casein complexation and phase behavior

Accepted Manuscript Title: Impact of pH, ionic strength and chitosan charge density on chitosan/casein complexation and phase behavior Authors: Lan Ding, Yan Huang, XiXi Cai, Shaoyun Wang PII: DOI: Reference:

S0144-8617(18)31449-8 https://doi.org/10.1016/j.carbpol.2018.12.015 CARP 14369

To appear in: Received date: Revised date: Accepted date:

20 April 2018 19 September 2018 7 December 2018

Please cite this article as: Ding L, Huang Y, Cai X, Wang S, Impact of pH, ionic strength and chitosan charge density on chitosan/casein complexation and phase behavior, Carbohydrate Polymers (2018), https://doi.org/10.1016/j.carbpol.2018.12.015 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Impact of pH, ionic strength and chitosan charge density on chitosan/casein complexation and phase behavior Lan Ding, Yan Huang*, XiXi Cai, Shaoyun Wang* College of Biological Science and Technology, Fuzhou University, Fuzhou,

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350108, China

* Corresponding author: Tel: +86-591-22866375

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Fax: +86-591-22866278

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E-mail address: [email protected] (Y. Huang)

Highlights

Stability and phase behavior of chitosan/casein complexes were systematically

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[email protected] (S.Y. Wang)

studied for the first time.

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Bridging flocculation was determined to be highly dependent on the charge density of polysaccharide. The complexation granted casein with higher stability over wider range of pH.

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The effect of pH and ionic strength on the complexation behavior was thoroughly

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investigated.

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Abstract The interaction between polysaccharides and proteins has attracted great interests and the phase behavior and colloidal properties of complexes between chitosan, a

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positively charged polysaccharide, and proteins still need careful investigation. Here,

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we use chitosan/casein as model system and monitor their complex size, zeta-potential, light scattering intensity and phase behavior as functions of chitosan deacetylation degree, pH and ionic strength. Chitosan flocculates casein at low polysaccharide

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concentrations, while stabilizes casein at moderate to high concentrations. Such

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flocculation and stabilization ability are dependent on the linear charge density of

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chitosan, where high charge density chitosan saturates casein surface charge and stabilizes the complexes at lower polysaccharide concentration. The complexes are

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stable at acidic pH where chitosan is highly protonated and the addition of NaCl leads

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to the reduction in both size and zeta-potential and facilitates complex aggregation. The

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phase maps are constructed to provide guidelines for further studies on chitosan/casein and other chitosan/protein complexes.

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Keywords: chitosan; casein; complexation; phase behavior

1. Introduction In many pharmaceutical and food formulations, proteins and polysaccharides are two of the major components whose interaction determines the functionality, stability 2

and texture of the system. The interaction between proteins and polysaccharides depends strongly on the types of macromolecules, (their molecular weight, functional groups distributions, biopolymer ratio and concentrations), and the pH and ionic strength of the suspending medium (Dickinson, 1998; Schmitt, Sanchez, Desobry-

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Banon, & Hardy, 1998). When proteins are mixed with polysaccharides, the interactions

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may lead to various types of phase behavior including homogeneous co-solubilization, segregative phase separation, associative phase separation, etc. (J.-L. Doublier, 2000; Xu, Luo, Liu, & McClements, 2017). These phase behaviors play a critical role in

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determining the properties of food hydrocolloids such as stability, rheology, texture,

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shelf life and so on (Chun et al., 2014; C. G De Kruif & Tuinier, 2001). Thus, the study

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on the phase behavior and interactions between proteins and polysaccharides is crucial

formulations.

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for understanding complex multi-component food systems and designing dairy

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Chitosan, obtained via deacetylation of chitin, is the only natural cationic

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polysaccharide which consists of β-(1-4)-2-acetamido-2-deoxy-β-D-glucopyranose and 2-amino-2-deoxy-β-D-glucopyranose (Bodmeier, Chen, & Paeratakul, 1989; Garcia-Fuentes & Alonso, 2012). It is the second most abundant polysaccharide in the

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world, and has been widely applied in food, biomedical and cosmetics industries, due to its low cytotoxicity, biocompatibility, biodegradability, antibacterial and mucoadhesive properties (Garcia-Fuentes & Alonso, 2012). Chitosan is hydrophilic, but the percentage of deacetylated monomers and the distribution of acetyl group on the chains 3

has a critical effect on its solubility and conformation in aqueous media (Lamarque, Lucas, Viton, & Domard, 2005). In general, larger degree of deacetylation leads to more glucosamine monomer and higher linear charge density in acidic medium. Meanwhile, a higher content of acetyl groups may increase the stiffness of the chains due to steric

