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Formation of concentrated particles composed of oppositely charged biopolymers for food applications – impact of processing conditions
Food Structure 10 (2016) 10–20
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Research paper
Formation of concentrated particles composed of oppositely charged biopolymers for food applications – impact of processing conditions Benjamin Zeeba , Catrin Stengera , Jörg Hinrichsb , Jochen Weissa,* a Department of Food Physics and Meat Science, Institute of Food Science and Biotechnology, University of Hohenheim, Garbenstrasse 21/25, 70599 Stuttgart, Germany b Department of Soft Matter Science and Dairy Technology, Institute of Food Science and Biotechnology, University of Hohenheim, Garbenstrasse 21, 70599 Stuttgart, Germany
A R T I C L E I N F O
Article history: Received 30 May 2016 Received in revised form 13 October 2016 Accepted 19 October 2016 Available online 21 October 2016 Keywords: Whey protein isolate Degree of esterification Pectin Complex coacervation Concentrated complexes Biopolymer particles
A B S T R A C T
Associative complexation approaches are promising tools to generate mixed biopolymer particles with tailor-made physicochemical properties. However, association of oppositely charged biopolymers typically occurs under acidic conditions and low biopolymer concentrations. The present study aims to investigate intrinsic and extrinsic parameters to generate concentrated biopolymer dispersions which could be utilized as structuring agents in food matrices. We therefore utilized a multistep procedure. Firstly, solutions of whey protein isolate (WPI) and pectins with different degrees of esterification (DE) were mixed under neutral conditions at various ratios (WPI:Pectin 1:2, 1:1, 2:1, 5:1, 8:1). Secondly, the pH was adjusted to 3 to promote complex coacervation. Thirdly, complex dispersions were subsequently heat-treated to tailor their overall water content, whereas temperatures below (q = 50–55 C) and above (q = 90–95 C) the protein’s denaturation were applied. Phase separation behavior, microstructural, rheological and electrical properties of the complexes were investigated by surface charge, turbidity, particle size, rheometry and light microscopy measurements. Results revealed that complexes composed of WPI and pectin with a low DE formed rather small and dense particles, whereas large aggregates were observed when the pectins DE was increased. Concentrated complex dispersions with water contents 80% could be manufactured regardless of heating temperatures, whereas complexes maintained their particulate structures upon re-dispersion. Results are of importance for future studies where we intend to incorporate concentrated biopolymer particles as fat replacer in meat and dairy matrices. ã 2016 Elsevier Ltd. All rights reserved.
1. Introduction Numerous studies focus on fabrication methods to design novel food structures possessing specific functionalities in terms of texture, appearance, mouthfeel, and flavor (Scholten, Moschakis, & Biliaderis, 2014). In particular, self-assembly techniques gained enormous interest along food manufacturers and technologists allowing ones to precisely control the interaction forces between molecules and colloids on a molecular, nano, colloidal, and microscale level (Zeeb, Fischer, & Weiss, 2014). In general, structural design principles based on self-assembly are rather cost effective, feasible, and ecofriendly, but require fundamental knowledge of the relationship between different hierarchical levels. In particular, directed self-assembly approaches such as
* Corresponding author. E-mail address: [email protected] (J. Weiss). http://dx.doi.org/10.1016/j.foostr.2016.10.002 2213-3291/ã 2016 Elsevier Ltd. All rights reserved.
associative coacervation has been widely reported in literature (Dickinson, 1998; de Kruif & Tuinier, 2001; Jones & McClements, 2010b, 2011; Schmitt, Sanchez, Desobry-Banon, & Hardy, 1998). Although biopolymer complexes generated by associative phase separation are known to have promising functional properties, their use in real food matrices as texture modifier or fat replacer is rather limited. One major drawback of complexes used for food applications is their low concentration range to be applied to prepare stable biopolymer particles without gelation (Turgeon, Laneuville, Stefan, Ian, & Johan, 2009). In general, associative complexation of mixed protein-polysaccharide systems occurs under conditions were charged polymers electrostatically attract each other, whereas the physicochemical properties of the complexes generated can be tailored by both, intrinsic (e.g. biopolymer type, biopolymer flexibility, branching, protein:pectin ratio) and extrinsic factors (e.g. pH, ionic strength, temperature, shear stress, overall biopolymer concentration) (Azarikia, Wu, Abbasi, & McClements, 2015; Jones, Decker, &
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McClements, 2009b; Jones, Lesmes, Dubin, & McClements, 2010b; Jones & McClements, 2010a; Krzeminski, Prell, Weiss, & Hinrichs, 2014; Samant, Singhal, Kulkarni, & Rege, 1993; Schmitt et al., 1998). In a previous study conducted in our lab, we could demonstrate that high WPI:Citrus pectin ratios led to positive surface charges promoting the formation of large and dense particles that easily precipitated. In addition, complexes composed of WPI and highmethoxylated pectins had a relatively small and monomodal particle size distribution (Salminen & Weiss, 2013). Moreover, it was shown that the local overall charge density of pectins can be controlled by their degree of methyl-esterification – a key parameter to modulate the electrostatic attraction between pectins and their oppositely charged counterpart (Sperber, Schols, Cohen Stuart, Norde, & Voragen, 2009). As such, low-methylated pectins (DE < 50 %) are known to have a high overall charge, whereas high-methylated pectins (DE > 50 %) are pectins with a low overall charge (Akhtar, Dickinson, Mazoyer, & Langendorff, 2002; May, 1990). In the current study, whey protein isolate and pectins with different degrees of esterification were utilized since they are commercially available as texture modifier, thickener, emulsifier, or gelling agents in many food products (Akhtar et al., 2002; Bryant & McClements, 1998; Leroux, Langendorff, Schick, Vaishnav, & Mazoyer, 2003). Krzeminski et al. (2014) demonstrated that WPI:Pectin complexes could be embedded into a skim milk yoghurt as fat replacer having a similar texture profile than their full-fat control. In addition, we recently conducted a study to generate concentrated biopolymer dispersions by a simple heat treatment (q = 90–95 C) to simulate textural and sensorial properties of low-fat products. It has been shown that concentrated complex dispersions with water contents 80 % could be manufactured and easily re-dispersed, whereas the complexes maintained their particulate matter (Stenger, Zeeb, Hinrichs, & Weiss, 2016). The objective of the present study, however, was to gain further insights in intrinsic and extrinsic factors affecting the formation of heat-concentrated biopolymer particles. We hypothesized that heating conditions above the protein’s denaturation temperature might yield in biopolymer complexes which are stable against redispersion, whereas incorporated pectins having a low DE may form small and dense structures due to strong electrostatic attraction. Therefore, whey protein isolate (WPI) and pectin (DE 5.4%, 55%, 71%) solutions were mixed at pH 7 at various biopolymer ratios and the pH was subsequently lowered to acidic conditions to promote complexation. Biopolymer dispersions were then heattreated using two different temperature ranges, namely q = 50– 55 C and q = 90–95 C, to tailor the overall water content. The results might have important implications for the preparation of biopolymer particles that could be utilized as fat replacers in dairy or meat products. 2. Materials and methods 2.1. Materials Whey protein isolate (WPI 895) was obtained from Fonterra (Auckland, New Zealand). As stated by the manufacturer, the composition of the WPI used was 93.9 % protein (69.2 % b-lactoglobulin, 14.2% a-lactalbumin, 3.3% bovine serum albumin, 2.1% immunoglobulin G, 1.6% glycomacropeptide, 1.2% proteose peptone 5), 4.7% moisture, 0.3 % fat, 0.4 % carbohydrates, and 1.5 % minerals. Pectin samples with various degrees of esterification (DE) were donated by Herbstreith & Fox KG (Neuenbürg, Germany), namely citrus pectin (DE 71%, P71, Pectin Classic CU 201, 84% galacturonic acid), sugar beet pectin (DE 55%, P55, Betapec RU 301, 65% galacturonic acid), and citrus pectin (DE 5.4%, P5.4, Pectin Classic CU 902, 85% galacturonic acid). The pectin samples were
