isolate. Colloids and Surfaces B: Biointerfaces, 74, 178–185. Madene, A., Jacquot, M., Scher, J., & Desobry, S. (2006). Flavour encapsulation and controlled release – A review. International Journal of Food Science & Technology, 41, 1–21. Matalanis, A., Jones, O. G., & McClements, D. J. (2011). Structured biopolymer-based delivery systems for encapsulation, protection, and release of lipophilic compounds. Food Hydrocolloids, 25, 1865–1880. Matias, R., Ribeiro, P., Sarraguça, M., & Lopes, J. (2014). A UV spectrophotometric method for the determination of folic acid in pharmaceutical tablets and dissolution tests. Analytical Methods, 6, 3065–3071. Mendanha, D. V., Ortiz, S. E. M., Favaro-Trindade, C. S., Mauri, A., Monterrey-Quintero, E. S., & Thomazini, M. (2009). Microencapsulation of casein hydrolysate by complex coacervation with SPI/pectin. Food Research International, 42, 1099–1104. Mohammadi, A., Jafari, S. M., Assadpour, E., & Esfanjani, A. F. (2016). Nano-encapsulation of olive leaf phenolic compounds through WPC–pectin complexes and evaluating their release rate. International Journal of Biological Macromolecules, 82, 816–822. Peinado, I., Lesmes, U., Andrés, A., & McClements, J. D. (2010). Fabrication and morphological characterization of biopolymer particles formed by electrostatic complexation of heat treated lactoferrin and anionic polysaccharides. Langmuir, 26, 9827–9834. Rader, J. I., Weaver, C. M., & Angyal, G. (2000). Total folate in enriched cereal-grain products in the United States following fortification. Food Chemistry, 70, 275–289. Tan Khang, W., Tang Siah, Y., Thomas, R., Vasanthakumari, N., & Manickam, S. (2016). Curcumin-loaded sterically stabilized nanodispersion based on non-ionic colloidal system induced by ultrasound and solvent diffusion-evaporation. Pure and Applied Chemistry, 88, 37–43. Tang, S. Y., Shridharan, P., & Sivakumar, M. (2013a). Impact of process parameters in the generation of novel aspirin nanoemulsions – Comparative studies between ultrasound cavitation and microfluidizer. Ultrasonics Sonochemistry, 20(1), 485–497. Tang, S. Y., Sivakumar, M., & Nashiru, B. (2013b). Impact of osmotic pressure and gelling in the generation of highly stable single core water-in-oil-in-water (W/O/W) nano multiple emulsions of aspirin assisted by two-stage ultrasonic cavitational emulsification. Colloids and Surfaces B: Biointerfaces, 102, 653–658. Teng, Z., Luo, Y., & Wang, Q. (2013). Carboxymethyl chitosan–soy protein complex nanoparticles for the encapsulation and controlled release of vitamin D3. Food Chemistry, 141, 524–532. Wagoner, T. B., & Foegeding, E. A. (2017). Whey protein–pectin soluble complexes for beverage applications. Food Hydrocolloids, 63, 130–138. Weinbreck, F., Nieuwenhuijse, H., Robijn, G. W., & de Kruif, C. G. (2004). Complexation of whey proteins with carrageenan. Journal of Agricultural and Food Chemistry, 52, 3550–3555. Ye, A. (2008). Complexation between milk proteins and polysaccharides via electrostatic interaction: Principles and applications – A review. International Journal of Food Science & Technology, 43, 406–415. Yeo, Y., Bellas, E., Firestone, W., Langer, R., & Kohane, D. S. (2005). Complex coacervates for thermally sensitive controlled release of flavor compounds. Journal of Agricultural and Food Chemistry, 53, 7518–7525. Zimet, P., & Livney, Y. D. (2009). Beta-lactoglobulin and its nanocomplexes with pectin as vehicles for ω-3 polyunsaturated fatty acids. Food Hydrocolloids, 23, 1120–1126.
