Adsorptive Surface Modification of Cellulose Nanocrystals to Stabilize Nutraceuticals Loaded Lipid Carriers for Food Application

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Avinash S. Patel University of Maine, [email protected]

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Recommended Citation Patel, Avinash S., "Adsorptive Surface Modification of Cellulose Nanocrystals to Stabilize Nutraceuticals Loaded Lipid Carriers for Food Application" (2020). Electronic Theses and Dissertations. 3303. https://digitalcommons.library.umaine.edu/etd/3303

This Open-Access Thesis is brought to you for free and open access by DigitalCommons@UMaine. It has been accepted for inclusion in Electronic Theses and Dissertations by an authorized administrator of DigitalCommons@UMaine. For more information, please contact [email protected]. ADSORPTIVE SURFACE MODIFICATION OF CELLULOSE

NANOCRYSTALS TO STABILIZE NUTRACEUTICALS LOADED LIPID

CARRIERS FOR FOOD APPLICATION

Avinash Singh Patel

M.S. Banaras Hindu University, India, 2014

B.S. Mahatma Gandhi Kashi Vidyapith, India, 2012

A DISSERTATION

Submitted in Partial Fulfillment of the

Requirements for the Degree of

Doctor of Philosophy

(in Food and Nutrition Sciences)

The Graduate School

The University of Maine

December 2020

Advisory Committee:

Mary E. Camire, Professor of Food Science & Human Nutrition, Co-Advisor

Robert C. Causey, Professor of Animal and Veterinary Sciences, Co-Advisor

Carl P. Tripp, Professor of Chemistry

Mehdi Tajvidi, Associate Professor of Renewable Nanomaterials

Denise I. Skonberg, Professor of Food Science and Human Nutrition

ADSORPTIVE SURFACE MODIFICATION OF CELLULOSE NANOCRYSTALS

TO STABILIZE NUTRACEUTICALS LOADED LIPID CARRIERS FOR FOOD

APPLICATION

By Avinash Singh Patel

Dissertation Co-Advisors: Dr. Mary Ellen Camire and Dr. Robert Causey

An Abstract of the Dissertation Presented in Partial Fulfillment of the Requirements for the Degree of Doctor Philosophy (in Food and Nutrition Sciences) December 2020

Lipid carriers such as emulsions and liposomes have been widely studied as

carriers for different nutraceuticals. However, lipid carriers' physicochemical

instability results in the leakage of loaded nutraceuticals and the low solubility into the

digestion medium, thus reducing the bioaccessibility. Cellulose nanocrystals (CNC) are

nano-sized cellulose derivatives obtained after hydrolysis of cellulosic matrices that

possess potential applications in functional foods and nutraceuticals. However,

hydrophilicity, anionic surface potential, and poor re-dispersibility limit CNC’s

applications. The primary objectives of this research were 1) to modify CNC via

adsorbing polyethylene glycol (PEG) to stabilize liposomes and to study its

physiochemical stability at different in vitro gastrointestinal tract (GIT) conditions, pH

levels (1-11), temperatures (4-100 ºC) and illumination periods (4-72 hours); 2) to

modify CNC via adsorbing lauric acid to stabilize O/W emulsion and to study its

physicochemical stability at different pH (1-11), dilution factors (1-10 times), storage

period (up to 30 days) and GIT conditions. Phycobiliprotein and beta-carotene were

chosen as model nutraceuticals. iii

In study 1, particle size and microscopic analysis revealed a better physical/kinetic stability for the liposomes stabilized with PEG-adsorbed CNC than the liposomes stabilized with unmodified CNC, resulting in significantly (p≤0.05) smaller average particle size during the initial and mouth phases of the in vitro GIT study, nonetheless a significant (p≤0.05) increase in the particle size from ~300 nm to ~5500 nm at gastric phase due to lower pH. Moreover, stabilized liposomes retained the attributes of phycobiliprotein against pH, illumination, and a temperature of 60 ̊C. In study 2, lauric acid-adsorbed CNC affirmed the kinetic/physical stability of emulsions throughout the

GIT phases. Results exhibited a maximum of 65% beta-carotene bioaccessibility for the emulsion stabilized with modified CNC, and 71% for the emulsion stabilized with unmodified CNC. Moreover, a higher amount of lipid oxidation was recorded for the emulsion with stabilized modified CNC than the emulsion stabilized with gum Arabic.

Although modified CNC retains the kinetic/physical stability of lipid carriers; however, it increases lipid carriers' chemical instability and decreases the loaded nutraceuticals' bioaccessibility. A strategic modification of CNC is needed to ensure the stability of lipid carriers and increased nutraceuticals bioaccessibility.

DEDICATION

I dedicate this dissertation to my beloved mother, who taught me to believe in myself,

my father, who taught me to believe in hard work and dedication, and my teachers

and friends for providing me knowledge, devotion, and encouragement.

iii

ACKNOWLEDGMENTS

Fervently and modestly, I extol the genuine cooperation, inspiration and affection offered to me by the Co-Chairman of the advisory committee, Dr. Mary Ellen Camire,

Professor of Food Science & Human Nutrition, in completing the research work in time, successfully. The present work bears the impression of her concrete suggestions, guidance, and meticulous attention to detail.

I humbly place on record my respect and gratitude to Dr. Balunkeswar Nayak

(former advisor), previously Associate Professor, School of Food and Agriculture, for his valuable guidance and cooperation towards completing my research and study. I am grateful to members of my advisory committee, Dr. Robert Causey, Dr. Carl P. Tripp,

Dr. Mehdi Tajvidi, and Dr. Denise Skonberg, for constant motivation, rendered at all stages of this endeavor and my study period.

I record my profound thanks to the India Council of Agricultural Research (ICAR),

New Delhi, India, for awarding me with a prestigious Netaji Subhas International

Fellow award (award number: 18(02)/2016-EQR/Edn.) and financial support to perform my doctoral degree at the University of Maine. I want to thank the USDA

National Institute of Food and Agriculture Hatch project ME-021914, for financial support of my research work through the Maine Agricultural and Forest Experiment

Station (MAFES).

I owe a deep debt of thankfulness and heartfelt regards to Dr. Abhijit Kar, ICAR-

IARI, New Delhi for his endless inspirational help and Mr. Kuldeep Katiyar, Nicholas

Shoes, Pvt. Ltd., Greater Noida, India for his generous financial supports providing a

iv security bond of $10,000 FD (fix deposit) in favor of the ICAR, India until successful completion of the degree program.

My sincere thanks to the Intensive English Institute and Office of International

Programme for their kind support in my first year of classes and creating an overwhelming and friendly environment around the campus making this place feel like home.

I also record my sincere thanks to Dr. L. Brian Perkins, Katherine Davis-Dentici,

David Labrecque, and Kelly Edwards for their guidance and assistance on instrumentation and analysis. I would like to thank SuriyaPrakash

Lakshmibalasubrmaniam and my other past and current labmates and friends (Praveen

Sappati, Richa Arya, Adoum Fadaya Arabi, Samuel Akomea Frempong, Zach

Dijkhoff, Holly Leung, Rosanna Woodruff, Islam Hafez and all my SFA family members, UMaine, for helping me in one or the other manner during my research work.

Mere words of acknowledgment will never express my deepest sense of gratitude and indebtedness to my father Shri. Umashankar Patel, my brothers (Anurag and

Ashwani) and sisters (Mrs. Shalendri Patel and Mrs. Arati Patel), my family members, whose much-cherished love, inspiration ever encouraging moral support; selfless sacrifices enabled me to pursue my studies with unfailing zeal and devotion. To them,

I owe more than I say.

The words are not eloquent enough to express my special feeling for my mother

Smt. Basmati Devi, whose moral support, deep affection, encouragement, ever willing help and understanding always envisioned me throughout this research work.

Last but not least, none is forgotten, but all of them may not be mentioned.