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used without further purification. Analytical grade sodium hydroxide (NaOH) and hydrochloric acid (HCl) were obtained from Carl Roth GmbH + Co. KG (Karlsruhe, Germany). All samples were prepared in double-distilled water. All concentrations are expressed in mass percentage (%w/w). 2.2. Fabrication of biopolymer complexes Biopolymer complexes were generated as previously described (Stenger et al., 2016). In brief, powdered WPI (max. 20%) and pectin (max. 4.5%) were dissolved in double-distilled water and stirred at room temperature overnight. After complete hydration, the biopolymer solutions were readjusted to pH 7.0 using 0.1 M and 1 M HCl and/or NaOH, respectively. Both, WPI and pectin stock solutions were then mixed at various WPI:Pectin ratios (1:2, 1:1, 2:1, 5:1, 8:1) and the pH was adjusted to pH 7, 5, and 3, respectively. The final biopolymer concentration ranged between cbiopolymer = 0.6 –6.0%, whereas single WPI and pectin solutions were used as control samples. 2.3. Particle size determination Particle size measurements were performed using a static light scattering instrument (Horiba LA-950, Retsch Technology GmbH, Haan, Germany). Samples from the biopolymer dispersions were withdrawn and diluted to a concentration of approximately 0.005% with an appropriate buffer to prevent multiple scattering effects. The measurement principle is based on the angular dependence of the intensity of the scattered light from a laser beam by the dilute colloidal dispersion. The Mie theory was used to calculate the droplet size distributions. A refractive index of 1.42 for the WPI: Pectin complexes was used as previously determined by Krzeminski et al. (2014). However, it has to be noted that particle size measurements have to be treated with some caution since the biopolymer complexes generated are not entirely spherical in shape. 2.4. Turbidity measurements A photometer (Agilent 8453 UV–vis Spectroscopy System, Agilent Technologies Inc., CA, USA) was used to determine the phase separation behavior of mixed biopolymer systems by measuring the turbidity at 600 nm. High turbidity values of samples indicate milky dispersions and were thus samples containing particles which were large enough to scatter light. Double distilled water was used as reference. 2.5. z-Potential measurements A particle electrophoresis instrument (Nano ZS, Malvern Instruments, Malvern, UK) was used to determine the z-potential of single biopolymer and complex dispersions. Samples were loaded into an appropriate cuvette and the z-potential was determined by measuring the direction and velocity that the droplets moved in the electric field applied. The Smoluchowski equation was utilized to calculate the z-potential. The z-potential measurements were made from two freshly prepared samples, and were carried out with four readings per sample. All samples were made in duplicate. 2.6. Rheological analysis Rheological analysis was conducted using a rheometer MCR300 (Anton Paar GmbH, Graz, Austria; Software: Rheoplus/32 V3.40). The measurement system used was a plate-to-plate geometry (d = 25 mm, PP25, Part No. 79044, Anton Paar). An amplitude
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sweep was conducted with 40 measuring points in the strain range between 0.1 and 100 % (frequency f = 1 Hz; measuring gap = 1 mm; q = 25 C). The measuring point duration was set to 10 s. At the beginning of each measurement, a relaxation time of 30 s was applied. Each sample was measured in duplicate, whereas the storage modulus G', loss modulus G”, and damping factor tan d were utilized as a key parameter to determine the rheological properties of heat-concentrated dispersions. However, storage moduli <0.1 Pa should be treated with some caution due to the detection limit of the instrument. 2.7. Optical microscopy All samples were gently mixed prior to analysis to ensure homogeneity. 10 mL of each sample were placed on an objective plate and a cover glass was carefully slid on top. Light microscopy images were taken with an axial mounted Canon Powershot G10 digital camera (Canon, Tokyo, Japan) mounted on an Axio Scope optical microscope (A1, Carl Zeiss Microimaging GmbH, Göttingen, Germany). 2.8. Heat-induced concentration of biopolymer complexes Complex dispersions (WPI:Pectin ratio 1:2, 2:1, 8:1, cbiopolymer = 6.0%) formed at pH 3 were subjected to a heat-induced concentration step, whereas temperatures below and above the proteins denaturation were applied: q = 50–55 C (Modell400, Memmert GmbH & Co.KG, Schwabach, Germany) and q = 90–95 C (Professional Hot Plate Stirrer, VWR, Radnor, PA, USA) (Dickinson, 2010). Dispersions were heated at a heating rate of 0.5 C/min and low shear rates (<100 s1). At regular time intervals, aliquots were withdrawn to gravimetrically determine the water content according to Eqs. (1) and (2). mi;biopolymer ð1Þ 100% W i ð%Þ ¼ 1 mi;total with mi;biopolymer ¼
1
W i1 mi1;total 100%
ð2Þ
where Wi is the water content of sample i at a given time. Phase separation behavior and rheological properties of heat-treated samples were determined. In addition, single WPI and pectin solutions served as control samples.
2.9. Reconstitution of heat-treated complexes Concentrated complex dispersions (WPI:Pectin ratio 1:2, 2:1, 8:1) by various temperature ranges (q = 50–55 C vs. q = 90–95 C) were redispersed to their initial water content (94.0%) using double-distilled water which was adjusted to pH 3. Particle size distributions and microstructure were then assessed after 12 h of reconstitution. 2.10. Statistical analysis All samples were prepared in duplicate. The averages and the standard deviations were calculated from all measurements of the duplicate samples using Excel Software (Microsoft Corporation, Redmond, WA, USA). 3. Results and discussion 3.1. Characterization of base biopolymer solutions Initially, we examined the influence of pH (7–3) on the electrical properties of the biopolymer solutions since electrostatic attraction is the major driving force for the association of WPI: Pectin complexes (Azarikia et al., 2015). In particular, the affect of the pectins DE (5.4%, 55%, 71%) on the z-potential and turbidity of biopolymer dispersions were identified when the pH was moved from neutral to acidic (Table 1). The electrical charge of WPI changed from 29.3 2.0 mV to +27.9 2.5 mV as the pH was decreased from 7 to 3 which could be attributed to the fact that the pH was altered from above to below the isoelectric point (pI) of the protein (Azarikia et al., 2015). In earlier published literature the pI of WPI was already identified between pH 5 and 4 (Salminen & Weiss, 2014; Zeeb, Salminen, Fischer, & Weiss, 2014). In addition, turbidity measurements revealed high WPI solubility above and below the proteins’ pI – a fact that could be attributed to strong electrostatic repulsion between protein molecules (Majhi et al., 2006; Verheul, Roefs, & De Kruif, 1998). All pectin samples used remained negatively charged when the pH was decreased from neutral to acidic conditions regardless of DE, whereas a slight increase in the z-potential was measured between pH 4.5 and 3.0 – a fact that can be attributed to the pKa values of the pectins studied (Neirynck et al., 2007). However, our results revealed that the DE had a clear impact on the electrical properties of the pectin dispersions. As such, low-methoxylated
Table 1 Surface charge properties (z-potential) and turbidity of single biopolymer solutions (cbiopolymer = 0.6%) as a function of pH (3–7): Whey protein isolate (WPI) and pectin with DE of 5.4%, 55%, and 71%, respectively. pH
WPI
P5.4
P55
P71
z-Pot. (mV)
T600nm (%)
z-Pot. (mV)
T600nm (%)
z-Pot. (mV)
T600nm (%)
z-Pot. (mV)
T600nm (%)
3.0
27.9 2.5
29.7 0.4
29.5 1.0
14.4 0.8
90.6 0.4
14.6 2.1
31.3 0.5
3.5
23.9 1.9
35.3 0.5
28.6 1.0
22.6 0.9
90.2 1.0
24.1 2.0
30.8 0.5
4.0 4.5 5.0 5.5
15.6 1.0 3.4 0.8 10.8 0.8 17.5 0.7
43.3 1.7 46.7 2.9 47.3 0.3 47.8 0.7
27.4 0.8 26.3 0.5 25.0 0.8 23.5 1.4
29.0 0.8 32.7 1.3 34.2 1.0 34.4 1.2
90.1 0.7 89.8 0.5 89.8 0.8 88.6 0.7
30.6 1.7 35.8 0.9 37.0 1.7 38.4 1.3
29.7 1.0 28.2 0.4 26.5 0.7 25.1 0.9
6.0
22.6 1.5
48.0 0.7
22.9 1.2
34.1 0.9
88.8 1.2
37.6 1.3
22.4 1.1
6.5
26.2 1.3
47.8 0.6
22.1 1.4
33.7 0.9
88.3 0.9
38.4 1.2
23.1 1.0
7.0
29.3 2.0
3.1 0.9 3.8 1.1 19.0 5.2 79.4 2.1 52.7 10.2 4.1 0.6 3.6 2.3 3.1 1.0 2.0 1.2
44.6 4.6
21.9 1.6
34.1 1.1
88.4 0.8
38.5 1.0
22.6 0.9
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Fig. 1. Impact of degree of esterification (DE 5.4%, 55%, 71%) on the electrical properties (z-potential) of mixed biopolymer dispersions (WPI:Pectin ratio 1:2, 1:1, 2:1, 5:1, 8:1, cbiopolymer = 0.6%) fabricated at pH 7 (A), 5 (B), and 3 (C).
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Fig. 2. Impact of degree of esterification (DE 5.4%, 55%, 71%) on the turbidity (T600nm) of mixed biopolymer dispersions (WPI:Pectin ratio 1:2, 1:1, 2:1, 5:1, 8:1, cbiopolymer = 0.6%) fabricated at pH 7 (A), 5 (B), and 3 (C).