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effect of larger acetyl group comparing to hydrogen atom (Berth, Dautzenberg, & Peter,

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1998). The pKa of chitosan is around 6.5, depending on the degree of deacetylation

(Sorlier, Denuzière, Viton, & Domard, 2001). At pH values below its pKa, the primary amine groups on chitosan become protonated and lead to the dissolution of chitosan

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(Gan & Wang, 2007). The resulting polycation allows strong electrostatic interactions

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with negatively charged small molecules and polyanions to form various types of

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complexes (e.g. colloidal particles, aggregates and coacervates) (Al-Qadi, Grenha, Carrion-Recio, Seijo, & Remunan-Lopez, 2012; Yuan, Wan, Yang, & Yin, 2014).

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Caseins are the major ingredient of milk proteins, mainly consisted of four

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phosphoproteins, αS1-, αS2-, β-, and κ-casein, with weight ratios of 4:1:4:1,

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respectively (Y. Liu & Guo, 2008). Their molecular weights are between 19 and 25 kDa and the average isoelectric point (pI) is between 4.6 and 4.8 (Elzoghby, El-Fotoh, & Elgindy, 2011). Caseins are amphiphilic proteins with a high heat stability, and the

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distinct hydrophobic and hydrophilic domains result in the self-assembly of casein molecules into stable micellar structures in aqueous solutions (Elzoghby et al., 2011). The four phosphoproteins are held together by hydrophobic interactions and the bridging of calcium phosphate nanoclusters that bound to phosphorylated serine 4

residues of the casein side chains (De Kruif & Holt, 2003). A large proportion of κcasein is located on micelle surface which endows steric stabilization by protruding into the serum phase with its hydrophilic glycosylated portion (Corredig, Sharafbafi, & Kristo, 2011; Kethireddipalli, Hill, & Dalgleish, 2011; C. G. De Kruif & Zhulina, 1996).

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The amphiphilicity and special peptide structure of casein facilitate their surface-

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activity and excellent emulsifying, gelling and water holding capacities, and ions/small molecules binding ability (Livney, 2010). Additionally, casein can interact with biomolecules to form complexes and conjugates with synergistic combinations of

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properties. L. Liu et al. (2012), for example, reported that the casein/xanthan gum

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complex at oil-water interface could grant protein stabilized food emulsion with higher

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stability over wider range of pH and ionic strength which was superior to casein or xanthan gum alone. Hence, the interaction between charged polysaccharides with milk

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proteins has been continuously exploited.

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While most studies have focused on the interactions between neutral and anionic

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polysaccharides with proteins(Chun et al., 2014; Dai, Jiang, Shah, & Corke, 2017; Girard & Schaffer-Lequart, 2008; Jones, Decker, & McClements, 2010; Kontogiorgos, Tosh, & Wood, 2009; Tobin, Fitzsimons, Chaurin, Kelly, & Fenelon, 2012), increasing

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attention has also been paid on investigating the phase behavior of chitosan/protein complexes and the effect of pH, ionic strength and biomacromolecule’s structure on chitosan/protein complexation (Elmer, Karaca, Low, & Nickerson, 2011; Kayitmazer, Shaw, & Dubin, 2005; Kayitmazer, Strand, Tribet, Jaeger, & Dubin, 2007; Koo et al., 5

2018; Xiong et al., 2016; Yuan et al., 2014). However, a comprehensive study on chitosan/casein complexation, and, the effect of chitosan deacetylation degree (DD) and polymer charge density on chitosan/protein complexation, has yet been reported, to the best of our knowledge. Chitosan has been demonstrated as a great encapsulation and

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delivery material due to their muco-adhesive and penetrating capability that promote

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the absorption of active compounds (Kawashima, 2001; Sogias, Williams, &

Khutoryanskiy, 2008). Therefore, some recent studies have focused on the potential of using chitosan/casein complexes as functional delivery system (Bing Hu, Ting, Zeng,

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& Huang, 2013; B. Hu, Wang, Li, Zeng, & Huang, 2011; Kurukji, Norton, &

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Spyropoulos, 2016; Razmi et al., 2013; Zhang et al., 2014). While systematic phase

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behavior analysis and phase map construction is the foundation for understanding the physicochemical properties of a colloidal system, the lack of such information would

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largely limit the further development of chitosan/casein and potentially other colloidal

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chitosan/protein complexes. To this end, we aim to carefully investigate the interaction

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between chitosan and casein, build a thorough phase map of chitosan/casein complexation, and illustrate the effects of chitosan deacetylation degree, pH and ionic strength on the phase behavior of chitosan/casein complexes.