TABLE OF CONTENTS

DEDICATION ...... v

ACKNOWLEDGMENTS ...... vi

CHAPTERS

1. INTRODUCTION ...... 1

1.1. Background ...... 1

1.2. Objectives ...... 5

2. LITERATURE REVIEW ...... 7

2.1. Nutraceuticals ...... 8

2.1.1. Phycobiliproteins (PBP) ...... 10

2.1.2. Characteristics of PBP ...... 11

2.1.3. Stability of PBP ...... 12

2.1.4. Carotenoids ...... 14

2.1.5. Stability of carotenoids ...... 15

2.2. Lipid carriers ...... 16

2.2.1. Oil-in-water emulsion: preparation and physicochemical stability ...18

2.2.2. Stabilization of O/W emulsion ...... 21

2.2.3. Pickering emulsion ...... 22

2.2.4. O/W emulsion and modified CNC ...... 24

2.2.5. Liposomes and stability ...... 25

2.3. Cellulose ...... 27

2.3.1. Cellulose nanomaterials ...... 29

2.3.2. Application of cellulose nanomaterials ...... 30

2.3.3. Cellulose nanocrystals ...... 32

3. STERIC STABILIZATION OF PHYCOBILIPROTEIN LOADED LIPOSOMES

WITH POLYETHYLENE GLYCOL ADSORBED CELLULOSE

NANOCRYSTALS AND THEIR IMPACT ON THE GASTROINTESTINAL

TRACT ...... 35

3.1. Introduction ...... 35

3.2. Materials and Methods ...... 38

3.2.1. Chemicals and reagents...... 38

3.2.2. Purification and characterization of phycobiliprotein...... 39

3.2.3. Quantification of phycobiliprotein ...... 40

3.2.4. Gel electrophoresis analysis of phycobiliproteins ...... 41

3.2.5. Adsorption of PEG with CNC ...... 42

3.2.6. Preparation and stabilization of PBP loaded liposomes ...... 42

3.2.7. In vitro gastrointestinal tract (GIT) study ...... 43

3.2.8. Determination of particle size and zeta potential ...... 46

3.2.9. Spectrofluorometer analysis of phycobiliprotein ...... 46

3.2.10. FTIR analysis ...... 46

3.2.11. Confocal laser scanning microscopic ...... 48

3.2.12. Statistical analysis ...... 48

3.3. Results and Discussion ...... 48

3.3.1. Phycobiliproteins in content the extract ...... 48

3.3.2. Gel electrophoresis of purified PBP ...... 49

3.3.3. Adsorption of PEG with CNC ...... 50

vii

3.3.4. Effect of PEG’s molecular weight on average particle size and zeta

potential of desulfated cellulose nanocrystals (DCs) ...... 51

3.3.5. Effect of gastrointestinal tract (GIT) phase on the quality

characteristics of phycobiliprotein (PBP) loaded liposomes ...... 54

3.3.5.1. Effect on average particle size ...... 54

3.3.5.2. Effect on zeta potential ...... 60

3.3.5.3. Stability of phycobiliprotein ...... 62

3.4. Conclusion ...... 63

4. IMPROVED STABILITY OF PHYCOBILIPROTEIN WITHIN LIPOSOME

STABILIZED BY POLYETHYLENE GLYCOL ADSORBED CELLULOSE

NANOCRYSTALS ...... 64

4.1. Introduction ...... 64

4.2. Materials and Methods ...... 68

4.2.1. Adsorption of PEG with CNC ...... 68

4.2.2. Encapsulation and stabilization of phycobiliproteins ...... 69

4.2.3. Encapsulation efficiency of phycobiliproteins ...... 70

4.2.4. Stability analysis of phycobiliproteins ...... 71

4.2.5. Particle size and zeta potential analysis ...... 72

4.2.6. Microscopic analysis ...... 72

4.2.7. Statistical analysis ...... 73

4.3. Results and Discussion ...... 73

4.3.1. Adsorption of PEG with desulfated CNC ...... 73

4.3.2. Effect of pH on average particle size ...... 74

viii

4.3.3. Effect of pH on zeta potential ...... 79

4.3.4. Effect on the coalescence and agglomeration of behavior of liposomes

and cellulose nanocrystals...... 81

4.3.5. Effect of pH on the stability of phycobiliproteins ...... 83

4.3.6. Effect of temperatures on the stability of phycobiliproteins ...... 86

4.3.7. Effect of illumination on the stability of the phycobiliproteins ...... 89

4.4. Conclusion ...... 90

5. PHYSICOCHEMICAL STABILITY OF O/W PICKERING EMULSION

STABILIZED WITH LAURIC ACID ADSORBED CELLULOSE

NANOCRYSTALS AND THE EFFECT OF DILUTIONS, pH, AND STORAGE

PERIODS ...... 92

5.1. Introduction ...... 92

5.2. Materials and Methods ...... 95

5.2.1. Adsorption of lauric acid with CNC ...... 95

5.2.2. Preparation of beta-carotene emulsion and stabilization ...... 98

5.2.3. Stability study of emulsions ...... 98

5.2.3.1. Effect of dilution ...... 99

5.2.3.2. Effect of pH...... 99

5.2.3.3. Effect of storage periods ...... 99

5.2.4. Particle size and zeta potential analysis ...... 100

5.2.5. Optical microscopic analysis ...... 100

5.2.6. Creaming index ...... 100

5.2.7. TBARS analysis ...... 101

5.2.8. Beta-carotene stability ...... 103

5.2.9. Mineral analysis ...... 103

5.2.10. Statistical analysis ...... 104

5.3. Results and Discussion ...... 104

5.3.1. Characterization of lauric acid adsorbed CNC (CL) ...... 104

5.3.2. Stability of Pickering emulsion ...... 106

5.3.2.1. Effect of dilutions on average particle size ...... 106

5.3.2.2. Effect of pH on average particle size ...... 111

5.3.2.3. Effect of storage on average particle size ...... 113

5.3.2.4. Effect on creaming index ...... 114

5.3.3. Oxidative stability of emulsions ...... 119

5.3.4. Beta-carotene stability ...... 122

5.4. Conclusion ...... 124

6. DEVELOPMENT OF O/W PICKERING EMULSION USING LAURIC ACID

ADSORBED CELLULOSE NANOCRYSTALS (CNC): AN IN VITRO

GASTROINTESTINAL TRACT STUDY ...... 125

6.1. Introduction ...... 125

6.2. Materials and Methods ...... 129

6.2.1. Adsorption of lauric acid with CNC ...... 129

6.2.2. Preparation of emulsions ...... 130

6.2.3. In vitro gastrointestinal tract (GIT) study ...... 131

6.2.3.1. Initial phase of the in vitro GIT ...... 132

6.2.3.2. Simulated saliva juice (SSJ) phase of the in vitro GIT ...... 132

6.2.3.3. Simulated gastric juice (SGJ) phase of the in vitro GIT ....132

6.2.3.4. Small-intestinal juice (SIJ) phase of the in vitro GIT ...... 132

6.2.4. Particle size and zeta potential analysis ...... 133

6.2.5. Creaming index ...... 133

6.2.6. Confocal laser scanning microscopic analysis ...... 134

6.2.7. Determination of lipid digestion ...... 134

6.2.8. Beta-carotene bioaccessibility ...... 135

6.2.9. Statistical analysis ...... 136

6.3. Results and Discussion ...... 136

6.3.1. Characterization of lauric acid adsorbed CNC (CL) ...... 136

6.3.2. Morphology of CNC and CL ...... 137

6.3.3. In vitro GIT fate of the Pickering emulsion ...... 138

6.3.3.1. Initial phase ...... 138

6.3.3.2. Simulated saliva juice (SSJ) phase ...... 140

6.3.3.3. Simulated gastric juice (SGJ) phase ...... 143

6.3.3.4. Simulated small-intestinal juice (SIJ) phase ...... 147

6.3.3.5. Lipid digestion ...... 149

6.3.3.6. Bioaccessibility of beta-carotene ...... 150

6.4. Conclusion ...... 152

7. OVERALL CONCLUSION AND RECOMMENDATIONS ...... 153

REFERENCES ...... 155

BIOGRAPHY OF THE AUTHOR ...... 184

LIST OF TABLES ...... xv

LIST OF FIGURES ...... xvi

xii

LIST OF TABLES Table 3.1. Chemical composition of simulated gastrointestinal tract (GIT) medium

used in vitro study ...... 45

Table 3.2. Phycobiliprotein contents in Palmaria palmata (Dulse) ...... 49

Table 5.1. Mineral contents of cellulose nanocrystals (CNC) ...... 121

xiii

LIST OF FIGURES

Figure 2.1. Hierarchical structure of cellulose from wood to monomeric unit ...... 28

Figure 3.1. Size exclusion-based column separation of phycobiliproteins ...... 40

Figure 3.2. Macroscopic images of purified PBP (A) and freeze-dried PBP (B) ...... 49

Figure 3.3. SDS-PAGE of phycobiliproteins. Marker is in far left ...... 50

Figure 3.4. A schematic of polyethylene glycol adsorption and FTIR spectra ...... 51

Figure 3.5. Average particle size and zeta potential of PEG-modified CNC ...... 52

Figure 3.6. Effect of the gastrointestinal phases on the average particle size ...... 55

Figure 3.7. Confocal laser scanning microscopic images of stabilized liposomes .....57

Figure 3.8. FTIR spectra of CNC before and after gastric phase of GIT ...... 59

Figure 3.9. Effect of gastrointestinal (GIT) phase on zeta-potential ...... 61

Figure 3.10. Stability of PBP during GIT phases ...... 62

Figure 4.1. FTIR spectra of PEG adsorbed CNC ...... 74

Figure 4.2. Effect of pH on the average particle size of stabilized liposomes ...... 76

Figure 4.3. Schematic illustration of liposomes and CNC behavior at lower pH ...... 78

Figure 4.4. Effect of pH range on the zeta potential value of stabilized liposomes ...80

Figure 4.5. Confocal laser scanning microscopic images of stabilized liposomes .....81

Figure 4.6. Optical microscopic image of samples at different pH ...... 82

Figure 4.7. Effect of stressors on the fluorescence intensity of phycobiliproteins .....83

Figure 4.8. Effect stressors on the % loss of fluorescence of phycobiliproteins ...... 87

Figure 5.1. FTIR spectra lauric acid adsorbed CNC and calibration curve ...... 97

Figure 5.2. Calibration curve of 1,1,3,3-tetraethoxypropane (TEP) ...... 102

Figure 5.3. Calibration curve of beta-carotene ...... 103

xiv

Figure 5.4. FTIR spectra of cellulose nanocrystal and lauric adsorbed CNC ...... 105

Figure 5.5. Effect of dilution factors on the average particle size emulsions ...... 108

Figure 5.6. Optical microscopic image of the stabilized emulsions ...... 109

Figure 5.7. Schematic illustration of the stabilization of emulsions ...... 110

Figure 5.8. Macroscopic image of stabilized emulsions ...... 110

Figure 5.9. Effect of pH on the average particle size of stabilized emulsions ...... 112

Figure 5.10. Effect of storage periods on the average particle size of emulsions ...... 114

Figure 5.11. Creaming index of the stabilized emulsions ...... 114

Figure 5.12. UV-Vis spectra of the beta-carotene of stabilized emulsions ...... 117

Figure 5.13. TBARS value of stabilized emulsions ...... 120

Figure 5.14. Stability of beta-carotene during storage ...... 123

Figure 6.1. FTIR spectra of CNC, lauric acid and modified CNC ...... 137

Figure 6.2. TEM images of CNC and modified CNC ...... 138

Figure 6.3. Effect of GIT phases on average particle size of emulsions ...... 139