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pectin (P5.4) typically possessed a large number of deprotonated carboxyl groups contributing to a high overall charge density, whereas high-methoxylation (P55 and P71) leads to a lower magnitude of negative surface charges indicated by lower z-potential values regardless of pH (Table 1). Moreover, turbidity values remained unchanged over the entire pH range due to strong electrostatic repulsion a fact that is in good accordance with other studies (Jones, Lesmes, Dubin, & McClements, 2010a; Salminen & Weiss, 2013). According to these results, WPI molecules were positively and pectin molecules were negatively charged below pH 4.5, which would be expected to induce electrostatic attraction between these biopolymers. 3.2. Characterization of WPI:Pectin dispersions 3.2.1. Influence of pH and biopolymer ratio The purpose of this set of experiments was to investigate the impact of pH and biopolymer ratio on the formation of mixed complexes that are composed of WPI and pectins having different DE, namely 5.4%, 55%, and 71%. Therefore, stock biopolymer solutions were mixed at various WPI:Pectin ratios (1:2, 1:1, 2:1, 5:1, 8:1) under neutral conditions and then the pH was changed from 7 to 3, whereas the overall concentration remained constant (cbiopolymer = 0.6%). All biopolymer dispersions prepared were characterized in terms of z-potential, turbidity, optical light microscopy and particle size measurements (Figs. 1 and 3). z-Potential and turbidity measurements revealed that no complexes were formed under neutral conditions which can be attributed due to strong electrostatic repulsion between negatively charged WPI and pectin molecules. Moreover, no phase separation
of WPI and pectin molecules under neutral conditions occurred indicating that the phase separation threshold was not exceeded (de Kruif & Tuinier, 2001). When we decreased the pH to 5, the z-potential of the biopolymer dispersions remained still negative (Fig. 1), whereas turbidity values slightly increased indicating that both WPI and pectin molecules began to electrostatically attract each other regardless of DE. Previous studies have already shown that complexation of mixed biopolymer systems can be initiated slightly above the proteins pI due to positively charged patches on the proteins surface (Azarikia et al., 2015; Davidov-Pardo, Joye, & McClements, 2015; Mattison, Dubin, & Brittain, 1998; Park, Muhoberac, Dubin, & Xia, 1992; Weinbreck, de Vries, Schrooyen, & de Kruif, 2003). Complex coacervation was promoted by further lowering the pH to 3 due to strong electrostatic attraction between oppositely charged WPI and pectin molecules (Jones & McClements, 2010b; Thongkaew, Hinrichs, Gibis, & Weiss, 2015). In addition, higher turbidity and z-potential values as well as micrographs taken further proofed that complexes were formed (Fig. 3). In addition, the biopolymer ratio clearly affected the complex structures formed regardless of DE, in particular at pH 3 and 5. The surface charge of the complexes generated at pH 3 became increasingly positive as the protein concentration was increased a fact that could be attributed to a saturation of the pectin molecules by WPI. Similarities could be observed when multilayered emulsions are formed by electrostatic deposition method (Zeeb, Gibis, Fischer, & Weiss, 2012; Zeeb, Thongkaew, & Weiss, 2014). In general, the effects described at pH 3 were less pronounced at pH 5 as shown in Figs. 1–3 and are in line with our previous study (Stenger et al., 2016).
Fig. 3. Influence of degree of esterification (DE 5.4%, 55%, 71%) on the microstructure and phase separation behavior of mixed biopolymer dispersions (WPI:Pectin ratio 1:2, 1:1, 2:1, 5:1, 8:1, cbiopolymer = 0.6%) prepared at pH 3 (scale bar = 100 mm).
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Fig. 4. Rheological properties (storage modulus G' and loss modulus G'' at g = 0.3 %) of biopolymer dispersions (WPI, P55, P71) depending on processing temperature: q = 50– 55 C (A) and q = 90–95 C (B).
3.2.2. Influence of degree of esterification (DE) The purpose of this set of experiments was to determine the affect of the pectins DE incorporated into biopolymer complexes on their morphology, size and shape. For the clarity of sake, results obtained at pH 3 were highlighted since the impact of pectins DE was most pronounced under acidic conditions (Figs. 1–3). In general, our results demonstrated that the DE had a major impact on the surface charge, size and shape of the biopolymer complexes formed at pH 3 regardless of WPI:Pectin ratio. Images taken by optical light microscopy showed that mixed biopolymer complexes composed of WPI and P5.4 consisted of small and loosely linked particles forming a thick sediment layer, whereas a high DE led to the formation of large and dense aggregates, in particular at high WPI:Pectin ratios (Fig. 3). Moreover, a steep increase in turbidity was observed under acidic conditions, whereas WPI:Pectin samples phase-separated at biopolymer ratios 2:1, 5:1 and 8:1 indicating the formation of insoluble complexes that easily phase separated into an upper biopolymer-depleted and a lower biopolymer-enriched layer (Figs. 2 and 3). In addition, electrophoretic mobility measurements revealed higher surface
charges for WPI:P5.4 samples regardless of biopolymer ratio which indicates a strong electrostatic repulsion between the biopolymer complexes formed a fact that might help to explain the morphological differences between the mixed complexes. Jones et al. (2010b) already demonstrated that biopolymer complexes exhibited an increased surface charge when low-methoxylated pectins were utilized. In addition, similarities might be drawn to the aggregation process of proteins which is known to be chargedependent. As such, high electrostatic repulsion between native proteins lead to filamentous aggregates, whereas particulate matters might be formed at low electrostatic repulsion (Bryant & McClements, 1998; Langton & Hermansson, 1992). 3.3. Heat-induced concentration of biopolymer dispersions under acidic pH – impact of processing temperature The purpose of this set of experiments was to investigate the formation of concentrated biopolymer dispersions that could be potentially utilized as structure modulators for foods. Single and mixed dispersions (cbiopolymer = 6.0%, pH 3) were subjected to a
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simple heat treatment to tailor the overall water content, whereas two different processing temperatures were utilized (Figs. 4–7 , Table 2). In particular, temperature ranges below (q = 50–55 C) and above (q = 90–95 C) the protein’s denaturation were applied (Jones, Decker, & McClements, 2009a; Zeeb, Salminen, Fischer, & Weiss, 2013). We hypothesized that high processing temperatures and low water concentrations might promote the formation of irreversibly linked biopolymer particles which could not be incorporated into a food matrix. 3.3.1. Single biopolymers We initially evaluated the rheological behavior of single WPI and pectin solutions having various DE, namely 5.4%, 55%, and 71%, after water was removed by heat treatment (Fig. 4). Storage modulus G' and loss modulus G” damping factor tan d were utilized as key parameters to monitor the rheological changes as the water content was decreased over time. Typically, higher G' values represent an elastic behavior resulting in a gel-like network structure (Mezger, 2010). Biopolymer dispersions (cbiopolymer = 6.0%, pH 3) were therefore hydrated and subsequently subjected to temperature ranges below (q = 50–55 C) and above (q = 90–95 C) the proteins denaturation under continuous stirring. Interestingly, stock pectin samples with a DE of 5.4% could not be fully dissolved
at initial high concentrations without inducing a gel network. It has been previously reported that low-methoxylated pectins might interact with monovalent cations such as Na+ under acidic conditions promoting biopolymer gelation (Ström, Schuster, & Goh, 2014; Yoo, Fishman, Savary, & Hotchkiss Jr, 2003). We therefore excluded this pectin sample from all further heating experiments. In general, our results revealed that the water content had a major impact on the rheological properties of single biopolymer solutions regardless of processing temperature (Fig. 4). We observed a significant increase in G' of pectin dispersions with decreasing water content, whereas no gel-like behavior was determined at final water concentrations (G' > G”)’’’. Moreover, pectins with a DE of 71% showed larger G' values over the entire concentration range tested than pectins with an intermediate DE – a fact that could be attributed to the structural differences of the pectins tested. The citrus pectin is a high molecular weight (85 kDa) biopolymer with a highly branched configuration which leads to a large hydrated volume, whereas beet pectins are rather small (45 kDa) and globular-like biomolecules (Salminen & Weiss, 2013). In addition, a significant increase in G' of WPI solutions was observed when different heating temperature were applied. In particular, below a water content of 86% G' values steeply increased
Fig. 5. Impact of biopolymer ratio on rheological (storage modulus G' and loss modulus G'' at g = 0.3 %) properties of complex dispersions (generated at pH 3) obtained during heat treatment: WPI:P55 1:2 (A) and WPI:P55 8:1 (B).
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Fig. 6. Visual appearance of heat-treated WPI:P55 dispersions (pH 3) before and after heat treatment: q = 50–55 C (A) and q = 90–95 C (B).
at low heating, whereas elevated temperatures promoted an increase in elastic properties below 93% of water. This indicated that the temperature had a major impact on the rheological properties of WPI dispersions since globular proteins are known to be heat-sensitive leading to the formation of a gelled network (G' > G”). Previously, it was already shown that above the thermal denaturation temperature, globular proteins are prone to configurational changes thereby exposing reactive side groups (Hoffmann & Van Mil, 1997, 1999; Töpel, 2004). However, both processing temperatures utilized led to the formation of heat-induced WPIgels. 3.3.2. Mixed biopolymers Mixed WPI:Pectin dispersions generated at pH 3 were also subjected to various heat treatments (q = 50–55 C vs. q = 90–95 C) to fabricate concentrated dispersions, whereas three biopolymer ratios were tested (WPI:Pectin 1:2, 2:1, 8:1). Rheological and textural properties of heated biopolymer samples are shown in Figs. 5 and 6. The rheometry measurements revealed that higher processing temperatures led to an increase in G' values regardless of biopolymer ratio or degree of esterification. In addition, the elastic properties of mixed biopolymer dispersions were dominated by the pectin and protein proportion as the water content was decreased. High protein contents led to gel-like structures (biopolymer ratio 8:1), whereas G' values of biopolymer dispersions mainly composed of pectin (biopolymer ratio 1:2) were appreciably lower, in particular at high temperatures. These results displayed the behavior of single biopolymer solutions when subjected to various heat treatments. However, the concentration process had to be terminated at water concentrations between 60
and 70% for q = 50–55 C and 80% for q = 90–95 C, respectively, since sample viscosities steeply increased preventing a homogenously distributed biopolymer mixture by stirring. Moreover, final damping factors (tan d) decreased with decreasing water contents indicating the formation of gelled particles, in particular at high processing temperatures (Table 2). In addition, photographic images taken after heating the biopolymer dispersions noticeably showed differences in texture. As such, we observed fluid-like samples at low processing temperatures, whereas highly viscous and gritty structures were formed at high temperatures (Fig. 6). In previous studies, it was demonstrated that unfolded proteins increase their hydrodynamic volume due to the assimilation of large amounts of water which led to sponge-like aggregates – a fact that is in line with our findings (Kessler, 2002). Moreover, the thermal stability of WPI could be enhanced by the addition of pectin leading to differences in protein aggregation (Ibanoglu, 2005; Verheul & Roefs, 1998). The pH measured was not affected by the heating process and remained constant at pH 3. 3.4. Redispersion of concentrated WPI:Pectin complexes Biopolymer dispersions formed under acidic conditions and subsequently heat-concentrated under various temperatures (q = 50–55 C vs. q = 90–95 C) were redispersed in order to determine potential changes in size or morphology. We hypothesized that temperatures above the proteins denaturation might have induced configurational changes leading to the formation of large aggregates that could not be redispersed. As such, all samples concentrated were diluted to their initial water content of 94% using double-distilled water (pH 3), followed by stirring overnight
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Fig. 7. Particle size distribution of redispersed biopolymer dispersions (pH 3, q = 50–55 C): WPI:P55 1:2 (A), WPI:P55 8:1 (B), WPI:P71 1:2 (C), WPI:P71 8:1 (D).