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2. Materials and methods 2.1 Materials

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Casein (protein content > 90%) was provided by Shanghai Macklin Biochemical Co., Ltd (Shanghai, China). Chitosan (low molecular weight) was purchased from Sigma-Aldrich Life Science & Technology Co., Ltd (Wuxi, China). Sodium chloride (99.5%), Sodium hydroxide (99.5%), acetic acid (99.5%) and hydrochloride acid

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China). Deionized water was used throughout to prepare all solutions.

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solution (36%) were purchased from Sino pharm Chemical Reagent Co., Ltd. (Fuzhou,

2.2 Preparation of chitosan with different deacetylation degree (DD)

Chitosan samples with lower DD-values than the commercial chitosan were

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prepared by previously reported method (Huang, Cai, & Lapitsky, 2015). Briefly, 3 g

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of the commercial chitosan were dissolved in 100 mL 2% acetic acid solution, into

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which 100 mL ethanol were added, and the air bubbles were removed by sonication. Next, 160, 330 and 480 μL of neat acetic anhydride were added to the mixture to

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reacetylate commercial chitosan to various DD-values. The mixtures were stirred at

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room temperature overnight before dialysis, once against 15 mM NaCl and twice

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against distilled water in two days. The purified solutions were freeze dried to obtain the final products. The DD-value of commercial chitosan was 88%, while the DDvalues of the three reacetylated chitosan were 78, 67, and 61% as determined by NMR

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(Lavertu et al., 2003). The NMR data and DD-value calculation was shown in Fig. S1 and Table 1 in the supplementary data.

2.3 Preparation of stock solutions

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Casein (CA) solution was prepared according to procedures used with minor modifications (Song, Zhang, Yang, & Yan, 2009). Briefly, CA powder was dissolved in 0.01 M NaOH solution by heating at 70 ℃ for 10 min to obtain 6 mg/mL CA solution which was then centrifuged at 4 ℃ and 8000 rpm for 15 min to remove the undissolved

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solid (less than 5% of original weight). The 6 mg/mL stock chitosan (CS) solution was prepared by dissolving chitosan in 0.6% acetic acid by stirring constantly at room

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temperature for 1h at 500 rpm. All samples were kept at 4 ℃ overnight to ensure

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complete hydration.

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2.4 Preparation of CA-CS complexes and phase map construction

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CA and CS solutions were prepared at different concentrations (0.2, 0.6, 1, 2, 3, 4,

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5 and 6 mg/mL) by diluting the stock solutions. For samples with different ionic

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strength, NaCl was added into both CA and CS solutions at matching concentrations prior to the mixing. CA-CS complexes at different mass ratios were prepared by adding

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chitosan solutions into casein solutions of different concentrations at 1:1 volume ratio

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with 500 rpm stirring, after which the mixture pH was adjusted to the desired value using 1.0 M NaOH and 1.0 M hydrochloric acid and kept stir mixed for 1 hour. For all

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phase map samples, chitosan with 88% DD was used and their complex size, polydispersity index (PDI), light scattering intensity, and zeta-potential were monitored at room temperature for one month. Sodium azide (0.2 mg/mL final concentration) was

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added in mixtures to prevent microbial growth (Chang, Gupta, Timilsena, & Adhikari, 2016). 2.5 Dynamic and electrophoretic light scattering The particle size, PDI and light scattering intensity (expressed as derived count

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rate with unit of kcps) of all samples were determined by dynamic light scattering (DLS)

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with light source wavelength of 633 nm at 173°backscatter detection angle to avoid multiple light scattering. The zeta-potential was measured by electrophoretic light scattering (ELS) using Smoluchowski model. A Malvern Zetasizer (Nano ZS, Malvern

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2.6 Electron microscopy of CA-CS complexes

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Instruments Ltd., Worcestershire, UK) was used for these measurements.