Figure 6.4. Effect of GIT phases on zeta-potential of stabilized emulsions ...... 141

Figure 6.5. Confocal laser scanning microscopic images of stabilized emulsions ...142

Figure 6.6. Schematic illustration of fate of emulsions in small intestine ...... 145

Figure 6.7. Effect of GIT phases on creaming index of stabilized emulsion ...... 146

Figure 6.8. Release of free fatty acid during the small intestinal phase ...... 150

Figure 6.9. Bioaccessibility of the beta-carotene ...... 151

CHAPTER 1

INTRODUCTION

1.1. Background

In the last few decades, market demands have shifted towards healthy and low- calorie foods due to increased public health concerns, leading to the design and development of nutraceutical and functional foods (Nahr et al., 2018; Banerjee et al.,

2019; Bai et al., 2020a). Lipid nanocarriers such as emulsion and liposome-based delivery systems have been extensively utilized for several years as a carrier for hydrophilic and hydrophobic nutrients and phytochemical compounds to improve their stability, solubility, targeted delivery, and controlled release (Cagnasso et al., 2010;

McClements, 2015; Woodle et al., 2018; Ojeda-Serna et al., 2019). However, oil-in- water (O/W) and liposome-based delivery systems have several drawbacks that include kinetic or physical instability, oxidation of fatty acids, hydrolysis of the ester of the bond, leakage of loaded nutraceuticals, and poor solubility in the digestion medium

(Yadav et al., 2011; Feng et al., 2018; Mehrad et al., 2018; Peng et al., 2019). Synthetic and natural emulsifiers and stabilizers such as Tween®, sodium dodecyl sulfate (SDS), polyethylene glycol, glycerol, fatty acids, starch, gelatin, carrageenan, protein isolates, agar chitosan, and lecithin have been investigated to address these limitations of the lipid-based delivery systems (Carocho et al., 2014; Kono et al., 2018; Shen et al., 2018;

Zalipsky, 2018). Considering increased public health awareness and environmental sustainability, there has been a progressive effort to design and develop non-toxic, non- synthetic, biocompatible, low cost, and greener biopolymers for their potential applications in food and nutraceuticals industries. Due to unique features, cellulose and

1 its nanomaterials have inspired researchers to explore its potential application as fat replacer (Bai et al., 2020a), emulsifier (Angkuratipakorn et al., 2017), thickener

(Sviderskyi et al., 2018), gum (Jimenez et al., 2020) and food packaging material (Stark et al., 2016).

Cellulose is the most abundant biopolymer on the planet Earth, is a polysaccharide, and is a linear structure of anhydro D-glucose units condensed through a β-1,4 glycosidic bond (Peciulyte et al., 2015). Cellulose is predominantly present in wood, cotton, hemp, some bacterial pellicles, and other plant-based products as a principal structural material (Kargarzadeh et al., 2018). Due to its abundance in nature, researchers have investigated the isolation, purification, and characterization of cellulose and its derivatives, such as cellulose nanomaterials from different sources

(Chirayil et al., 2016; Prado and Spinace, 2019). Cellulose nanomaterials are micro/nano-scaled cellulose derivatives obtained after a series of processing methods, which include grinding, bleaching, homogenization/mechanical crushing, and chemical hydrolysis (Kargarzadeh et al., 2018). Nano-scaled biopolymers have distinctive features such as smaller dimension and higher surface area (Brinkmann et al., 2016), compatibility in nano-formulations, better dispersion and homogeneity, and more active surfaces such as hydroxyl groups (O-H) for targeted modifications (Cheng et al.,

2015; Natterodt et al., 2017; Kargarzadeh et al., 2018).

Cellulose nanocrystals (CNC), also known as crystalline nanocellulose, nanocrystalline cellulose, or cellulose nanowhiskers, are rod or needle-shaped nanoparticles with a dimension of 5-15 nm in width and 150-200 nm in length and up to 99% of crystallinity (Jordan et al., 2019; Abiaziem et al., 2019). CNC is produced

2 through different techniques such as acid hydrolysis (Kargarzadeh et al., 2018), acid- alkali hydrolysis (Jordan et al., 2019), enzymatic hydrolysis (Peciulyte et al., 2015), and metal salt catalyzed hydrolysis (Yahya et al., 2015). Among the CNC production methods, acid hydrolysis with sulfuric acid is most preferred because it contributes

- sulfate ions (-SO3 ) to the CNC. Sulfate ions on the surface of the produced CNC help in maintaining the dispersibility of the CNC particles into an aqueous medium by creating an electrostatic repulsion. Despite having a nano-scaled dimension and dispersibility into an aqueous medium, CNC also comprises free surface hydroxyl groups (O-H) (Natterodt et al., 2017), high charge potential due to sulfate ions

(Prathapan et al., 2016), and non-toxicity (Seabra et al., 2018). These properties of CNC offer an opportunity to explore potential applications.

Commercially produced CNC possesses a significant amount of sulfate anions (-

-3 OSO4 ), resulting in a zeta potential of approximately −55 mV (Cheng et al., 2015).

CNC with anionic properties and anionic liposomes (zeta potential −25 mV) may encounter a strong electrostatic repulsion that may limit the adsorption of CNC onto the surface of liposomes (Navon et al., 2017). Although a few researchers have reported the adsorption of anionic polymers on the interface of anionic liposomes due to Van der Waals attraction (Liu et al., 2013; Navon et al., 2017), a reduced zeta potential of

CNC can also assist in adsorption onto the interface of the liposome (Navon et al.,

2017). Moreover, it has been reported that CNC with reduced electric potential has shown agglomeration and precipitation behavior in an aqueous medium (Cheng et al.,

2015). Besides the instability and precipitation behavior of CNC with low electrical potential, Cheng et al. (2015) have reported that dried CNC had poor re-dispersibility

3 into an aqueous medium due to irreversible agglomeration of CNC particles caused by existing surface O-H groups. Therefore, it is necessary to ensure the dispersibility and the stability of CNC into aqueous medium sustaining its potential applications. Studies have shown that the adsorption of hydrophilic molecules such as polyethylene glycol

(PEG) maintained the dispersibility of the CNC (Cheng et al., 2015). Hence, we hypothesized that the adsorption of water-soluble molecules on the surface of CNC would help to maintain the dispersibility of CNC into the aqueous medium and stabilize liposome via adsorbing onto the interface, thus maintaining the stability of the liposome and the loaded nutraceuticals.

On the other hand, CNC in stabilizing O/W emulsions have shown a significant effect on maintaining the physicochemical stability of emulsion (Rayner et al., 2014;

Hedjazi and Razavi, 2018; Sharif and Zahid, 2018; Aswathanarayan and Vittal, 2019) by either electrostatic (Prathapan et al., 2016) or steric repulsion mechanism (Zoppe et al., 2012). Adsorption of CNC onto the interface of oil droplets may not be an easy target due to the hydrophilic nature of CNC, which has many hydroxyl groups (O-H).

However, several researchers have found that the addition of 0.06% of NaCl to the aqueous phase facilitated the adsorption of CNC particles onto the interface of oil droplets due to increased surface pressure (Bergfreund and Fischer, 2019; Bertsch et al., 2019). The addition of NaCl to the aqueous medium also increased the CNC-CNC interaction due to the counter-ion effect of sodium ions (Na+) altering the rheological properties, such as increasing the gel strength of the aqueous phase leading to higher viscosity. The increased gel strength of the continuous phase restricts the mobility of the free oil droplets (with no CNC on the interface), resulting in less interaction among

4 oil droplets, and thus a stable emulsion (Aaen et al., 2019). Moreover, a significant agglomeration and an increased particle size of CNC stabilized O/W emulsion have been reported due to a gradual dilution of the stabilized emulsion with simulated digestive juices and the addition of counter ions (Maphosa and Jideani, 2018; Bai et al., 2020a). Dilution of a CNC stabilized emulsion can lead to a separation of the oil phase from the aqueous phase due to the following reasons: (a) reduced surface pressure, thus desorption of polymer from the interface of oil droplet reducing steric or electrostatic repulsion among droplets, and (b) weaker gel strength of the dispersing medium, thereby increased droplets interaction thus coalescence/Ostwald ripening

(Wooster, 2006; Maphosa and Jideani, 2018; Yokota et al., 2019). Hence, we hypothesized that modifying CNC with hydrophobic or amphiphilic molecules properties can help to create a stable O/W emulsion by maintaining the hydrophilic and hydrophobic balance between CNC and lipids droplets.

1.2. Objectives

The main objective of this study was to modify the surface of CNC via adsorbing compatible food-grade polymers such as polyethylene glycol and fatty acids to ensure the dispersibility of CNC into aqueous medium and its suitability for the stabilization of liposomes and O/W type emulsion. One hydrophilic (phycobiliprotein) and one hydrophobic (beta-carotene) compound were chosen as model nutraceuticals. To ensure the physicochemical stability of the stabilized liposome, O/W emulsion, and loaded nutraceuticals, the effect of different stressors and static in vitro gastrointestinal tract (GIT) conditions were performed.

The specific objectives of this study are as follows:

1). To create a stable liposome using polyethylene glycol-adsorbed CNC and to study

the stability of the stabilized liposome during static in vitro GIT conditions.

2). To study the effect of different pH, illumination (exposure to light), and temperature

on the behavior of stabilized liposomes and the stabilization capacity for the

encapsulated phycobiliprotein.

3). Create a stable O/W type Pickering emulsion using lauric acid-adsorbed CNC and

to study the physicochemical behavior of the stabilized Pickering emulsion under

different dilution factors, pH, and storage period.

4). To study the physicochemical behavior of the stabilized Pickering emulsion during

static in vitro GIT conditions.