Table 2 Final damping factor tan d (strain g = 0.3%) of heat-concentrated biopolymer complexes (pH 3) composed of WPI and pectin with various degree of esterification (DE 55 and 71%). WPI:Pectin dispersion
q = 50–55 C
q = 90–95 C
WPI:Pectin ratio
WPI:P55 WPI:P71
1:2
2:1
8:1
1:2
2:1
8:1
1.36 0.28 0.89 0.02
1.11 0.05 0.94 0.01
1.44 0.19 0.43 0.12
0.44 0.01 1.34 0.41
0.24 0.01 0.33 0.05
0.22 0.00 0.22 0.00
to fully hydrate the biopolymer complexes. Static light scattering was then used to characterize the reconstituted samples (Fig. 7). For the clarity of sake, particle size distributions of WPI:Pectin complexes heated to 50–55 C are shown, however, similar results were obtained at higher heating temperatures (Data not shown).
Light scattering results clearly indicated that all complexes subjected to elevated temperatures could be redispersed regardless of the WPI:Pectin ratio or degree of esterification (Fig. 7). The particle sizes of the biopolymer particles measured remained unchanged after an overnight redispersion – a fact that highlights the particulated structure of the biopolymer complexes regardless
B. Zeeb et al. / Food Structure 10 (2016) 10–20
of their electrical properties. However, a question that was raised is whether the biopolymer complexes are held together by covalent rather than non-covalent hydrophobic forces. Previous studies have already shown that proteins change their configuration above the thermal denaturation temperature under neutral conditions which might induce aggregation or flocculation due to the exposure of reactive sulfhydryl groups and hydrophobic patches (Hoffmann & Van Mil, 1997, 1999). However, Shimada and Cheftel (1989) have demonstrated that the formation of disulfide bonds under acidic conditions is negligible. This indicates that further investigations are needed to gain deeper insights into the interaction forces involved in heat-concentrated biopolymer complexes formed at low pH. 4. Conclusions The present study showed that mixed biopolymer dispersions composed of WPI and pectin could be generated by associative complexation under acidic conditions and subsequently concentrated utilizing a simple thermal treatment. In particular, the morphology, density and size of the biopolymer complexes could be modulated by varying the biopolymer ratio and the pectins DE, respectively. As such, low-methoxylated pectins led to the formation of small and dense particles due to strong electrostatic attraction between oppositely charged biopolymers regardless of WPI:Pectin ratio. Textural properties of biopolymer dispersions highly depend on the processing temperatures applied, whereas fluid-like samples at low processing temperatures and gritty structures were formed at high temperatures. Moreover, redispersed complexes remained their particulated structure without promoting aggregation or flocculation. Our results might have important implications in the fabrication of novel foods with enhanced functionalities utilizing biopolymer particles as potential fat replacer and texture modifier. Acknowledgements and funding This research project was supported by the German Ministry of Economics and Energy (via AiF) and the FEI (Forschungskreis der Ernährungsindustrie e.V., Bonn): Project AiF 17876 N. References Akhtar, M., Dickinson, E., Mazoyer, J., & Langendorff, V. (2002). Emulsion stabilizing properties of depolymerized pectin. Food Hydrocolloids, 16, 249–256. Azarikia, F., Wu, B.-C., Abbasi, S., & McClements, D. J. (2015). Stabilization of biopolymer microgels formed by electrostatic complexation: Influence of enzyme (laccase) cross-linking on pH, thermal, and mechanical stability. Food Research International, 78, 18–26. Bryant, C. M., & McClements, D. J. (1998). Molecular basis of protein functionality with special consideration of cold-set gels derived from heat-denatured whey. Trends in Food Science and Technology, 9, 143–151. Davidov-Pardo, G., Joye, I. J., & McClements, D. J. (2015). Food-Grade protein-Based nanoparticles and microparticles for bioactive delivery: Fabrication, characterization, and utilization. Advances in Protein Chemistry and Structural Biology. de Kruif, C. G., & Tuinier, R. (2001). Polysaccharide protein interactions. Food Hydrocolloids, 15, 555–563. Dickinson, E. (1998). Stability and rheological implications of electrostatic milk protein – Polysaccharide interactions. Trends in Food Science and Technology, 9, 347–354. Dickinson, E. (2010). Flocculation of protein-stabilized oil-in-water emulsions. Colloids and Surfaces B: Biointerfaces, 81, 130–140. Hoffmann, M. A. M., & Van Mil, P. J. J. M. (1997). Heat-Induced aggregation of b-Lactoglobulin: Role of the free thiol group and disulfide bonds. Journal of Agricultural and Food Chemistry, 45, 2942–2948. Hoffmann, M. A. M., & Van Mil, P. J. J. M. (1999). Heat-induced aggregation of b-lactoglobulin as a function of pH. Journal of Agricultural and Food Chemistry, 47, 1898–1905. Ibanoglu, E. (2005). Effect of hydrocolloids on the thermal denaturation of proteins. Food Chemistry, 90, 621–626.
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Jones, O. G., & McClements, D. J. (2010a). Biopolymer nanoparticles from heattreated electrostatic protein-polysaccharide complexes: Factors affecting particle characteristics. Journal of Food Science, 75, N36–N43. Jones, O. G., & McClements, D. J. (2010b). Functional biopolymer particles: Design, fabrication, and applications. Comprehensive Reviews in Food Science and Food Safety, 9, 374–397. Jones, O. G., & McClements, D. J. (2011). Recent progress in biopolymer nanoparticle and microparticle formation by heat-treating electrostatic proteinpolysaccharide complexes. Advances in Colloid and Interface Science, 167, 49–62. Jones, O. G., Decker, E. A., & McClements, D. J. (2009a). Formation of biopolymer particles by thermal treatment of beta-lactoglobulin-pectin complexes. Food Hydrocolloids, 23, 1312–1321. Jones, O. G., Decker, E. A., & McClements, D. J. (2009b). Formation of biopolymer particles by thermal treatment of b-lactoglobulin-pectin complexes. Food Hydrocolloids, 23, 1312–1321. Jones, O. G., Lesmes, U., Dubin, P., & McClements, D. J. (2010a). Effect of polysaccharide charge on formation and properties of biopolymer nanoparticles created by heat treatment of beta-lactoglobulin-pectin complexes. Food Hydrocolloids, 24, 374–383. Jones, O. G., Lesmes, U., Dubin, P., & McClements, D. J. (2010b). Effect of polysaccharide charge on formation and properties of biopolymer nanoparticles created by heat treatment of b-lactoglobulin-pectin complexes. Food Hydrocolloids, 24, 374–383. Kessler, H. G. (2002). Food and bio process engineering – dairy technology, Vol. 5, München: Publishing House A. Kessler. Krzeminski, A., Prell, K. A., Weiss, J., & Hinrichs, J. (2014). Environmental response of pectin-stabilized whey protein aggregates. Food Hydrocolloids, 35, 332–340. Langton, M., & Hermansson, A.-M. (1992). Fine-stranded and particulate gels of b-lactoglobulin and whey protein at varying pH. Food Hydrocolloids, 5, 523–539. 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Colloids and Surfaces A: Physicochemical and Engineering Aspects, 298, 99–107. Park, J. M., Muhoberac, B. B., Dubin, P. L., & Xia, J. (1992). Effects of protein charge heterogeneity in protein-polyelectrolyte complexation. Macromolecules, 25, 290–295. Salminen, H., & Weiss, J. (2013). Effect of pectin type on association and pH stability of whey Protein—Pectin complexes. Food Biophysics1–10. Salminen, H., & Weiss, J. (2014). Effect of pectin type on association and pH stability of whey protein-Pectin complexes. Food Biophysics, 9, 29–38. Samant, S. K., Singhal, R. S., Kulkarni, P. R., & Rege, D. V. (1993). Proteinpolysaccharide interactions: A new approach in food formulations. International Journal of Food Science & Technology, 28, 547–562. Schmitt, C., Sanchez, C., Desobry-Banon, S., & Hardy, J. (1998). Structure and technofunctional properties of protein-polysaccharide complexes: A review. Critical Reviews in Food Science and Nutrition, 38, 689–753. Scholten, E., Moschakis, T., & Biliaderis, C. G. (2014). Biopolymer composites for engineering food structures to control product functionality. Food Structure, 1, 39–54. Shimada, K., & Cheftel, J. C. (1989). Sulfhydryl group/disulfide bond interchange reactions during heat-induced gelation of whey protein isolate. Journal of Agricultural and Food Chemistry, 37, 161–168. Sperber, B. L. H. M., Schols, H. A., Cohen Stuart, M. A., Norde, W., & Voragen, A. G. J. (2009). Influence of the overall charge and local charge density of pectin on the complex formation between pectin and b-lactoglobulin. Food Hydrocolloids, 23, 765–772. Stenger, C., Zeeb, B., Hinrichs, J., & Weiss, J. (2016). Formation of concentrated biopolymer particles composed of oppositely charged WPI and pectin for food applications. Journal of Dispersion Science and Technology [null-null]. Ström, A., Schuster, E., & Goh, S. M. (2014). Rheological characterization of acid pectin samples in the absence and presence of monovalent ions. Carbohydrate Polymers, 113, 336–343. Töpel, A. (2004). Chemie und Physik der Milch, Vol. 3, Hamburg: Behr’s Verlag. Thongkaew, C., Hinrichs, J., Gibis, M., & Weiss, J. (2015). Sequential modulation of pH and ionic strength in phase separated whey protein isolate – Pectin dispersions: Effect on structural organization. Food Hydrocolloids, 47, 21–31. Turgeon, S. L., Laneuville, S. I., Stefan, K., Ian, T. N., & Johan, B. U. (2009). Protein + polysaccharide coacervates and complexes: from scientific background to their application as functional ingredients in food products. Modern biopolymer science. San Diego: Academic Press (pp. 327–363). Verheul, M., & Roefs, S. P. F. M. (1998). Structure of particulate whey protein gels: Effect of NaCl concentration, pH, heating temperature, and protein composition. Journal of Agricultural and Food Chemistry, 46, 4909–4916.