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A high-performance digital imaging TEM (JEOL H-7650, Hitachi HighTechnologies Corporation, Tokyo, Japan) was used to characterize complex

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morphology. To prepare TEM samples, one drop of the complex suspension was placed

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on a copper grid and stained with 2% phosphotungstic acid, which was then air dried

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overnight. The grid was then placed in the microscope for imaging at 100 kV accelerating voltage, while the images were taken on a Gatan electron energy loss

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spectrometry system using a 6 eV energy slit.

2.7 Statistical analysis All the phase map experiments were conducted at least twice, and the other all conducted in triplicate. The data was expressed as the mean ± standard deviation where 9

feasible. All statistical analysis was carried out using Microsoft Excel™ and Origin Pro 2017 software.

3. Results and discussion

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3.1 Characterization of pure and mixed biopolymer solutions The pH plays a key role in complexation process, as it controls the ionization

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degree of the ionic groups and the strength of the electrostatic interaction between

charged moieties. Chitosan molecules were positively charged at acidic pHs due to the

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protonation of primary amine groups and gave a positive electrophoretic mobility of

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4.86×10-8 m2·V-1·s-1 (corresponding to a zeta-potential around 60 mV when assuming

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a spherical morphology, which will be used in this work for the convenience of

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discussion and comparison between biomacromolecules and their complexes) at pH 4.0,

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which gradually decreased to 1.83×10-8 m2·V-1·s-1 (zeta-potential of 25 mV) at its pKa of 6.5 where chitosan became insoluble (Fig. 1A). Meanwhile, with pI around 4.6, the

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zeta-potential of casein changed from positive to negative as pH increased from 4.0 to

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6.5 with a crossover point at pH 4.6, where the zeta-potential became 0 mV and the protein precipitated (Fig. 1A). To achieve the strongest interaction between chitosan

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and casein, pH 5.5, where the difference between chitosan and casein zeta-potential was the largest, was chosen for further study on the interaction between casein and chitosan (Fig. 1A) (Espinosa-Andrews et al., 2013; Weinbreck, Wientjes, Nieuwenhuijse, Robijn, & Kruif, 2004). 10

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Fig. 1. (A) The zeta-potential evolution of casein/chitosan mixture at different chitosan concentrations (CA = 1 mg/mL) and (B) the zeta-potential evolution of casein/chitosan

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mixture at different chitosan concentrations (CA = 1 mg/mL, 88% DD chitosan), where

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the shaded area represents the flocculation and phase separation of complexes and (C)

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the critical chitosan concentration of stablization (CA = 1 mg/mL, pHCA = 5.5). When adding chitosan, the zeta-potential of casein solution (1 mg/mL) increased

from -27 mV to a constant value at 35 mV as the polysaccharide concentration exceeded

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0.1 mg/mL (CA:CS = 1:0.1) (Fig. 1B). With 0.05 mg/mL CS (CA:CS = 1:0.05), the zeta-potential became zero, revealing the full neutralization of casein charge by chitosan at this mass ratio, which was accompanied by precipitation. This trend was similar to previous studies with chitosan and anionic surfactant micelles, where the 11

surfactant micelle surface was gradually covered by chitosan with a zeta-potential increase and a plus-minus crossover point at certain chitosan concentration (B. Hu et al., 2011; Li & McClements, 2013). In this experiment, at chitosan concentrations lower than 0.095 mg/mL, casein flocculated and precipitated likely due to the insufficiency

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of chitosan to fully cover the casein surface (Fig. 1B), where the reduced surface

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electrostatic potential increased the collision rate between casein micelles, and the

chitosan-free patches on casein surface allowed the chitosan molecules to attach simultaneously to more than one casein micelle upon contact to cause bridging

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flocculation (Dickinson, 1998), which led to the phase separation observed at low

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chitosan concentrations. Thus, an appropriate CA:CS ratio where a complete coverage

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of the casein micelle surfaces by chitosan molecules is crucial for preparing stable CACS complexes.

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It is worth mentioning that the chitosan concentrations for bridging flocculation

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and casein surface full saturation were directly related to chitosan linear charge density.