CHAPTER 2

LITERATURE REVIEW

According to the World Health Organization (WHO), good health is not just simply the absence of illness or debility but a complete physical, mental, and social well-being (Sartorius, 2006). Lifestyle behavior defines the state of human health, which can be constructive to health, but it can also harm. For example, a balanced and healthy diet improves our health; however, a diet deficient in essential nutrients or too high in energy results in malnutrition, decreased immunity, vulnerability to several preventable diseases, physical and mental discomfort, and compromised overall health status (Corzo et al., 2020). Due to hectic lifestyles, increased household income, urbanization, and availability of a variety of food options, a drastic change in regular diet habits have arisen in the last decade among the majority of the world population.

The consequences of several chronic diseases or conditions, including chronic heart disease (CVD), obesity, diabetes mellitus, high blood pressure, stroke, and certain types of cancer, have affected millions of people globally, resulting in the development of infirmity and premature death (Lynch and Smith, 2005; Silberner, 2018; Corzo et al.,

2020). These chronic diseases or conditions create immense burdens on the health and economy of developed areas like the United States, European Union, and Australia.

According to the National Center for Chronic Disease Prevention and Health

Promotion (NCCDPHP) of the CDC, 6 in 10 adults in the United States have a chronic disease, and 4 in 10 have two or more preexisting health conditions, costing around

$3.5 trillion annually (NCCDPHP, 2020). Even though the annual budget for public health is high, an average of 174 (Type 2 diabetes) and 134 (hypertensive disease)

7 deaths per million population was recorded in 2016 in the United States, which was higher than the average global deaths of 134 and 121 per million of the population

(Lynch and Smith, 2005; Editor, 2018). In the elderly, the prevalence and seriousness of chronic diseases are higher as compared to the younger age groups (Ge et al., 2019).

In 2016, the age group of 65 years and older was estimated to be 15% of the U.S. population, and is further estimated to increase up to 17% by the end of 2020 and 21% in 2030 (Vespa et al., 2019). According to the United States Census, Maine was the second oldest state in the country in 2015 and one-fifth of its population was above 65 years old. The median age of “Mainers” was 42.7 years, which was greater than the national median age of 37.2 years (U.S. Census, 2016). Therefore, with its significantly larger aged population, Maine residents are more likely to suffer from chronic diseases and face an increased healthcare burden.

Studies have shown a significant impact of diet and eating habits in modulating health and preventing chronic diseases (Costa et al., 2017). Diets rich in bioactive compounds, also known as nutraceuticals, have the potential in controlling and preventing several pathological mechanisms of chronic diseases and other health issues

(Ortega et al., 2019; Maqsood et al., 2020).

2.1. Nutraceuticals

Significant scientific explorations have led to notable developments of expensive, highly innovative techniques in pharmaceutical, medical, and surgical procedures. An emphasis on good health maintenance and the significance of proper diet has contributed in recent decades to increased interest in alternative medical therapies

(Kuttruff et al., 2014; Hettle et al., 2017; Zhang et al., 2020). Nutraceuticals are

8 substances that might be regarded as foods or part of foods and are responsible for providing medical or health interests. However, the United States government does not recognize a definition for these products. While consumer interests involve the prevention and cure of illnesses, the U.S. Food and Drug Administration restricts claims for those abilities to drugs, not foods (Burdock et al., 2006). Nutraceutical products extensively include special foods, isolated nutrients, diets, and dietary supplements. Foods fortified with nutraceuticals tend to be considered as functional foods that signify the food products consisting of health benefits that stretch beyond basic nutrition. Depending on the nation, the federal food authority may allow different products to claim to improve health status, delay aging, increase life expectancy, support body structure and function, and prevent chronic illnesses (Pandey et al., 2013;

Santini et al., 2017; Corzo et al., 2020).

Nutraceuticals are grouped based on food sources, mechanisms of action, and chemical nature. Food sources includes plants, animals, and microbes (Bao et al., 2019;

Corzo et al., 2020). Nutraceuticals are categorized according to their mechanisms of action as anti-cancerous, anti-inflammatory, osteogenic, antioxidants, anti-cardiac disease (Medeiros and Wildman, 2018). Other categories of nutraceuticals are based on chemical nature such as carotenoids, collagen hydrolysate, dietary fiber, fatty acids, flavonoids, glucosinolates, indoles, phenols, plant steroids, soy protein, phytoestrogens, and prebiotics (Medeiros and Wildman, 2018; Bao et al., 2019; Corzo et al., 2020). Further nutraceuticals can be classified as potential or established nutraceuticals (Pandey et al., 2010; Medeiros and Wildman, 2018).

Potential nutraceuticals hold a significant promise for a particular health or medical advantage. Potential nutraceuticals are only developed after having enough clinical data; thus, demonstrations are made to convince consumers and regulatory agencies. However, there are substantial scientific interventions underway to validate the nutraceuticals in diversified health applications (Sauer and Plauth, 2017; Sharif and

Khalid, 2018).

2.1.1. Phycobiliproteins (PBP)

PBP are regarded as the main light-harvesting chromo-proteins present in some seaweeds (Lage-Yusty et al., 2013). PBP are classified into four major classes according to their properties in light absorption as well as the types of bilins: phycoerythrins, phycocyanins, allophycocyanins, and the phycoerythrocyanins.

Phycoerythrins can further be divided into four classes: R-phycoerythrin (R-PE), B- phycoerythrin (B-PE), C-phycoerythrin (C-PE), and B-phycoerythrin (B-PE); they are grouped according to their origin and absorption spectrum. These absorption properties are determined by various types of bilin prosthetic groups (Pagels et al., 2019; Nikolic et al., 2020). R-PE has been identified as the most plentiful phycobiliprotein in red algae and marine unicellular cyanobacteria (Denis et al., 2010). Microalgae are a diversified category of photosynthetic aquatic microorganisms capable of growing in an extended range of habitats, pH, and temperatures. Microalgae tend to have uncomplicated growth requirements and use light, carbon dioxide, and other inorganic components efficiently (Manirafasha et al., 2016). Recently, microalgae have drawn researchers’ attention because of their high rate of growth and compound diversification, capable of accumulating large amounts of bioactive compounds such

10 as proteins, lipids, carbohydrates, carotenoids, and PBP (Eyley et al., 2015; Sadhukhan et al., 2019; Sappati et al., 2019; Nikolic et al., 2020).

Phycocyanin and phycoerythrin are photosynthetic pigments that are associated with a blue and red color in most of the red macro/microalgae. They are also found in cyanobacteria, rhodophytes, and cryptomonads (Falkeborg et al., 2018; Hsieh-Lo, et al., 2019). These pigments are grouped in the family of PBP, where the main commercial interest in these compounds stems from their high protein content and simple extraction techniques. Reports indicate that these proteins are identified to have health benefits like antioxidant, anti-inflammatory, anti-hypertension and anti-cancer properties (Lee et al., 2013; Furuta et al., 2016; Manirafasha et al., 2016; Pagels et al.,

2019). PBP is identified as food-grade if the ratio of Amax PBP / A280 is greater or equal to 0.7, reagent grade when Amax PBP / A280 ranges between 0.7 and 3.9 and analytical grade when Amax PBP / A280 is greater than or equal to 4.0 (Dumay et al., 2013; Rastogi et al., 2015).

2.1.2. Characteristics of PBP

Chlorophyll is a pigment responsible for photosynthesis by trapping light energy present in most plants and algae. It is worth noting that red algae lack chlorophyll b pigment; hence absorption of light occurs in the blue and red region within the visible spectrum because of chlorophyll a (Hsieh-Lo, et al., 2019). Compensation of significant absorption gaps and optimization of light energy gathered is achieved by assembling supramolecular complexes identified as phycobilisomes within the thylakoid membrane having an absorption range of 500-600 nm. Each phycobilisome consists of colored proteins known as PBP. Arrangement of these molecules can be

11 made in an antenna-like shape. The absorbed energy is channeled towards the center of the photosynthesis II reaction process, attaining an efficiency higher than 95%. PBP, like phycoerythrin, are regarded as chlorophyll accessory pigments that contain two primary subunits; α subunit (MW 12-19 kDa), β subunit (MW 14-21 kDa) and γ subunit

(MW 30-40 kDa), where each type consists of 160-180 amino acids (Galland-Irmouli et al., 2000; Furuta et al., 2016; Hsieh-Lo, et al., 2019). These molecules' color is mainly provided by the chromophore (prosthetic group) of the PBP, which is a linear tetrapyrrole group responsible for binding the apoprotein through a thioether bond to cysteine residues with its absorption spectrum depending upon the attached protein

(Kuddus et al., 2013; Martelli et al., 2014; Hsieh-lo et al., 2019).

2.1.3. Stability of PBP

PBP are soluble in water and produce intense colors. However, their utilization in foods has been insubstantial because of their low stability (Rastogi et al., 2015).

Notably, these molecules are extremely unstable in the presence of light, low pH, high temperatures, and as well in the presence of alcohol (Rastogi et al., 2015). pH changes might lead to disruption in the electrostatic properties and hydrogen bonds that are associated with protein’s structure, which has the capability of inducing structural chromophore changes (Kuddus et al., 2013; Martelli et al., 2014; Hsieh-lo et al., 2019).

Lower levels of pH might lead to the denaturation of PBP, which includes the dissociation of trimers to monomers (Ratogi et al., 2015). Moreover, under light, there is a high tendency to lose chromophores (PBP fluorescence capacity) (Munier et al.,

2014; Rastogi et al., 2015).