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Verheul, M., Roefs, S. P. F. M., & De Kruif, K. G. (1998). Kinetics of heat-Induced aggregation of bLactoglobulin. Journal of Agricultural and Food Chemistry, 46, 896–903. Weinbreck, F., de Vries, R., Schrooyen, P., & de Kruif, C. G. (2003). Complex coacervation of whey proteins and gum arabic. Biomacromolecules, 4, 293–303. Yoo, S. H., Fishman, M. L., Savary, B. J., & Hotchkiss, A. T. Jr. (2003). Monovalent saltInduced gelation of enzymatically deesterified pectin. Journal of Agricultural and Food Chemistry, 51, 7410–7417. Zeeb, B., Gibis, M., Fischer, L., & Weiss, J. (2012). Crosslinking of interfacial layers in multilayered oil-in-water emulsions using laccase: Characterization and pHstability. Food Hydrocolloids, 27, 126–136.
Zeeb, B., Salminen, H., Fischer, L., & Weiss, J. (2013). Impact of heat and laccase on the pH and freeze-Thaw stability of oil-in-Water emulsions stabilized by adsorbed biopolymer nanoparticles. Food Biophysics1–13. Zeeb, B., Thongkaew, C., & Weiss, J. (2014). Theoretical and practical considerations in electrostatic depositioning of charged polymers. Journal of Applied Polymer Science, 131, [n/a–n/a. Zeeb, B., Fischer, L., & Weiss, J. (2014). Stabilization of food dispersions by enzymes. Food and Function, 5, 198–213. Zeeb, B., Salminen, H., Fischer, L., & Weiss, J. (2014). Impact of heat and laccase on the pH and freeze-Thaw stability of oil-in-Water emulsions stabilized by adsorbed biopolymer nanoparticles. Food Biophysics, 9, 125–137.
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Research paper
Formation of concentrated particles composed of oppositely charged biopolymers for food applications – impact of processing conditions Benjamin Zeeba , Catrin Stengera , Jörg Hinrichsb , Jochen Weissa,* a Department of Food Physics and Meat Science, Institute of Food Science and Biotechnology, University of Hohenheim, Garbenstrasse 21/25, 70599 Stuttgart, Germany b Department of Soft Matter Science and Dairy Technology, Institute of Food Science and Biotechnology, University of Hohenheim, Garbenstrasse 21, 70599 Stuttgart, Germany
A R T I C L E I N F O
Article history: Received 30 May 2016 Received in revised form 13 October 2016 Accepted 19 October 2016 Available online 21 October 2016 Keywords: Whey protein isolate Degree of esterification Pectin Complex coacervation Concentrated complexes Biopolymer particles
A B S T R A C T
Associative complexation approaches are promising tools to generate mixed biopolymer particles with tailor-made physicochemical properties. However, association of oppositely charged biopolymers typically occurs under acidic conditions and low biopolymer concentrations. The present study aims to investigate intrinsic and extrinsic parameters to generate concentrated biopolymer dispersions which could be utilized as structuring agents in food matrices. We therefore utilized a multistep procedure. Firstly, solutions of whey protein isolate (WPI) and pectins with different degrees of esterification (DE) were mixed under neutral conditions at various ratios (WPI:Pectin 1:2, 1:1, 2:1, 5:1, 8:1). Secondly, the pH was adjusted to 3 to promote complex coacervation. Thirdly, complex dispersions were subsequently heat-treated to tailor their overall water content, whereas temperatures below (q = 50–55 C) and above (q = 90–95 C) the protein’s denaturation were applied. Phase separation behavior, microstructural, rheological and electrical properties of the complexes were investigated by surface charge, turbidity, particle size, rheometry and light microscopy measurements. Results revealed that complexes composed of WPI and pectin with a low DE formed rather small and dense particles, whereas large aggregates were observed when the pectins DE was increased. Concentrated complex dispersions with water contents 80% could be manufactured regardless of heating temperatures, whereas complexes maintained their particulate structures upon re-dispersion. Results are of importance for future studies where we intend to incorporate concentrated biopolymer particles as fat replacer in meat and dairy matrices. ã 2016 Elsevier Ltd. All rights reserved.
1. Introduction Numerous studies focus on fabrication methods to design novel food structures possessing specific functionalities in terms of texture, appearance, mouthfeel, and flavor (Scholten, Moschakis, & Biliaderis, 2014). In particular, self-assembly techniques gained enormous interest along food manufacturers and technologists allowing ones to precisely control the interaction forces between molecules and colloids on a molecular, nano, colloidal, and microscale level (Zeeb, Fischer, & Weiss, 2014). In general, structural design principles based on self-assembly are rather cost effective, feasible, and ecofriendly, but require fundamental knowledge of the relationship between different hierarchical levels. In particular, directed self-assembly approaches such as
* Corresponding author. E-mail address: [email protected] (J. Weiss). http://dx.doi.org/10.1016/j.foostr.2016.10.002 2213-3291/ã 2016 Elsevier Ltd. All rights reserved.