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Figure. 1C shows the critical chitosan concentrations below which casein would be flocculated. For commercial chitosan with 88% DD-value, the chitosan charge density (charged monomer ratio) increased from 59% at pH 6.0 to 88% at pH 4.0 (Sorlier et al.,

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2001), while the critical chitosan concentration decreased from 0.115 mg/mL to 0.02 mg/mL. For chitosan with different DD-values (61% to 88 %) at pH 5.5, the charged chitosan monomer ratio was proportional to its DD-value, which increased from 58% (61% deacetylated chitosan) to 73% (88% deacetylated chitosan). The critical 12

concentration, as a result, decreased from 0.115 mg/mL for 61% deacetylated chitosan to 0.095 mg/mL for 88% deacetylated chitosan. Thus, at high charge density, less chitosan was needed to fully cover casein surface to prevent bridging flocculation, and vice versa. Interestingly, the two curves of critical flocculation chitosan concentration

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perfectly overlapped, suggesting that, regardless of whether the pH or deacetylation

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degree was varied, chitosan charge density was the key determinant of the chitosan

concentration needed for casein bridging flocculation and surface saturation. This experimental result was also in agreement with previous modeling studies on Monte

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Carlo simulation of polyelectrolyte/protein interactions and chitosan induced latex

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flocculation (Ashmore, Hearn, & Karpowicz, 2001; Carlsson, Malmsten, & Linse,

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2003). Such close correlation between chitosan charge density and critical chitosan concentration also revealed that the interaction between chitosan and casein was mostly

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driven by electrostatic forces.

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3.2 Characteristics of CA-CS complexes

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3.2.1 Effect of chitosan concentration and DD on CA-CS complexes To further investigate the effect of chitosan DD-values on chitosan/casein

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complexation, 2 mg/mL casein solution was mixed with chitosan solutions with different concentrations and DD-values (61 ~ 88%) at 1:1 volume ratio to reach different final chitosan concentrations. When mixed with 88% deacetylated chitosan, the particle size increased from 300 nm to 1200 nm and PDI increased from 0.3 to 0.5 13

as the chitosan concentration increased from 0.1 to 3 mg/mL (Fig. 2A and 2B), indicating the larger complexes formation with broader size distributions. The zetapotential of CA-CS complexes remained stable around 35 mV for commercial chitosan (Fig. 2C), which is consistent with previous results in Fig. 1C. The light scattering

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intensity increased from 1.5 × 105 to 2.5 × 105 kcps as chitosan concentration increased

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from 0.1 mg/mL to 0.5 mg/mL, which reflected the formation of colloidal complexes. At chitosan concentrations above 0.5 mg/mL, however, the light scattering intensity dropped slightly likely due to the form factor effect (Pedersen, 1997) (see

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supplementary data for detailed discussion).

Fig. 2. Impact of deacetylation degree of chitosan on the (A) size; (B) PDI; (C) zeta14

potential; (D) light scattering intensity of CA-CS complexes (CA = 1 mg/mL). When the DD was reduced, the particle size dropped several hundred nanometers at all chitosan concentrations, while the PDI, zeta-potential and light scattering intensity all dropped slightly. Compared to the samples with higher DD-values, the light

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scattering intensity of samples with 61% DD chitosan increased slowly. This could be

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explained by the reduced charge density of chitosan at low DD, which led to less

extended chitosan molecules when adsorbed at casein surface and, as a result, a smaller complex structure. Besides, lower chitosan charge could lead to a higher surface

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coverage and enhanced steric stabilization of casein, which reduced the degree of

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aggregation and the size of the complexes. At high DD-value, chitosan chains with

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higher charge density could form a more extended structure on casein surface and attach to multiple CA, both of which could lead to larger complex sizes. The zeta-potential of

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complexes decreased with chitosan DD and reached a value of 25 mV with 61% DD

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chitosan. The reduction in zeta-potential could be easily explained by the reduced

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charge density of chitosan and a lower surface charge density of complexes when the casein surface was fully covered by chitosan. Generally, the complex size and PDI increased with the chitosan addition while the zeta-potential was not sensitive to the

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chitosan concentrations. At a fixed chitosan concentration, the complex size and surface potential increased significantly with the chitosan DD-values, and, the complexes with lowest chitosan DD (61%) possessed the smallest size with the weakest light scattering ability. 15

At 0.1 mg/mL CS (CA:CS = 1:0.1), the complex formed by chitosan with 88% DD was stable with particle size around 300 nm. As the DD-values reduced to 61, 67 and 78%, however, the bridging flocculation of complex occurred at 0.1 mg/mL chitosan concentration (no data point at 0.1 mg/mL chitosan concentration as shown in

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Fig. 2). This could be ascribed to the lower chitosan linear charge density which

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required more chitosan polymers to saturate casein surface charge, thus, prolonged the

occurrence of bridging flocculation at low chitosan concentrations. This trend was in good agreement with previous discussion (Section 3.1) and, again, suggested that the

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bridging flocculation is highly related to the charge density of chitosan.