Researchers have come up with few strategies to improve pigment stability. The primary and most straightforward way to enhance PBP’s stability is by utilization of additives. Various studies have been established to investigate the application of additives, thus improving their thermal stability (Martelli et al., 2014; Gonzalez-

Ramirez et al., 2014; Braga et al., 2016). Nonetheless, low toxicity additives must be used due to the use of large amounts of additives that might be necessary. Martelli et al. (2014) recorded that high glucose concentration increases the thermal stability PBP.

Sugars and salts tend to act as protein stabilizing agents. The addition of sugars leads to an increase in surface tension in water, and as a result, the thermal stability of proteins is increased. A study by Braga et al. (2016) found improved thermal stability of PBP when nanofiber was incorporated into the solution, followed by the addition of polyethylene oxide and sorbitol as additives. Braga et al. (2016) indicated that the addition of nanofibers enhanced pigment thermal stability by lowering the system's enthalpy, thus reducing the denaturing of proteins. Benzoic acid is another additive that seems to be very interesting in improving PBP stability. Benzoic acid is known to act as an antimicrobial agent, thus inhibiting bacterial growth as well as antioxidant activity

(Kannaujiya et al., 2016). Crosslinking is a method where two chemical groups present on the protein surface tend to be covalently bound by the use of biofunctional reagents either intramolecularly or intermolecularly, thus reinforcing the folded structure (Wu et al., 2017). Bekasova et al. (2016) reported that when intramolecular crosslinking is applied between the protein molecule and silver nanoparticles, the stability of phycoerythrin was increased while the protein aggregation was unchanged.

Phycocyanin stabilized through covalent crosslinking between the subunits (alpha and

13 the beta) by formaldehyde successfully inhibits the dissociation of PBP into its subunits during treatment in SDS buffer at different temperatures (Sun et al., 2006). However, the use of formaldehyde in food-grade ingredients is questionable from a regulatory perspective. Alternatively, a complex formation can be another stabilization method such that proteins can be linked covalently to a polysaccharide, thus maintaining the extended tetrapyrrole structure (Hsieh-lo et al., 2019).

PBP products are primarily used in consumer products because of their possible application as natural food colorants in products like ice cream, chewing gum and candy (Sekar and Chandramohan, 2008). The application of PBP in food products is limited since several challenges are encountered in preserving these products for a longer duration without degradation (Rastogi et al., 2015). However, more studies are needed to improve PBP stability and thereby increasing the number of products in the market using PBP pigments as additives (Sekar et al., 2013; Furuta et al., 2016).

2.1.4. Carotenoids

Carotenoids are a group of hydrophobic compounds found in primarily terrestrial plants such as carrot, tomato, sweet potato, marigold flower, and colored mature leaves of several plants (Maoka, 2020). The carotenoids are long-chain hydrocarbon (CH) compounds containing 40 carbon atoms with an alternating double bond. The double bonds of the carotenoids are collectively known as isoprene units. Carotenoids comprise a total of eight isoprene units. Structurally, carotenoids are categorized into two groups: carotenes and xanthophylls. Carotenoids containing only carbon and hydrogen in its structure are recognized as carotene, while others containing carbon, hydrogen, and oxygen are xanthophylls (Xiao et al., 2018; Maoka, 2020). Typically,

14 carotenoids play a crucial role in photosynthesis and photoprotection processes in plant tissues. An essential role of carotenoid pigments included in the diets of humans and other animals is their potential to serve as precursors of vitamin A. The antioxidant function of carotenoids is responsible for reducing the risks of cancer, cataracts, and atherosclerosis (Maoka, 2020). Usually, carotenoids can be categorized based on their functionality, and whether they are primary or secondary carotenoids. These primary carotenoids are substances necessary for the photosynthetic process, while secondary carotenoids are elements that are not directly related to the plant's survival (Xiao et al.,

2018).

2.1.5. Stability of carotenoids

Among various food characteristics, color may be the most significant since it is often a consumer’s first impression or sensation (Singh, 2006; Viera et al., 2019).

Diverse food colorants have been studied to intensify food's original color. Carotenoids are responsible for many of the brilliant reds, orange, and yellow coloring in fruits, vegetables, and flowers (Maoka, 2020). Regrettably, several food processing functions, particularly ones that include the application of heat, are generally known to degrade food ingredients, and even change food coloring and its functionality (Boon et al.,

2010). Isomerization and oxidation are the significant reactions responsible for degradation of carotenoids. Generally, conjugated double bonds in carotenoids are known to exist in all-trans configuration. The carotenoids in plants exist mostly as trans-isomers, and trans- to cis-isomerization mainly occurs during food processing due to their large number of conjugated double bonds (Xiao et al., 2018). Several factors, such as heat, light, low pH, transition metals and other pro-oxidants, promote

15 the oxidation and isomerization of carotenoids. Moreover, exposure to organic solvents, irradiation and ozone can also induce oxidative degradation of carotenoids

(Boon et al., 2010).

Carotenoids are relatively stable typically during storage and as well as during the handling of most fresh and minimally processed fruits and vegetables. The apparent carotene content slightly tends to be affected by an increase in temperature, such as the blanching process. Blanching affects cell walls and thus makes carotenoids easier to extract than from raw plant tissues (Cilla et al., 2018; Mehmood and Zeb, 2020).

Natively, carotenes are considered reasonably stable at a lower temperature; however, heating at a higher temperature, such as those that occur during sterilization, can facilitate the induction of trans to cis isomerization reaction processes (Xiao et al.,

2018). Nevertheless, carotenoid isomerization results in a slight decline in biological activities and the color saturation, while oxidation terminates all operations and color in carotenoids. The creation of cis isomers provides a slight hypsochromic or blue shift of 3 to 5 nm, affecting mainly the color of the product. This can decrease the pro- vitamin A activity of the products (Xiao et al., 2018).

2.2. Lipid carriers

Lipids are prime nutrients widely applied in foods such as confections, butter/spreads, salad dressings, and ice creams. Physical, biotechnological, chemical, and biochemical techniques have been invented to modify lipids, thus coping with the present diversification of market demands within the lipids industry (Damodaran et al.,

2007). Among these diversifications, different practices such as replacement of saturated fatty acids with alternatives, reduction of trans-fatty acids, and improvement

16 of functionality of the end products have been widely used (Kabeva et al., 2017;

Wilczek et al., 2017; Bai et al., 2020b). Lipid contributes different physicochemical, tribological and mechanical attributes to end products; these attributes include glossiness, melting, texture, lubrication, friction, and spreadability (Guichard et al.,

2018; Tan et al., 2019).

Nowadays, the demand for food is not only limited to good sensory, tribological and satiety value but also more about a balance between these attributes with additional health implications. Thus, consumers are expecting food with multifunctional or nutraceutical ingredients besides essential nutrients. Foods with these multifunctional or nutraceutical ingredients are recognized as a functional food, which provides additional health benefits besides the nutritional values (Singh et al., 2018; Banerjee et al., 2019). In a functional food system, nano/micronization of multifunctional or nutraceutical ingredients with a compatible polymer or carrier/encapsulating materials is considered an ideal tool to improve its functionality and health benefits (Ko et al.,

2017; Da Silva et al., 2020).

The Definition of the nanomaterials/nanoparticles are commonly described as

"the understanding and monitoring the properties of materials at the nanoscale, which ranged from 1 to 100 nm” (Khan et al., 2019; Wang et al., 2019). At the nano dimension, the physicochemical properties such as color, solubility, reactivity, texture and toxicity of the particle vary, and in turn, so do the functional applications (Hydbring and Du, 2019; Khan et al., 2019; Wang et al., 2019). On the other hand, microparticles range from 1 to 1000 µm and have a relatively lesser influence on the physicochemical properties (Lengyel et al., 2019). The particle size of matrices/ingredients plays a key

17 role in designing and developing desirable functional foods, nutraceuticals, and medicines. The size-dependent application primarily includes enhanced bioavailability of bioactive compounds, efficient and targeted delivery, and increased solubility

(McClements, 2015; McClements, 2018; Lengyel et al., 2019).

Lipid’s biocompatibility and versatility traits have extensively led to several nano and microparticulate preparations in the past two decades. There are many merits portrayed by lipid nano/microparticles over the polymeric nanoparticles in delivering drugs and other bioactive compounds, hence they have a widespread application

(Tamjidi et al., 2013). Lipid nano/microparticle’s biocompatibility appears better than that of the polymeric nanoparticles since they consist of lipids that have a similarity to their physiological counterparts (McClements, 2018). Lipid particles or lipid carriers are nano/micro-sized lipids particles or droplets and are an ideal carrier for labile multifunctional and nutraceutical ingredients. Lipid carriers are categorized into two groups: solid lipid nanoparticles (SLN) and nano-lipid carriers (NLC) (Beloqui et al.,

2016). Nano-lipid carriers are considered more pertinent in delivering nutraceuticals than SLN due to higher loading capacity, faster release, and less expulsion capacity of

NLC. The most common types of NLC are nanoemulsion, microemulsion and liposome

(Beloqui et al., 2016; Khosa et al., 2018; Nahr et al., 2018).

2.2.1. Oil-in-water (O/W) emulsion: preparation and physicochemical stability

A wide range of food products that consumers use in daily life is produced from different immiscible phases, for example, a mixture of oil and water in salad dressing or mayonnaise. Consequently, the droplets of one liquid phase tend to be dispersed into the other, creating an emulsion (Gupta et al., 2016). There are two major categories of

18 emulsions: first, oil-in-water (O/W) emulsions, which involves dispersing of oil droplets into a continuous phase, and second, water-in-oil (W/O) emulsion, which involves dispersion of water droplets into an ongoing oil phase (Yang et al., 2017). In our food system, O/W type emulsions are found in milk, salad dressings, mayonnaise, ice-cream, and yogurt; however, a W/O type emulsion is found in butter, and margarine. A triple-phase emulsion can also be created by mixing oil-in-water-oil

(O/W/O) or water-oil-water (W/O/W) emulsions. Triple-phase emulsions are generally used in drug delivery (Tamjidi et al., 2013).