associative coacervation has been widely reported in literature (Dickinson, 1998; de Kruif & Tuinier, 2001; Jones & McClements, 2010b, 2011; Schmitt, Sanchez, Desobry-Banon, & Hardy, 1998). Although biopolymer complexes generated by associative phase separation are known to have promising functional properties, their use in real food matrices as texture modifier or fat replacer is rather limited. One major drawback of complexes used for food applications is their low concentration range to be applied to prepare stable biopolymer particles without gelation (Turgeon, Laneuville, Stefan, Ian, & Johan, 2009). In general, associative complexation of mixed protein-polysaccharide systems occurs under conditions were charged polymers electrostatically attract each other, whereas the physicochemical properties of the complexes generated can be tailored by both, intrinsic (e.g. biopolymer type, biopolymer flexibility, branching, protein:pectin ratio) and extrinsic factors (e.g. pH, ionic strength, temperature, shear stress, overall biopolymer concentration) (Azarikia, Wu, Abbasi, & McClements, 2015; Jones, Decker, &
B. Zeeb et al. / Food Structure 10 (2016) 10–20
McClements, 2009b; Jones, Lesmes, Dubin, & McClements, 2010b; Jones & McClements, 2010a; Krzeminski, Prell, Weiss, & Hinrichs, 2014; Samant, Singhal, Kulkarni, & Rege, 1993; Schmitt et al., 1998). In a previous study conducted in our lab, we could demonstrate that high WPI:Citrus pectin ratios led to positive surface charges promoting the formation of large and dense particles that easily precipitated. In addition, complexes composed of WPI and highmethoxylated pectins had a relatively small and monomodal particle size distribution (Salminen & Weiss, 2013). Moreover, it was shown that the local overall charge density of pectins can be controlled by their degree of methyl-esterification – a key parameter to modulate the electrostatic attraction between pectins and their oppositely charged counterpart (Sperber, Schols, Cohen Stuart, Norde, & Voragen, 2009). As such, low-methylated pectins (DE < 50 %) are known to have a high overall charge, whereas high-methylated pectins (DE > 50 %) are pectins with a low overall charge (Akhtar, Dickinson, Mazoyer, & Langendorff, 2002; May, 1990). In the current study, whey protein isolate and pectins with different degrees of esterification were utilized since they are commercially available as texture modifier, thickener, emulsifier, or gelling agents in many food products (Akhtar et al., 2002; Bryant & McClements, 1998; Leroux, Langendorff, Schick, Vaishnav, & Mazoyer, 2003). Krzeminski et al. (2014) demonstrated that WPI:Pectin complexes could be embedded into a skim milk yoghurt as fat replacer having a similar texture profile than their full-fat control. In addition, we recently conducted a study to generate concentrated biopolymer dispersions by a simple heat treatment (q = 90–95 C) to simulate textural and sensorial properties of low-fat products. It has been shown that concentrated complex dispersions with water contents 80 % could be manufactured and easily re-dispersed, whereas the complexes maintained their particulate matter (Stenger, Zeeb, Hinrichs, & Weiss, 2016). The objective of the present study, however, was to gain further insights in intrinsic and extrinsic factors affecting the formation of heat-concentrated biopolymer particles. We hypothesized that heating conditions above the protein’s denaturation temperature might yield in biopolymer complexes which are stable against redispersion, whereas incorporated pectins having a low DE may form small and dense structures due to strong electrostatic attraction. Therefore, whey protein isolate (WPI) and pectin (DE 5.4%, 55%, 71%) solutions were mixed at pH 7 at various biopolymer ratios and the pH was subsequently lowered to acidic conditions to promote complexation. Biopolymer dispersions were then heattreated using two different temperature ranges, namely q = 50– 55 C and q = 90–95 C, to tailor the overall water content. The results might have important implications for the preparation of biopolymer particles that could be utilized as fat replacers in dairy or meat products. 2. Materials and methods 2.1. Materials Whey protein isolate (WPI 895) was obtained from Fonterra (Auckland, New Zealand). As stated by the manufacturer, the composition of the WPI used was 93.9 % protein (69.2 % b-lactoglobulin, 14.2% a-lactalbumin, 3.3% bovine serum albumin, 2.1% immunoglobulin G, 1.6% glycomacropeptide, 1.2% proteose peptone 5), 4.7% moisture, 0.3 % fat, 0.4 % carbohydrates, and 1.5 % minerals. Pectin samples with various degrees of esterification (DE) were donated by Herbstreith & Fox KG (Neuenbürg, Germany), namely citrus pectin (DE 71%, P71, Pectin Classic CU 201, 84% galacturonic acid), sugar beet pectin (DE 55%, P55, Betapec RU 301, 65% galacturonic acid), and citrus pectin (DE 5.4%, P5.4, Pectin Classic CU 902, 85% galacturonic acid). The pectin samples were
11
used without further purification. Analytical grade sodium hydroxide (NaOH) and hydrochloric acid (HCl) were obtained from Carl Roth GmbH + Co. KG (Karlsruhe, Germany). All samples were prepared in double-distilled water. All concentrations are expressed in mass percentage (%w/w). 2.2. Fabrication of biopolymer complexes Biopolymer complexes were generated as previously described (Stenger et al., 2016). In brief, powdered WPI (max. 20%) and pectin (max. 4.5%) were dissolved in double-distilled water and stirred at room temperature overnight. After complete hydration, the biopolymer solutions were readjusted to pH 7.0 using 0.1 M and 1 M HCl and/or NaOH, respectively. Both, WPI and pectin stock solutions were then mixed at various WPI:Pectin ratios (1:2, 1:1, 2:1, 5:1, 8:1) and the pH was adjusted to pH 7, 5, and 3, respectively. The final biopolymer concentration ranged between cbiopolymer = 0.6 –6.0%, whereas single WPI and pectin solutions were used as control samples. 2.3. Particle size determination Particle size measurements were performed using a static light scattering instrument (Horiba LA-950, Retsch Technology GmbH, Haan, Germany). Samples from the biopolymer dispersions were withdrawn and diluted to a concentration of approximately 0.005% with an appropriate buffer to prevent multiple scattering effects. The measurement principle is based on the angular dependence of the intensity of the scattered light from a laser beam by the dilute colloidal dispersion. The Mie theory was used to calculate the droplet size distributions. A refractive index of 1.42 for the WPI: Pectin complexes was used as previously determined by Krzeminski et al. (2014). However, it has to be noted that particle size measurements have to be treated with some caution since the biopolymer complexes generated are not entirely spherical in shape. 2.4. Turbidity measurements A photometer (Agilent 8453 UV–vis Spectroscopy System, Agilent Technologies Inc., CA, USA) was used to determine the phase separation behavior of mixed biopolymer systems by measuring the turbidity at 600 nm. High turbidity values of samples indicate milky dispersions and were thus samples containing particles which were large enough to scatter light. Double distilled water was used as reference. 2.5. z-Potential measurements A particle electrophoresis instrument (Nano ZS, Malvern Instruments, Malvern, UK) was used to determine the z-potential of single biopolymer and complex dispersions. Samples were loaded into an appropriate cuvette and the z-potential was determined by measuring the direction and velocity that the droplets moved in the electric field applied. The Smoluchowski equation was utilized to calculate the z-potential. The z-potential measurements were made from two freshly prepared samples, and were carried out with four readings per sample. All samples were made in duplicate. 2.6. Rheological analysis Rheological analysis was conducted using a rheometer MCR300 (Anton Paar GmbH, Graz, Austria; Software: Rheoplus/32 V3.40). The measurement system used was a plate-to-plate geometry (d = 25 mm, PP25, Part No. 79044, Anton Paar). An amplitude
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B. Zeeb et al. / Food Structure 10 (2016) 10–20
sweep was conducted with 40 measuring points in the strain range between 0.1 and 100 % (frequency f = 1 Hz; measuring gap = 1 mm; q = 25 C). The measuring point duration was set to 10 s. At the beginning of each measurement, a relaxation time of 30 s was applied. Each sample was measured in duplicate, whereas the storage modulus G', loss modulus G”, and damping factor tan d were utilized as a key parameter to determine the rheological properties of heat-concentrated dispersions. However, storage moduli <0.1 Pa should be treated with some caution due to the detection limit of the instrument. 2.7. Optical microscopy All samples were gently mixed prior to analysis to ensure homogeneity. 10 mL of each sample were placed on an objective plate and a cover glass was carefully slid on top. Light microscopy images were taken with an axial mounted Canon Powershot G10 digital camera (Canon, Tokyo, Japan) mounted on an Axio Scope optical microscope (A1, Carl Zeiss Microimaging GmbH, Göttingen, Germany). 2.8. Heat-induced concentration of biopolymer complexes Complex dispersions (WPI:Pectin ratio 1:2, 2:1, 8:1, cbiopolymer = 6.0%) formed at pH 3 were subjected to a heat-induced concentration step, whereas temperatures below and above the proteins denaturation were applied: q = 50–55 C (Modell400, Memmert GmbH & Co.KG, Schwabach, Germany) and q = 90–95 C (Professional Hot Plate Stirrer, VWR, Radnor, PA, USA) (Dickinson, 2010). Dispersions were heated at a heating rate of 0.5 C/min and low shear rates (<100 s1). At regular time intervals, aliquots were withdrawn to gravimetrically determine the water content according to Eqs. (1) and (2). mi;biopolymer ð1Þ 100% W i ð%Þ ¼ 1 mi;total with mi;biopolymer ¼
1
W i1 mi1;total 100%
ð2Þ
where Wi is the water content of sample i at a given time. Phase separation behavior and rheological properties of heat-treated samples were determined. In addition, single WPI and pectin solutions served as control samples.
2.9. Reconstitution of heat-treated complexes Concentrated complex dispersions (WPI:Pectin ratio 1:2, 2:1, 8:1) by various temperature ranges (q = 50–55 C vs. q = 90–95 C) were redispersed to their initial water content (94.0%) using double-distilled water which was adjusted to pH 3. Particle size distributions and microstructure were then assessed after 12 h of reconstitution. 2.10. Statistical analysis All samples were prepared in duplicate. The averages and the standard deviations were calculated from all measurements of the duplicate samples using Excel Software (Microsoft Corporation, Redmond, WA, USA). 3. Results and discussion 3.1. Characterization of base biopolymer solutions Initially, we examined the influence of pH (7–3) on the electrical properties of the biopolymer solutions since electrostatic attraction is the major driving force for the association of WPI: Pectin complexes (Azarikia et al., 2015). In particular, the affect of the pectins DE (5.4%, 55%, 71%) on the z-potential and turbidity of biopolymer dispersions were identified when the pH was moved from neutral to acidic (Table 1). The electrical charge of WPI changed from 29.3 2.0 mV to +27.9 2.5 mV as the pH was decreased from 7 to 3 which could be attributed to the fact that the pH was altered from above to below the isoelectric point (pI) of the protein (Azarikia et al., 2015). In earlier published literature the pI of WPI was already identified between pH 5 and 4 (Salminen & Weiss, 2014; Zeeb, Salminen, Fischer, & Weiss, 2014). In addition, turbidity measurements revealed high WPI solubility above and below the proteins’ pI – a fact that could be attributed to strong electrostatic repulsion between protein molecules (Majhi et al., 2006; Verheul, Roefs, & De Kruif, 1998). All pectin samples used remained negatively charged when the pH was decreased from neutral to acidic conditions regardless of DE, whereas a slight increase in the z-potential was measured between pH 4.5 and 3.0 – a fact that can be attributed to the pKa values of the pectins studied (Neirynck et al., 2007). However, our results revealed that the DE had a clear impact on the electrical properties of the pectin dispersions. As such, low-methoxylated
Table 1 Surface charge properties (z-potential) and turbidity of single biopolymer solutions (cbiopolymer = 0.6%) as a function of pH (3–7): Whey protein isolate (WPI) and pectin with DE of 5.4%, 55%, and 71%, respectively. pH
WPI
P5.4
P55
P71
z-Pot. (mV)
T600nm (%)
z-Pot. (mV)
T600nm (%)
z-Pot. (mV)
T600nm (%)
z-Pot. (mV)
T600nm (%)
3.0
27.9 2.5
29.7 0.4
29.5 1.0
14.4 0.8
90.6 0.4
14.6 2.1
31.3 0.5
3.5
23.9 1.9
35.3 0.5
28.6 1.0
22.6 0.9
90.2 1.0
24.1 2.0
30.8 0.5
4.0 4.5 5.0 5.5
15.6 1.0 3.4 0.8 10.8 0.8 17.5 0.7
43.3 1.7 46.7 2.9 47.3 0.3 47.8 0.7
27.4 0.8 26.3 0.5 25.0 0.8 23.5 1.4
29.0 0.8 32.7 1.3 34.2 1.0 34.4 1.2
90.1 0.7 89.8 0.5 89.8 0.8 88.6 0.7
30.6 1.7 35.8 0.9 37.0 1.7 38.4 1.3
29.7 1.0 28.2 0.4 26.5 0.7 25.1 0.9
6.0
22.6 1.5
48.0 0.7
22.9 1.2
34.1 0.9
88.8 1.2
37.6 1.3
22.4 1.1
6.5
26.2 1.3
47.8 0.6
22.1 1.4
33.7 0.9
88.3 0.9
38.4 1.2
23.1 1.0
7.0
29.3 2.0
3.1 0.9 3.8 1.1 19.0 5.2 79.4 2.1 52.7 10.2 4.1 0.6 3.6 2.3 3.1 1.0 2.0 1.2
44.6 4.6
21.9 1.6
34.1 1.1
88.4 0.8
38.5 1.0
22.6 0.9
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Fig. 1. Impact of degree of esterification (DE 5.4%, 55%, 71%) on the electrical properties (z-potential) of mixed biopolymer dispersions (WPI:Pectin ratio 1:2, 1:1, 2:1, 5:1, 8:1, cbiopolymer = 0.6%) fabricated at pH 7 (A), 5 (B), and 3 (C).