References Anal, A. K., Tobiassen, A., Flanagan, J., & Singh, H. (2008). Preparation and characterization of nanoparticles formed by chitosan–caseinate interactions. Colloids and Surfaces B: Biointerfaces, 64, 104–110. Arroyo-Maya, I. J., & McClements, D. J. (2015). Biopolymer nanoparticles as potential delivery systems for anthocyanins: Fabrication and properties. Food Research International, 69, 1–8. Assadpour, E., & Jafari, S.-M. (2017). Spray drying of folic acid within nano-emulsions: optimization by taguchi approach. Drying Technology, 35(9), 1152–1160. Assadpour, E., Maghsoudlou, Y., Jafari, S.-M., Ghorbani, M., & Aalami, M. (2016a). Evaluation of folic acid nano-encapsulation by double emulsions. Food and Bioprocess Technology, 9, 2024–2032. Assadpour, E., Maghsoudlou, Y., Jafari, S.-M., Ghorbani, M., & Aalami, M. (2016b). Optimization of folic acid nano-emulsification and encapsulation by maltodextrinwhey protein double emulsions. International Journal of Biological Macromolecules, 86, 197–207. Assadpour, E., Jafari, S.-M., & Maghsoudlou, Y. (2017). Evaluation of folic acid release from spray dried powder particles of pectin-whey protein nano-capsules. International Journal of Biological Macromolecules, 95, 238–247. Bédié, G. K., Turgeon, S. L., & Makhlouf, J. (2008). Formation of native whey protein isolate–low methoxyl pectin complexes as a matrix for hydro-soluble food ingredient entrapment in acidic foods. Food Hydrocolloids, 22, 836–844. Bosnea, L. A., Moschakis, T., & Biliaderis, C. G. (2014). Complex coacervation as a novel microencapsulation technique to improve viability of probiotics under different stresses. Food and Bioprocess Technology, 7, 2767–2781. Chegini, G., & Ghobadian, B. (2005). Effect of spray-drying conditions on physical properties of orange juice powder. Drying Technology, 23, 657–668. Esfanjani, A. F., Jafari, S. M., Assadpoor, E., & Mohammadi, A. (2015). Nano-encapsulation of saffron extract through double-layered multiple emulsions of pectin and whey protein concentrate. Journal of Food Engineering, 165, 149–155. Gujska, E., Michalak, J., & Klepacka, J. (2009). Folates stability in two types of rye breads during processing and frozen storage. Plant Foods for Human Nutrition, 64, 129–134. Hosseini, S. F., Zandi, M., Rezaei, M., & Farahmandghavi, F. (2013). Two-step method for encapsulation of oregano essential oil in chitosan nanoparticles: preparation, characterization and in vitro release study. Carbohydrate Polymers, 95, 50–56. Hosseini, S. M. H., Emam-Djomeh, Z., Sabatino, P., & Van der Meeren, P. (2015). Nanocomplexes arising from protein-polysaccharide electrostatic interaction as a promising carrier for nutraceutical compounds. Food Hydrocolloids, 50, 16–26. Jafari, S. M., He, Y., & Bhandari, B. (2007a). Encapsulation of nanoparticles of d-limonene by spray drying: Role of emulsifiers and emulsifying techniques. Drying Technology, 25, 1069–1079. Jafari, S. M., He, Y., & Bhandari, B. (2007b). Production of sub-micron emulsions by ultrasound and microfluidization techniques. Journal of Food Engineering, 82, 478–488. Jun-xia, X., Hai-yan, Y., & Jian, Y. (2011). Microencapsulation of sweet orange oil by complex coacervation with soybean protein isolate/gum Arabic. Food Chemistry, 125, 1267–1272. Khazaei, K. M., Jafari, S., Ghorbani, M., & Kakhki, A. H. (2014). Application of maltodextrin and gum Arabic in microencapsulation of saffron petal's anthocyanins and evaluating their storage stability and color. Carbohydrate Polymers, 105, 57–62. Li, Y., Hu, M., & McClements, D. J. (2011). Factors affecting lipase digestibility of emulsified lipids using an in vitro digestion model: Proposal for a standardised pHstat method. Food Chemistry, 126, 498–505. Livney, Y. D. (2010). Milk proteins as vehicles for bioactives. Current Opinion in Colloid & Interface Science, 15, 73–83. Luo, Y., Zhang, B., Whent, M., Yu, L. L., & Wang, Q. (2011). Preparation and
377