The utilization of O/W emulsion-based delivery systems in functional foods has been extensively explored (Siro et al., 2008; Sharif and Zahid, 2018) because an O/W emulsion is an ideal tool for the encapsulation of lipophilic micronutrients, protecting them against degradation during processing, storage, and consumption. The preparation of O/W emulsions, which is available commercially, can be done by applying low energy and high-energy methods. Low-energy approaches require an input energy density of 103-105 W kg-1; however, high-energy approaches utilize an input energy density of about 108-1010 W kg-1 (Gupta et al., 2016). Low-energy approaches include emulsion inversion or phase inversion method and input energy is not induced by any mechanical means other than the chemical activity of the components. A low-energy approach allows a narrower size distribution of the resulting emulsion. On the other hand, a high-energy approach utilizes energy input from mechanical devices such as microfluidizer, high-pressure homogenizer, or high shear blender. High-energy approach method results in wider size distribution of the oil droplets. However, due to higher emulsion production efficiency and scalability, the high-energy method is most

19 commercially applicable (Yang et al., 2012; Liu et al., 2020). High-energy methods also help in retaining the rheological attributes of emulsions and a higher concentration of dissolved nutraceuticals; however, the emulsion is diluted gradually, resulting in thinner emulsions with a lower nutraceutical concentration (Yang et al., 2012; Gupta et al., Liu et al., 2020). In high-energy methods, two steps are carried out separately by using a microfluidizer or high-pressure homogenizer. The first step involves a higher- shear blend to create a coarse emulsion by mixing up water, oil, and emulsifier. The second step consists of an action where the emulsion is forced through the microfluidizer (Silva et al., 2012). The micro-fluidization pressure facilitates the determination of the oil droplet size, emulsifier concentration, and emulsion stability.

Nevertheless, it is worth noting that the system has to be fed with sufficient emulsifiers that should be rapidly adsorbing to the oil droplet surfaces and stabilizing them against the aggregation (Al-Bukhaitia et al., 2018).

Since the O/W emulsion system consists of an oil-water interface, it is associated with positive free energy and is thus regarded as thermodynamically unstable. On this account, oil droplets tend to break down over time; therefore, reducing the contact area between both phases. This reduced contact results in separation of the oil phase from the aqueous phase over time (Vignati et al., 2003; Marquez et al., 2018). Additionally, oxidation of fatty acids and hydrolysis of the ester bond of the triglycerides results in the leakage of loaded nutraceuticals and poor solubility in the digestion media, thus reducing the bioaccessibility of loaded/dissolved nutraceuticals (McClements, 2015;

Da Silva et al., 2020; Hu et al., 2020). An emulsion’s physiochemical instability is usually identified as a kinetically unstable system when there is a continuous

20 breakdown over a period because of the destabilization of chemical stability and other physical mechanisms like gravitational separation, coalescence, and flocculation (Silva et al., 2012; Berton-Carabin, and Schroen, 2015). Gravitational separation occurs because of contrasting relative densities between the dispersed and the continuous phases leading to creaming or sedimentation. The other physical mechanisms involve droplet aggregation like flocculation or coalescence and appear to be minimally encountered in nanoemulsions since the droplet particles are relatively small

(Dickinson, 2012; Whitby et al., 2011). Colloidal interactivity relates to droplets’ size and is due to attractive interactions (hydrophobic and van der Waals) and the repulsive interactions (electrostatic and stearic) between adjacent droplets (Dickson, 2012).

2.2.2. Stabilization of O/W emulsion

Several synthetic and natural polymers can be used to create an interfacial hindrance among oil droplets/particles, which can create a thermodynamically stable emulsion, and the polymers are known as emulsifiers. Emulsifiers are required to adsorb on the interface of the oil droplet to generate strong repulsive forces, thus preventing the droplets from coalescing. These repulsive forces can be either electrostatic or steric (Bai et al., 2020a). Various emulsifiers, such as protein isolates, starch, gum arabic, PEG, Tween-80®, glucan, and maltodextrin, have commonly been applied in the food industry for the formation and stabilization of emulsions (Silva et al., 2012; Da Silva et al., 2020). In the past few decades, consumers have been much more informed about food consumption-related matters and how healthy lifestyles can be enhanced. Thus, market demands shift towards healthy and low-calorie foods because of increased public health concerns have captured researchers’ attention in

21 designing and developing nutraceuticals and functional foods (Siro et al., 2008; Sharif and Zahid, 2018). Without a doubt, consumers opt for products that are natively obtained from their natural sources instead of synthetic products (Roman et al., 2017;

Saleh et al., 2019).

2.2.3. Pickering emulsion

Considerable investigations have been carried out on the development of nutraceuticals and functional foods through Pickering emulsions and liposome technology (Siro et al., 2008; Sharif and Zahid, 2018). In the present, much interest has been directed towards utilizing particle-based stabilizers to create and stabilize emulsions. Nevertheless, over a century ago, there was a realization of emulsion stabilization using the small solid particles through the Pickering stabilization process

(Pickering, 1907). These colloidal particles used in Pickering emulsions demand balanced interfacial wettability, hence presenting affinity for both phases.

Consequently, droplets coated with particles tend to exhibit more stability to droplet coalescence than ones coated with molecular-based emulsifiers due to the higher steric repulsion and irreversible adsorption (Vignati et al., 2003). The electrical charge density of these particles depends on the method of preparation, such as acid hydrolysis or application of post-treatment, which increases the electrical density (Rayner et al.,

2014).

Pickering emulsions can be economically suitable for commercial applications.

In general, O/W Pickering emulsions have soundly been applied in various food-related emulsion systems like antibacterial emulsions, coloring and flavoring agents, and delivery of nutraceuticals (Silva et al., 2012; Berton-Carabin, and Schroen, 2015; Khan

22 et al., 2018). Flavors and coloring compounds used in food consist of esters, aldehydes, and ketones acting as functional groups that expose them to oxidative and photolytic degradation. Prevention of such harmful impacts can be done by the encapsulation process of emulsions within these ingredients to enhance the shelf life (Berton-Carabin, and Schroen, 2015). With widespread adoption and application of emulsions in the food, nutraceutical, pharmaceuticals and packaging industries, the sustainability of the emulsification process and its physicochemical suitability for the targeted product, is one of the most important aspect to be addressed (Calabrese et al., 2018; Sarkar and

Dickinson, 2020).

There is an increased interest in investigating particle-based stabilizers in the production of Pickering emulsions and modulate their digestion (Dai et al., 2018; Fu et al., 2019). Research has been done to examine the impacts of naturally derived particle stabilizers, cellulose nanocrystals (CNC), to establish a wide range of applications in food emulsion (Bai et al., 2020a; Bai et al., 2020b). Researchers have investigated the stabilization capacity of CNC for the O/W type emulsions and reported a significant stabilization activity (Bai et al., 2018; Wu et al., 2020). However, the critical element in applications of CNC-stabilized Pickering emulsions involves storage instability. The phase separation of droplet coalescence (gravitational separation) can be of great importance in systems containing large oil droplets and smaller colloid particles (Bai et al., 2020a). Exceptionally, the O/W emulsions containing CNC can also lead to a separation of the oil phase when diluted. This can be due to the following reasons: (a) desorption of the polymer from the oil droplet interface reducing steric or electrostatic repulsion among droplets, (b) weaker gel strength of the aqueous medium, thereby

23 increasing droplets’ interaction, thus coalescence/Ostwald ripening, and (c) reduced surface pressure on the oil droplets due to decreased salt concentration hence depletion/desorption of adsorbed polymers (Wooster, 2006; Maphosa and Jideani,

2018; Bai et al., 2020a).

2.2.4. O/W emulsion and modified CNC

As stated earlier that, bio-based derived nanomaterials are in the process of being investigated for their applications in foods to advance quality and as well to facilitate the functionality due to the abundant primary and secondary hydroxyls. The presence of these hydroxyls on the surface can easily be modified with other polymers to create the desired surface property. Surface modification can also produce other cellulose derivatives or else through grafting to various substances (Tao et al., 2020). Different studies have also indicated that the modification of CNC formed a stable O/W emulsion. The amphiphilic nature is considered as the main driving force for the stabilization of emulsion using the modified CNC. The amphiphilic behavior leads to suggestions that all CNC surfaces consist of dual traits- the hydrophilic and the lipophilic. (Cagnasso et al., 2010; Li and Woodle, 2018). Surface modifications of the

CNC mainly focus on achieving enhanced emulsion stability, where such studies have been focusing on the surface hydroxyl unit modifications (Cagnasso et al., 2010; Li et al., 2020). CNC is known to be an easily- modified substance to produce several derivatives and impart physiochemical characteristics like stiffness, transparency, lower density, and induction of tunable surface chemistry (Saidane et al., 2016;

Natterodt et al., 2017). These developments include the application of modified CNC as a drug carrier, composites, and emulsifier/stabilizer for the Pickering emulsifiers. In

24 essence, the discovery of new and practical approaches in producing stable Pickering emulsions using the unmodified CNC in emulsion systems containing low oil concentration remains the main challenge due to the hydrophilic nature of unmodified

CNC (Cheng et al., 2015). CNC exhibit an abundance state of functional moieties like the aldehyde and the active hydroxyl units, thus, ensuring that they can be modified to enable control over their properties, physical and chemical is essential. Modified CNC can achieve the anisotropy since they can comfortably take advantage of these aldehyde units attached in the ends of the molecules, thus facilitating the selective modification of the CNC (Saidane et al., 2016). Researchers' interest has significantly been directed towards selective modification of CNC to prepare functional CNC with unique features

(Heise et al., 2019; Tao et al., 2020). However, the chemical modification process' sustainability is a big concern; moreover, it is also a complex process, which requires a variety of non-food grade chemicals and reagents.