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Fig. 2. Impact of degree of esterification (DE 5.4%, 55%, 71%) on the turbidity (T600nm) of mixed biopolymer dispersions (WPI:Pectin ratio 1:2, 1:1, 2:1, 5:1, 8:1, cbiopolymer = 0.6%) fabricated at pH 7 (A), 5 (B), and 3 (C).
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pectin (P5.4) typically possessed a large number of deprotonated carboxyl groups contributing to a high overall charge density, whereas high-methoxylation (P55 and P71) leads to a lower magnitude of negative surface charges indicated by lower z-potential values regardless of pH (Table 1). Moreover, turbidity values remained unchanged over the entire pH range due to strong electrostatic repulsion a fact that is in good accordance with other studies (Jones, Lesmes, Dubin, & McClements, 2010a; Salminen & Weiss, 2013). According to these results, WPI molecules were positively and pectin molecules were negatively charged below pH 4.5, which would be expected to induce electrostatic attraction between these biopolymers. 3.2. Characterization of WPI:Pectin dispersions 3.2.1. Influence of pH and biopolymer ratio The purpose of this set of experiments was to investigate the impact of pH and biopolymer ratio on the formation of mixed complexes that are composed of WPI and pectins having different DE, namely 5.4%, 55%, and 71%. Therefore, stock biopolymer solutions were mixed at various WPI:Pectin ratios (1:2, 1:1, 2:1, 5:1, 8:1) under neutral conditions and then the pH was changed from 7 to 3, whereas the overall concentration remained constant (cbiopolymer = 0.6%). All biopolymer dispersions prepared were characterized in terms of z-potential, turbidity, optical light microscopy and particle size measurements (Figs. 1 and 3). z-Potential and turbidity measurements revealed that no complexes were formed under neutral conditions which can be attributed due to strong electrostatic repulsion between negatively charged WPI and pectin molecules. Moreover, no phase separation
of WPI and pectin molecules under neutral conditions occurred indicating that the phase separation threshold was not exceeded (de Kruif & Tuinier, 2001). When we decreased the pH to 5, the z-potential of the biopolymer dispersions remained still negative (Fig. 1), whereas turbidity values slightly increased indicating that both WPI and pectin molecules began to electrostatically attract each other regardless of DE. Previous studies have already shown that complexation of mixed biopolymer systems can be initiated slightly above the proteins pI due to positively charged patches on the proteins surface (Azarikia et al., 2015; Davidov-Pardo, Joye, & McClements, 2015; Mattison, Dubin, & Brittain, 1998; Park, Muhoberac, Dubin, & Xia, 1992; Weinbreck, de Vries, Schrooyen, & de Kruif, 2003). Complex coacervation was promoted by further lowering the pH to 3 due to strong electrostatic attraction between oppositely charged WPI and pectin molecules (Jones & McClements, 2010b; Thongkaew, Hinrichs, Gibis, & Weiss, 2015). In addition, higher turbidity and z-potential values as well as micrographs taken further proofed that complexes were formed (Fig. 3). In addition, the biopolymer ratio clearly affected the complex structures formed regardless of DE, in particular at pH 3 and 5. The surface charge of the complexes generated at pH 3 became increasingly positive as the protein concentration was increased a fact that could be attributed to a saturation of the pectin molecules by WPI. Similarities could be observed when multilayered emulsions are formed by electrostatic deposition method (Zeeb, Gibis, Fischer, & Weiss, 2012; Zeeb, Thongkaew, & Weiss, 2014). In general, the effects described at pH 3 were less pronounced at pH 5 as shown in Figs. 1–3 and are in line with our previous study (Stenger et al., 2016).
Fig. 3. Influence of degree of esterification (DE 5.4%, 55%, 71%) on the microstructure and phase separation behavior of mixed biopolymer dispersions (WPI:Pectin ratio 1:2, 1:1, 2:1, 5:1, 8:1, cbiopolymer = 0.6%) prepared at pH 3 (scale bar = 100 mm).
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Fig. 4. Rheological properties (storage modulus G' and loss modulus G'' at g = 0.3 %) of biopolymer dispersions (WPI, P55, P71) depending on processing temperature: q = 50– 55 C (A) and q = 90–95 C (B).
3.2.2. Influence of degree of esterification (DE) The purpose of this set of experiments was to determine the affect of the pectins DE incorporated into biopolymer complexes on their morphology, size and shape. For the clarity of sake, results obtained at pH 3 were highlighted since the impact of pectins DE was most pronounced under acidic conditions (Figs. 1–3). In general, our results demonstrated that the DE had a major impact on the surface charge, size and shape of the biopolymer complexes formed at pH 3 regardless of WPI:Pectin ratio. Images taken by optical light microscopy showed that mixed biopolymer complexes composed of WPI and P5.4 consisted of small and loosely linked particles forming a thick sediment layer, whereas a high DE led to the formation of large and dense aggregates, in particular at high WPI:Pectin ratios (Fig. 3). Moreover, a steep increase in turbidity was observed under acidic conditions, whereas WPI:Pectin samples phase-separated at biopolymer ratios 2:1, 5:1 and 8:1 indicating the formation of insoluble complexes that easily phase separated into an upper biopolymer-depleted and a lower biopolymer-enriched layer (Figs. 2 and 3). In addition, electrophoretic mobility measurements revealed higher surface
charges for WPI:P5.4 samples regardless of biopolymer ratio which indicates a strong electrostatic repulsion between the biopolymer complexes formed a fact that might help to explain the morphological differences between the mixed complexes. Jones et al. (2010b) already demonstrated that biopolymer complexes exhibited an increased surface charge when low-methoxylated pectins were utilized. In addition, similarities might be drawn to the aggregation process of proteins which is known to be chargedependent. As such, high electrostatic repulsion between native proteins lead to filamentous aggregates, whereas particulate matters might be formed at low electrostatic repulsion (Bryant & McClements, 1998; Langton & Hermansson, 1992). 3.3. Heat-induced concentration of biopolymer dispersions under acidic pH – impact of processing temperature The purpose of this set of experiments was to investigate the formation of concentrated biopolymer dispersions that could be potentially utilized as structure modulators for foods. Single and mixed dispersions (cbiopolymer = 6.0%, pH 3) were subjected to a
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simple heat treatment to tailor the overall water content, whereas two different processing temperatures were utilized (Figs. 4–7 , Table 2). In particular, temperature ranges below (q = 50–55 C) and above (q = 90–95 C) the protein’s denaturation were applied (Jones, Decker, & McClements, 2009a; Zeeb, Salminen, Fischer, & Weiss, 2013). We hypothesized that high processing temperatures and low water concentrations might promote the formation of irreversibly linked biopolymer particles which could not be incorporated into a food matrix. 3.3.1. Single biopolymers We initially evaluated the rheological behavior of single WPI and pectin solutions having various DE, namely 5.4%, 55%, and 71%, after water was removed by heat treatment (Fig. 4). Storage modulus G' and loss modulus G” damping factor tan d were utilized as key parameters to monitor the rheological changes as the water content was decreased over time. Typically, higher G' values represent an elastic behavior resulting in a gel-like network structure (Mezger, 2010). Biopolymer dispersions (cbiopolymer = 6.0%, pH 3) were therefore hydrated and subsequently subjected to temperature ranges below (q = 50–55 C) and above (q = 90–95 C) the proteins denaturation under continuous stirring. Interestingly, stock pectin samples with a DE of 5.4% could not be fully dissolved
at initial high concentrations without inducing a gel network. It has been previously reported that low-methoxylated pectins might interact with monovalent cations such as Na+ under acidic conditions promoting biopolymer gelation (Ström, Schuster, & Goh, 2014; Yoo, Fishman, Savary, & Hotchkiss Jr, 2003). We therefore excluded this pectin sample from all further heating experiments. In general, our results revealed that the water content had a major impact on the rheological properties of single biopolymer solutions regardless of processing temperature (Fig. 4). We observed a significant increase in G' of pectin dispersions with decreasing water content, whereas no gel-like behavior was determined at final water concentrations (G' > G”)’’’. Moreover, pectins with a DE of 71% showed larger G' values over the entire concentration range tested than pectins with an intermediate DE – a fact that could be attributed to the structural differences of the pectins tested. The citrus pectin is a high molecular weight (85 kDa) biopolymer with a highly branched configuration which leads to a large hydrated volume, whereas beet pectins are rather small (45 kDa) and globular-like biomolecules (Salminen & Weiss, 2013). In addition, a significant increase in G' of WPI solutions was observed when different heating temperature were applied. In particular, below a water content of 86% G' values steeply increased
Fig. 5. Impact of biopolymer ratio on rheological (storage modulus G' and loss modulus G'' at g = 0.3 %) properties of complex dispersions (generated at pH 3) obtained during heat treatment: WPI:P55 1:2 (A) and WPI:P55 8:1 (B).