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3.2.2 Electron microscopy of CA-CS complexes

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The morphology of casein and CA-CS complexes were observed by TEM. As

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shown in Fig. 3, both casein and CA-CS complexes shared the spherical shape and chitosan was coated on the casein micelle surface (Fig. 3). This is similar to previous

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TEM images of chitosan/casein phosphopeptide nanocomplexes (B. Hu et al., 2011).

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The CA-CS complexes also had larger size with a diameter around 200 nm compared to casein at 100 nm (Fig. 3). The complex size in TEM was smaller than the

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hydrodynamic diameters determined by DLS (Fig. 1 and Fig. 2), likely due to the dehydration during TEM sample preparation. The color difference between casein and CA-CS complexes was due to the phosphotungstic acid staining which did not affect the dark color of polysaccharides but led to light color of proteins, thus demonstrating 16

the full coverage of chitosan on the casein surface (K. Hu et al., 2015; Zhang et al.,

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2014; Zhu, Sun, Wang, Xu, & Wang, 2017).

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Fig. 3. TEM images of (A) 1 mg/mL casein; (B) CA-CS complexes at mass ratio 1:0.1.

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3.2.3 Effect of NaCl on CA-CS complexes

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To investigate the effect of ionic strength on the complexes, the casein/chitosan

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interaction was monitored at various NaCl concentrations. As shown in Fig. 4A, with

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the increase of NaCl concentration from 0 to 300 mM, the complex diameter and PDI decreased from 450 to 300 nm and 0.40 to 0.20, respectively. This could be explained

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by screening of electrostatic repulsion between chitosan chains at high ionic strength

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and diminishing of osmotic pressure difference between the solvent and the interior of complexes. Both effects could lead to deswelling and a closer pack of chitosan layer on

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casein surface and a smaller complex size (Protonotariou, Evageliou, Yanniotis, & Mandala, 2013; Rochefort, 1987). The smaller particle size also contributed to the decrease of light scattering intensity (Fig. 4B). It is noteworthy that there was a sharp decrease of light scattering intensity as the NaCl concentration increased from 100 and

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150 mM, above which this trend was not obvious. This could be ascribed to the reduction of particle size, and, the aggregation and sedimentation of particles at this high salt concentration during the storage. The electrostatic screening also led to the reduction of zeta-potential from 40 mV in salt free solution to 20 mV with 150 mM

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potential.

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and

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NaCl. In sum, the addition of monovalent salt, NaCl, led to the shrinkage of complexes

Fig. 4. Impact of NaCl concentration on the (A) size and PDI; (B) zeta-potential and light scattering intensity of complexes at 1 mg/mL CA (mass ratio CA:CS = 1:0.5).

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3.2.4 Effect of pH on CA-CS complexes Since the electrostatic interaction between chitosan and casein is highly dependent

on pH (Fig. 1), it is necessary to carefully investigate the effect of pH on the final complexes (Fig. 5A and 5B). At pH 4.0, where both casein and chitosan possess positive 18

charges (Fig. 1A) the complexation still occurred as indicated by the increased particle size (Fig. S2 in supplemented data). This may be because although casein micelles carried net positive charge under low pH, negatively charged patches could still exist on the micelle surface and interact with chitosan. Similar phenomenon was also

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reported in β-lactoglobulin/chitosan and pea protein/chitosan systems, where soluble

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complexes were formed at low pH because of the interaction between the positively charged chitosan and negatively charged patches on protein (Elmer et al., 2011; Mounsey, O’Kennedy, Fenelon, & Brodkorb, 2008). Upon raising the pH to around

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casein’s pI at 4.5 and 5, no casein precipitation occurred in contrast to the chitosan free

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casein solution. This could be attributed to the chitosan wrapped around casein, which

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provide steric repulsion and stabilization for casein micelles. When increasing pH, the size dropped from 660 to 450 nm at pH 5.5, and slightly increased to 480 nm at pH 6.0

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with PDI dropped from 0.5 to 0.4. Meanwhile, the light scattering intensity displayed

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an inverted U-shape, which increased from 1 × 105 to 3 × 105 kcps until pH 5.5, and

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then decreased remarkably to 1.8 × 105 kcps. This trend in size and light scattering intensity was ascribed to the strongest electrostatic interaction between casein and chitosan at pH 5.5 (Fig. 1A), which led to the most compact complexation that

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possessed the strongest light scattering ability.