2.2.5. Liposomes and stability

Liposomes are spherical vesicles that form with the hydration of surfactants like phospholipids after being mixed with water under low shear forces. Ideally, the liposome technique is used to encapsulate and stabilize hydrophilic compounds

(Kopechek et al., 2008; Aditya et al., 2017). Typically, it consists of a bilayer of phospholipids that are self-assembled due to attractive forces associated with the long hydrophobic chains of the fatty acids (Bally et al., 2010). The liposomes prepared through the phospholipid method possess the zwitterionic property (Yadav et al., 2011).

Despite consisting of positively charged choline groups, liposomes prepared with

25 phospholipids show moderate anionic surface property due to the presence of phosphate group (Yoshikawa et al., 2016).

Liposomes enhance the stability of the hydrophilic nutraceuticals during oral delivery; thus, the increased bioaccessibility and bioavailability in the body is achieved

(Couvreur et al., 2001; Eloy et al., 2014). Although liposomes possess few physiochemical limitations such as 1) progressive coalescence due to the presence of

Van-der-Waals attractions leading to the formation of giant liposomes; thus, leakage of the loaded compound (Tan et al., 2016; Haider et al., 2017); and 2) chemical and enzymatic hydrolysis of phospholipid ester of fatty acids of liposome bilayers (Peng et al., 2018; Peng et al., 2019). Developments are made to ensure that physiochemical stability of liposome using polyethylene glycol (PEG), chitosan, and other synthetic and natural polymers (Panja et al., 2019; Sopyan and Gozali, 2020). Previous studies report that a significant improvement in the physicochemical stability of liposomes during storage prepared with chemically modified phospholipids with PEG and cholesterol (Kono et al., 2018; Shen et al., 2018; Zalipsky, 2018). The chemical modification increases the puncture resistance behavior of the phospholipid bilayers by providing a stronger lipophilic-lipophilic interaction between tail groups of fatty acids and the lipophobic-lipophobic interaction between the inner head group (Yadav et al.,

2011; Sopyan et al., 2020). Since the chemical modification of phospholipids limits the application of the liposome system in food due to being extremely expensive, chemical residue from the modification process and feasibility. Thus, researchers have made an effort to explore some alternative greener, cheap and feasible additives to ensure the stability of the liposomes. In recent studies conducted by Lee et al. (2020) have

26 investigated the effect of fortification of stigmasterol (a sterol) on the stability of phospholipid bilayer, found enhanced physicochemical stability of the liposomes, thus enhanced stability of the loaded vitamin C. In another conducted by Panja et al. (2019) reported the enhanced stability of liposome stabilized with casein protein. Utilization of a hydrophilic colloid particle can be a good choice as a stabilizing material since it can easily be adsorbed onto the interface of the liposome due to Van-der-Waals of attraction (Liu et al., 2013).

2.3. Cellulose

Cellulose is the most abundant polymer found on the planet Earth, is the most significant structural component in plants (Kargarzadeh et al., 2018). Cellulose is found in the cell walls of plants, bacterial pellicles, and even in algae. Cellulose is responsible for the rigidity, firmness, and shape of the cells (García et al., 2016). Cellulose was discovered in 1838 by Anselme Payen, a French chemist. Payen isolated cellulose from plant matter and discovered its chemical structure and formula. Cellulose is a homopolysaccharide consists of D-glucose units linked through β-(1 → 4) glycosidic bonds forming linear structures. Cellulose is a complex organic compound chemically identified with the formula (C6H10O5)n thus, classified as polysaccharides, which are linearly aligned into chains and closely packed, causing rigidity. Below is the equation that describes the formation of cellulose (García et al., 2016):

n(C6 H12 O6 ) → (C6 H10 O5)n + nH2O

(Glucose) (Cellulose)

Figure 2.1 illustrates the hierarchical structure of cellulose from wood to its monomeric unit. In their native state, cellulose parallel chains that are closely packed with multiple hydrogen bonds, giving rise to long fibers, also known as microfibrils.

Microfibrils have a width of 4 -10 nm and tend not to dissolve in water and are relatively inert. The microfibrils are characterized by crystalline (highly ordered) and amorphous regions (less ordered) (Kargarzadeh et al., 2018). The estimated degree of polymerization of naturally existing cellulose is approximately 10000 to 15000

Fig. 2.1. Hierarchical structure of cellulose from wood to a monomeric unit.

repeating units of two anhydroglucose rings, so the molecules have a high molecular weight (Moon et al., 2011; Kargarzadeh et al., 2018). Macrofibrils can be formed by further aggregation of microfibrils with the lignin and hemicelluloses present in plant tissue. Due to a high degree of cellulose bonding, microfibrils and macrofibrils produce tall, high tensile strength structures that are generally insoluble in several solvents

(Moon et al., 2011; Kargarzadeh et al., 2018). The cellulose content in both softwoods and hardwoods is approximately 42% (Rowell et al., 2012).

2.3.1. Cellulose nanomaterials

Cellulose nanomaterials are structures, platelet-like particles, or fibers that are modified substances manufactured at nanoscales (Moon et al., 2011; Kargarzadeh et al., 2018). The primary objective for developing these materials is to produce unusual functionalities such as conductivity and increased chemical reactivity compared to similar material not at the nanoscale level (Kargarzadeh et al., 2018; Kasiri and Fathi,

2018). Typically, these nano substances provide some unique advantages such as a larger surface area to volume ratio, compatibility with nanoformulations, homogeneity, better dispersion property, and hydroxyl groups (O-H) as a functional group

(Brinkmann et al., 2016). Cellulose nanomaterials naturally occur with unique structural properties, mechanical and also the optical traits. The surface chemistry of cellulose nanomaterials plays an essential role in determining particle interactions with the environment, dispersion control in solvents, and agglomeration (Brinkmann et al.,

2016). Primarily, Cellulose nanomaterials can be grouped into two; the nanoobjects and nanostructures. Cellulose nanoobjects consist of cellulose nanocrystals (CNC) and cellulose nanofibers (CNFs) (Grishkewich et al., 2017).

There are two major approaches for fragmentation of cellulose source material during its refinement process into nano-sized structures: mechanical shear and acid hydrolysis (Kargarzadeh et al., 2018). The mechanical refinement process involves the application of high shear to tear off the cellulose source material apart. This technique produces the cellulose nanofibrils (CNF) or microfibrillated cellulose (MFC)

29 structures. On the other hand, production of cellulose nanocrystals (CNC) is done by several methods such as acid hydrolysis (Kargarzadeh et al., 2018), acid-alkali hydrolysis (Jordan et al., 2019) enzymatic hydrolysis (Hayashi et al., 2005; Peciulyte et al., 2015), and the metal salt catalyzed hydrolysis process (Kamireddy et al., 2013;

Yahya et al., 2015). The hydrolysis conditions are primarily the determinant of CNC surface chemistry, charge, and as well the aspect ratio, which is a ratio of sizes in different dimensions of CNC (Brinkmann et al., 2016).

2.3.2. Application of Cellulose nanomaterials

Cellulose nanomaterials have potential applications in different areas of food, biomedical, packaging, textile, construction industry due to its aforesaid unique physicochemical properties (Habibi et al., 2010; Grishkewich et al., 2017; Tang et al.,

2017; Amalraj et al., 2019). Food science researchers have investigated the application of cellulose nanomaterials as a fat replacer, thickener, food package, and food gel

(Amalraj et al., 2019; Bai et al., 2020b; Dai et al., 2020). Cellulose nanomaterials also have shown the characteristic of dietary fibers in the rat (Yan et al., 2018).

Microorganisms present in the digestive system’s large intestine can partly degrade cellulose to produce organic acids and gasses (Flint et al., 2012). Thus, as dietary fiber cellulose possesses an ability to absorb moisture inside the intestinal tract, thus easy defecation and prevention of constipation (Ekvall, 2017; Lever et al., 2019).

Technology has led to a new application of cellulose nanomaterials in foods ranged from coatings directly on fruit surfaces to applying as an emulsifier/stabilizer for lipid droplets and fat reducer (Hedjazi and Razavi, 2018; Aswathanarayan and

Vittal, 2019). Recent reviews demonstrate an advantage of Cellulose nanomaterials

30 when applied to fruit pre-harvest to curb skin cracking caused by excess rain, extending the shelf-life, and preserving the nutrients and nutraceuticals (Hubbe et al., 2017; Jung et al., 2020). The reduced environmental toxicities of cellulose nanomaterials, particularly the CNF, may in the future favor food applications (Hedjazi and Razavi,

2018; Aswathanarayan and Vittal, 2019).

Cellulose nanomaterials also have a significant impact on biomedical applications, mainly when used as a binder, due to their biocompatibility, low levels of cytotoxicity, tissue-friendliness, wound healing aspects, antimicrobial effects, and biodegradability (Amalraj et al., 2018; Kargarzadeh et al., 2018). In addition, cellulose nanomaterials have the property of potential binding to tissues through the presence of

O-H groups and negative interfacial charges that promote the cellulose nanomaterial’s electrostatic adsorption onto the tissue. Cellulose nanomaterial’s medical applications are widely studied concerning the delivery of drugs, tissue healing, tissue repair, and tissue replacement cases, and medical implants (Hosseinidoust et al., 2015). Cellulose nanomaterials have primarily been used in the dressing of the wounds due to its anti- infection characteristic and the ability to increase tensile properties associated with the tissue scaffolds. Adding CNC to collagen-based composite raises the levels of its biocompatibility and stabilizes its mechanical functionality. Moreover, a study demonstrated that the addition of CNC to poly [D, L-lactic-co-glycolic acid] (PLGA) produced a more robust composite, enhancing thermal, mechanical stability and facilitated cytocompatibility, spreadability, fibroblast adhesion, and proliferation as compared to neat poly [D, L-lactic-co-glycolic acid] (Mo et al., 2015). CNF is also applicable in the biomedical field by using them for filaments (threads) formation.