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Fig. 6. Visual appearance of heat-treated WPI:P55 dispersions (pH 3) before and after heat treatment: q = 50–55 C (A) and q = 90–95 C (B).
at low heating, whereas elevated temperatures promoted an increase in elastic properties below 93% of water. This indicated that the temperature had a major impact on the rheological properties of WPI dispersions since globular proteins are known to be heat-sensitive leading to the formation of a gelled network (G' > G”). Previously, it was already shown that above the thermal denaturation temperature, globular proteins are prone to configurational changes thereby exposing reactive side groups (Hoffmann & Van Mil, 1997, 1999; Töpel, 2004). However, both processing temperatures utilized led to the formation of heat-induced WPIgels. 3.3.2. Mixed biopolymers Mixed WPI:Pectin dispersions generated at pH 3 were also subjected to various heat treatments (q = 50–55 C vs. q = 90–95 C) to fabricate concentrated dispersions, whereas three biopolymer ratios were tested (WPI:Pectin 1:2, 2:1, 8:1). Rheological and textural properties of heated biopolymer samples are shown in Figs. 5 and 6. The rheometry measurements revealed that higher processing temperatures led to an increase in G' values regardless of biopolymer ratio or degree of esterification. In addition, the elastic properties of mixed biopolymer dispersions were dominated by the pectin and protein proportion as the water content was decreased. High protein contents led to gel-like structures (biopolymer ratio 8:1), whereas G' values of biopolymer dispersions mainly composed of pectin (biopolymer ratio 1:2) were appreciably lower, in particular at high temperatures. These results displayed the behavior of single biopolymer solutions when subjected to various heat treatments. However, the concentration process had to be terminated at water concentrations between 60
and 70% for q = 50–55 C and 80% for q = 90–95 C, respectively, since sample viscosities steeply increased preventing a homogenously distributed biopolymer mixture by stirring. Moreover, final damping factors (tan d) decreased with decreasing water contents indicating the formation of gelled particles, in particular at high processing temperatures (Table 2). In addition, photographic images taken after heating the biopolymer dispersions noticeably showed differences in texture. As such, we observed fluid-like samples at low processing temperatures, whereas highly viscous and gritty structures were formed at high temperatures (Fig. 6). In previous studies, it was demonstrated that unfolded proteins increase their hydrodynamic volume due to the assimilation of large amounts of water which led to sponge-like aggregates – a fact that is in line with our findings (Kessler, 2002). Moreover, the thermal stability of WPI could be enhanced by the addition of pectin leading to differences in protein aggregation (Ibanoglu, 2005; Verheul & Roefs, 1998). The pH measured was not affected by the heating process and remained constant at pH 3. 3.4. Redispersion of concentrated WPI:Pectin complexes Biopolymer dispersions formed under acidic conditions and subsequently heat-concentrated under various temperatures (q = 50–55 C vs. q = 90–95 C) were redispersed in order to determine potential changes in size or morphology. We hypothesized that temperatures above the proteins denaturation might have induced configurational changes leading to the formation of large aggregates that could not be redispersed. As such, all samples concentrated were diluted to their initial water content of 94% using double-distilled water (pH 3), followed by stirring overnight
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Fig. 7. Particle size distribution of redispersed biopolymer dispersions (pH 3, q = 50–55 C): WPI:P55 1:2 (A), WPI:P55 8:1 (B), WPI:P71 1:2 (C), WPI:P71 8:1 (D).
Table 2 Final damping factor tan d (strain g = 0.3%) of heat-concentrated biopolymer complexes (pH 3) composed of WPI and pectin with various degree of esterification (DE 55 and 71%). WPI:Pectin dispersion
q = 50–55 C
q = 90–95 C
WPI:Pectin ratio
WPI:P55 WPI:P71
1:2
2:1
8:1
1:2
2:1
8:1
1.36 0.28 0.89 0.02
1.11 0.05 0.94 0.01
1.44 0.19 0.43 0.12
0.44 0.01 1.34 0.41
0.24 0.01 0.33 0.05
0.22 0.00 0.22 0.00
to fully hydrate the biopolymer complexes. Static light scattering was then used to characterize the reconstituted samples (Fig. 7). For the clarity of sake, particle size distributions of WPI:Pectin complexes heated to 50–55 C are shown, however, similar results were obtained at higher heating temperatures (Data not shown).
Light scattering results clearly indicated that all complexes subjected to elevated temperatures could be redispersed regardless of the WPI:Pectin ratio or degree of esterification (Fig. 7). The particle sizes of the biopolymer particles measured remained unchanged after an overnight redispersion – a fact that highlights the particulated structure of the biopolymer complexes regardless
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of their electrical properties. However, a question that was raised is whether the biopolymer complexes are held together by covalent rather than non-covalent hydrophobic forces. Previous studies have already shown that proteins change their configuration above the thermal denaturation temperature under neutral conditions which might induce aggregation or flocculation due to the exposure of reactive sulfhydryl groups and hydrophobic patches (Hoffmann & Van Mil, 1997, 1999). However, Shimada and Cheftel (1989) have demonstrated that the formation of disulfide bonds under acidic conditions is negligible. This indicates that further investigations are needed to gain deeper insights into the interaction forces involved in heat-concentrated biopolymer complexes formed at low pH. 4. Conclusions The present study showed that mixed biopolymer dispersions composed of WPI and pectin could be generated by associative complexation under acidic conditions and subsequently concentrated utilizing a simple thermal treatment. In particular, the morphology, density and size of the biopolymer complexes could be modulated by varying the biopolymer ratio and the pectins DE, respectively. As such, low-methoxylated pectins led to the formation of small and dense particles due to strong electrostatic attraction between oppositely charged biopolymers regardless of WPI:Pectin ratio. Textural properties of biopolymer dispersions highly depend on the processing temperatures applied, whereas fluid-like samples at low processing temperatures and gritty structures were formed at high temperatures. Moreover, redispersed complexes remained their particulated structure without promoting aggregation or flocculation. Our results might have important implications in the fabrication of novel foods with enhanced functionalities utilizing biopolymer particles as potential fat replacer and texture modifier. Acknowledgements and funding This research project was supported by the German Ministry of Economics and Energy (via AiF) and the FEI (Forschungskreis der Ernährungsindustrie e.V., Bonn): Project AiF 17876 N. References Akhtar, M., Dickinson, E., Mazoyer, J., & Langendorff, V. (2002). Emulsion stabilizing properties of depolymerized pectin. Food Hydrocolloids, 16, 249–256. Azarikia, F., Wu, B.-C., Abbasi, S., & McClements, D. J. (2015). Stabilization of biopolymer microgels formed by electrostatic complexation: Influence of enzyme (laccase) cross-linking on pH, thermal, and mechanical stability. Food Research International, 78, 18–26. Bryant, C. M., & McClements, D. J. (1998). Molecular basis of protein functionality with special consideration of cold-set gels derived from heat-denatured whey. Trends in Food Science and Technology, 9, 143–151. Davidov-Pardo, G., Joye, I. J., & McClements, D. J. (2015). Food-Grade protein-Based nanoparticles and microparticles for bioactive delivery: Fabrication, characterization, and utilization. Advances in Protein Chemistry and Structural Biology. de Kruif, C. G., & Tuinier, R. (2001). Polysaccharide protein interactions. Food Hydrocolloids, 15, 555–563. Dickinson, E. (1998). Stability and rheological implications of electrostatic milk protein – Polysaccharide interactions. Trends in Food Science and Technology, 9, 347–354. Dickinson, E. (2010). Flocculation of protein-stabilized oil-in-water emulsions. Colloids and Surfaces B: Biointerfaces, 81, 130–140. Hoffmann, M. A. M., & Van Mil, P. J. J. M. (1997). Heat-Induced aggregation of b-Lactoglobulin: Role of the free thiol group and disulfide bonds. Journal of Agricultural and Food Chemistry, 45, 2942–2948. Hoffmann, M. A. M., & Van Mil, P. J. J. M. (1999). Heat-induced aggregation of b-lactoglobulin as a function of pH. Journal of Agricultural and Food Chemistry, 47, 1898–1905. Ibanoglu, E. (2005). Effect of hydrocolloids on the thermal denaturation of proteins. Food Chemistry, 90, 621–626.
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