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Fig. 5. Impact of pH on the (A) size and PDI; (B) zeta-potential and light scattering intensity of complexes at 1 mg/mL CA (mass ratio CA:CS = 1:0.5).

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3.3 Casein/chitosan phase diagram

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3.3.1 The effect of macromolecule concentrations on casein/chitosan phase behavior

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To systematically illustrate the phase behavior of chitosan/casein, their mixtures at pH 5.5 with macromolecule concentrations between 0.1 and 3 mg/mL were

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monitored over a month, and a phase map was constructed (Fig. 6A). The phase

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behavior of the mixtures was assessed by both DLS and visual observation as stable

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dispersion, flocculation or sedimentation.

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Fig.6. (A) Phase diagram of CA/CS complexes (: stable : flocculation : sedimentation); (B) picture of complexes in different states: 1) casein, 2) stable CA-CS complex, 3) flocculated CA-CS complex, 4) sedimented CA-CS complex. The white background in the middle of each vials was the labling sticker.

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As a control, casein solutions showed no phase separation at all tested

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concentrations and kept their original translucence over a month (Fig. 6B1). When

casein was mixed with chitosan, the solutions became opaquer indicating the formation of CA-CS complexes (Fig. 6B2). However, bridging flocculation occurred at low

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chitosan concentration (Fig. 6A and Fig. 6B3). This was consistent with previous data

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(Fig. 1C), where low chitosan concentration would lead to bridging flocculation due to

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insufficient casein surface coverage, so that one chitosan could attached to multiple casein molecules and micelles which caused rapid flocculation. As a result, most

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complexes rapidly precipitated, which led to the formation of a polymer rich precipitate

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phase and a clear supernatant phase. It is needed to clarify that, at casein concentrations

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lower than 1.5 mg/mL, flocculation also occurred at low chitosan concentrations, which is consistent with Fig. 1B, but was not labeled with data points due to the limited space

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on the phase diagram. At high CA concentrations (1.5 ~ 3 mg/mL), the complexes sedimented slowly

over time due to gravity and caused phase separation (Fig. 6A and Fig. 6B4). This was because of the slower Brownian motion and greater gravity force of large complex at high casein concentrations. Unlike the bridging flocculation at low chitosan 21

concentration, such sedimentation process was slow and might took several days to produce a visible phase separation layer at the bottom of vials. Interestingly, the sedimentation region of CA-CS complexes suggested that the process was more dependent on protein concentration rather than chitosan concentration. This can be

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attributed to complexes formed at a high protein concentration with a larger particle

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size and PDI which promoted the sedimentation by gravity. Although the high chitosan

concentration also increased the complex size and PDI, the excess chitosan that completely wrapped outside of casein provided strong electrostatic and steric repulsion

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and maintained the stability of complexes.

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3.3.2 The effect of NaCl concentration on casein/chitosan phase diagram

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To illustrate the effect of ionic strength on complex stability, we drew the phase diagram of complexes at various NaCl concentrations at pH 5.5. When casein was fixed

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at 1 mg/mL, no flocculation or sedimentation occurred at salt free condition (Fig. 7A).

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However, the addition of monovalent salt caused obvious flocculation at CA:CS = 1:0.1

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even at small NaCl concentration of 50 mM. As NaCl concentration further increased to 300 mM, the complexes all sedimented when chitosan concentrations exceeded 0.1 mg/mL. The promoted bridging flocculation at low chitosan concentration can be

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ascribed to the electrostatic screening by monovalent salt, which weakened casein/chitosan association, and more chitosan polymers were required to wrap casein to prevent the bridging flocculation. This salt effect was in accordance with the data in

22

Fig. 1C that chitosan charge density and electrostatic interaction was the determining

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factor of chitosan concentration needed for casein bridging flocculation.

Fig. 7. Phase diagram showing the influence of ionic strength (NaCl) at (A) 1 mg/mL

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casein concentration (B) 0.3 mg/mL chitosan concentration (: stable : flocculation

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: sedimentation).

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Comparing to polysaccharide concentration, the concentration of casein had a stronger impact on complex stability. When fixing CS concentration at 0.3 mg/mL, the

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increase of casein concentration resulted in a larger region of phase separation. At salt

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free condition, except the bridging flocculation point at CA:CS = 3:0.3, sediment

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formed as casein concentrations exceeded 1 mg/mL, which is consistent with previous data (Fig. 6A). Further increasing NaCl concentration to 50 ~ 300 mM promoted the bridging flocculation at CA:CS = 2.5:0.3. At higher casein concentrations (above 1

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mg/mL), sedimentation occurred easily with NaCl addition. In general, the addition of salt promoted flocculation and sedimentation, and

reduced complexes stability. The above phenomenon could be explained by influences of monovalent salt on casein/chitosan interactions. First, salt shielded the charge on the 23

surface of colloidal particles, resulting in a significant decrease of the electrostatic repulsion between the particles and accelerated the sedimentation, which was also confirmed by zeta-potential (Fig. 3B). Second, salt disrupted electrostatic interactions between protein and polysaccharides via shielding charged reactive sites on both

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biopolymers, so that more chitosan polymers were required for wrapping casein

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compactly, promoting the bridging flocculation at higher chitosan concentration

compared to free salt. The results suggest that casein/chitosan complexes only have a

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good stability in a narrow ionic strength range.

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3.3.3 The effect of pH on casein/chitosan phase diagram

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To investigate the effect of pH on CA-CS complex stability, the phase behavior of

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CA-CS complexes was investigated under various pH values (Fig. 8A and 8B). As a

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control, casein showed isoelectric point participation at pH 4.5 and 5.0, which was consistent with Fig. 1B. Meanwhile, with the addition of chitosan, casein no longer

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precipitated at pI, which has been discussed previously (Section 3.2.4). At pH 6.0,

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where was close to the pKa of chitosan, obvious phase separation occurred, and complexes flocculated at point CA:CS = 1:0.1. Meantime, sedimentation phase

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separation happened at chitosan concentrations beyond 0.1 mg/mL. At pH > pI, attraction interaction occurred between the positively charged chitosan and negatively charged casein.

24

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Fig. 8. Phase diagram showing the influence of pH at different mass ratio with (A) CA

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fixed at 1 mg/mL and (B) CS fixed at 0.3 mg/mL (: stable : flocculation : sedimentation).

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When fixing chitosan concentration at 0.3 mg/mL, the flocculation phase

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separation was found at CA:CS = 3:0.3 for all tested pH-values and at the point of

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CA:CS = 2.5:0.3, 2:0.3 at pH 6. At low casein concentrations, the complexes had good stability at pH 6, however, as casein concentration reached 1mg/mL, the sedimentation

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repulsion force.

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occurred easily at high pH-values due to reduced chitosan charge and electrostatic

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In summary, the colloidally non-stable region was located at high CA concentration and high pH-values. At high CA:CS mass ratios, when chitosan was not enough to completely wrap casein, one chitosan macromolecule bridged several casein

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and caused the bridging flocculation. As the pH-value increased, more chitosan macromolecules were required to wrap casein due to the charge reduction of chitosan, and, contributed to a broader chitosan concentration range for flocculation.

4. Conclusions 25

To investigate the complexation behavior between chitosan and casein, the phase map of chitosan/casein mixture was constructed for the first time with the effects of ionic strength and pH on the phase map systematically discussed. The chitosan flocculated casein at low polysaccharide concentrations and stabilized the complexes

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at higher concentrations where the amount of chitosan was sufficient to cover the casein

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surface. The critical chitosan concentrations for stable complexes were highly dependent on the chitosan linear charge density, which was adjusted by either changing pH or chitosan DD-value, and, provided identical critical concentration results with

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both methods. These results likely reflected that the electrostatic interaction was the

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major driving force for chitosan/casein complexation, while other forces (e.g. hydrogen

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bonding, hydrophobic interaction) played a minimal role during complexation. Further investigation on the phase map of chitosan/casein complexation revealed

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that colloidally stable complexes only existed at low casein concentrations. The

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addition of NaCl had little impact on complex stability until high salt concentrations

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where complexes flocculated or sedimented. The complexation with chitosan also granted casein a high stability at its pI and

provided colloidally stable complexes over broad pH ranges. The detailed and thorough

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information on CA-CS complexes obtained in this work provided theoretical support for future application of such complexes, and, could potentially benefit the design of other chitosan/protein complexes.

Acknowledgements 26

We are grateful to National Key R&D Program of China (No.2016YFD0400202)， National Natural Science Foundation of China (No.31801492), Qishan Scholar Program of Fuzhou University (XRC-1624) and Fuzhou University Open Measuring

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ED

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A

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Fund of Large Precious Apparatus (No.2017T023) for supporting this work.

27

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