Coating the CNF filaments with stem cells can produce sutures that positively influence post-surgery inflammation and wound healing. Hakkarainen et al. (2016) conducted a clinical study investigating CNF’s application in the dressing of wounds, particularly for severely burned patients. There was a successful grafting within the skin donor sites consequent to cellulose dehydration. Cellulose nanomaterials have also been effective in bone generation.

The study has shown that the modification of surface charge on the CNC can be useful in delivering small interfering RNA, which tend to have therapeutic effects.

RNA in their naked form cannot be delivered to the disease cell and can degrade quickly; therefore, it is advisable to employ a nanocarrier such as CNC since CNC is non-toxic, biodegradable, easily available, and a nano-biopolymer (Kargarzadeh et al.,

2018) that facilitates the therapeutic actions through binding RNA (Kim et al., 2020).

A study by Plackett et al. (2014) also reports that CNF and CNC could readily bind with water-soluble drugs when applied in drug delivery functions.

2.3.3. Cellulose nanocrystals (CNC)

CNC is also identified as crystalline nanocellulose or cellulose nanowhisker

(Kargarzadeh et al., 2018), which is a tiny needle or rod-like structure produced from cellulose. Due to CNC’s unique features, researchers have recommended it for potential use in foods, nutraceuticals, drugs, and other applications (Cherhal et al., 2016; Hedjazi

& Razavi, 2018; Bai et al., 2020a; Bai et al., 2020b). CNC derived from wood pulp have a diameter of 5-15 nm with a length of 150-200 nm (Grishkewich et al., 2017;

Tang et al., 2017). CNC accounts for up to 99% of the crystallinity in its structure compared to other cellulose nanomaterials that account for a maximum of 70%

(Abiaziem et al., 2019). CNC is a nano dimension structure with surface hydroxyl groups (O-H) (Natterodt et al., 2017). CNC has excellent dispersibility in aqueous

- media (Cheng et al., 2015), contains a high charge potential due to sulfate ions (-SO3 )

(Prathapan et al., 2016), and is a non-toxic biopolymer (Natterodt et al., 2017;

Kargarzadeh et al., 2018). These characteristics establish CNC as a better platform for developing functional foods (Bai et al., 2018; Dai et al., 2020).

CNC has been considerably utilized in stabilizing O/W emulsion and has shown significant (p≤0.05) effect in retaining physical and morphological attributes of the lipid droplets (Angkuratipakorn et al., 2017; Bai et al., 2018, Bai et al., 2020b; Dai et al., 2020). The stabilization of lipid droplets via CNC is due to the electrical screening behavior of added salt (NaCl). NaCl in the CNC stabilized emulsion improves the rate of adsorption of CNC on oil droplets' interface due to increased surface pressure

(Bergfreund & Fischer, 2019; Bertsch et al., 2019). It has been reported that the addition of salt to the O/W Pickering emulsion also increases the CNC-CNC interactions resulting in increased gel strength of the aqueous phase, which limits the motion and interfacial interaction of the oil droplets, thus stable emulsion (Bai et al.,

2018). However, the stability of these emulsion types can be compromised when diluted or the addition of opposite ions (Wooster, 2006; Maphosa and Jideani, 2018).

Dilution leads to a segregative separation of oil phase due to the following reasons: (a) detachment of the polymer from the oil droplet interface, reducing the steric or electrostatic repulsion; (b) weaker gel strength of the aqueous medium, thus increased droplets interaction consequently coalescence/Ostwald ripening and (c) reduced surface pressure on the oil droplets due to decreased salt concentration resulting in

33 detachment of adsorbed polymer (Wooster, 2006; Maphosa and Jideani, 2018; Bai et al., 2019a).

CNC modified with hydrophobic or amphiphilic molecules have significantly improved the physical stability of the O/W emulsion (Saidane et al., 2016; Natterodt et al., 2017; Li et al., 2020; Tao et al., 2020). The modified CNC helped in creating a structure that is stable to dilution, ionic stress, and pH due to its compatible hydrophobic-hydrophobic interaction between oil and CNC interface (Saidane et al.,

2016; Tang et al., 2018; Le et al., 2020). However, the modification process's sustainability and utilization of several non-food grade chemicals and reagents are a major concern and may preclude their use in food.

CHAPTER 3

STERIC STABILIZATION OF PHYCOBILIPROTEIN LOADED

LIPOSOMES THROUGH POLYETHYLENE GLYCOL ADSORBED

CELLULOSE NANOCRYSTALS AND THEIR IMPACT ON THE

GASTROINTESTINAL TRACT

3.1. Introduction

Recent advances in the nutraceutical, cosmeceutical and pharmaceutical sciences support the concept that bioactive phytochemicals extracted from plants and seaweeds can help in the reduction of some of the human chronic diseases (Kuć, 2007;

Manirafasha et al., 2016).

Phycobiliproteins (PBP) are common light-harvesting protein found in brown/red seaweeds, including dulse and several other marine algae and have potential effects in reducing human diseases such as anti-inflammatory, hepatoprotective, anti-aging and angiotensin (Riss et al., 2007; Furuta et al., 2016). However, PBP is embedded into the cell wall of the seaweed forming a conjugate complex with a broad range of polysaccharides. Due to lack of site-specific enzymes, this protein-polysaccharide conjugate complex limits PBP extractability and digestibility into the human gastrointestinal tract (GIT) and consequently hinders its applications (Mæhre et al.,

2016). Further, the stability of PBP is greatly affected by light, pH, oxidative stress, irradiation, and temperature (Munier et al., 2014). For example, Rastogi et al. (2015) reported that the concentration of PBP extracted from marine cyanobacterium

Phormidium sp. decreased significantly when the extract was exposed to above 40 °C, at a pH below 4.0 & above 8.0, and oxidative stress above 0.8%. Few other studies

35 have investigated the stabilization of extracted PBP with different encapsulating materials, such as pH-regulated multilayered polyelectrolytes and sol-gel glass mediated encapsulation (Chen et al., 1996; Li et al., 2012). However, there is a limited study on the fate and stability of PBP through various pH in human GIT. The stomach stage of GIT system has a most noticeable effect on the quality attributes on the bioactive food compounds due to its very low pH (2.0 to 3.0), however, the mouth and small intestine phases of the GIT have less significant effects (McClements, 2015; Xiao et al. .2017; Winuprasith et al., 2018). To protect the bioactive compounds from lower pH of the GIT and safely deliver into the small intestine and subsequent absorption into epithelial cell, oil-in-water (O/W) based delivery systems have been often tested using liposomes (Dhakane et al., 2017; Woodle et al., 2018; Zaborova et al., 2018).

Liposome-based delivery systems for hydrophilic and hydrophobic compounds improve the solubility and controlled release (Cagnasso et al., 2010; McClements,

2015; Woodle et al., 2018). However, preparation temperature, particle size, and method of preparation cause changes in the arrangement of the phospholipid bilayer within the liposomes. This affects the loosely packed head groups in the inner layer, and tightly packed alkyl chains in the outer layer of the bilayer and leads to physical instability, such as agglomeration or fusion of the liposomes (Kamatsu et al., 2001;

Yadav et al., 2011). Similarly, pH, ionic strength and oxidative effects can hydrolyze the ester linkage between fatty acids & phosphocholine glycerol, cause peroxidation of long-chain fatty acids and consequently lead to the formation of acetaldehyde (Yadav et al., 2011; Guldiken et al., 2018). To reduce these effects, a suitable coating material could be identified and used for encapsulation. A few studies have been investigated to

36 understand the physicochemical interaction between cellulose nanomaterials with liposomes (Kresse et al., 2016; Navon et al., 2017; Lombardo and Thielemans, 2018).

Few researchers have investigated cellulose nanomaterials as a stabilizer for an oil- based delivery system (Angkuratipakorn et al., 2017; Guo et al., 2017; Kasiri and Fathi,

2018; Li et al., 2018). Studies reported that the cellulose nanomaterials stabilize

Pickering emulsion and retain the desired quality attributes. Further, Winuprasith et al.

(2018) investigated the effect of different ratios of CNF (0.1 – 0.7 %) on the stability and digestibility of Pickering emulsion under in vitro simulated GIT conditions and reported that the increase in the concentration of CNFs increased the stability of the emulsion, which reversibly limited the digestibility. Findings concluded that the CNFs could stabilize the oil droplet due to its bulky molecular chain-like structure and large average particle size (>100 nm diameter to several µm in length) while limiting the lipase activity by keeping them away from lipid droplets. (Winuprasith et al., 2018).

Since CNC is tiny, rod-like particles with dimensions of approximately 5 nanometers

(nm) in diameter and 150-200 nm in length (Cheng et al., 2015), it can be a better stabilizer for O/W based delivery system. However, the dispersibility of cellulose nanomaterials into the aqueous medium is limited due to its irreversible agglomeration through surface hydrogen bonding. Adsorption of polyethylene glycol (PEG) via hydrogen bonding onto the surface of CNC could be a suitable way to reduce irreversible agglomerations (Yang et al., 2013; Cheng et al., 2015) through coating the free surface hydroxyl groups (O-H).

Understanding the surface and colloidal properties of CNC and phycobiliprotein, we hypothesize that the cellulose nanocrystals can be adsorbed with polyethylene

37 glycol to stabilize phycobiliprotein for better delivery of the compounds in the human gastrointestinal tract. The objectives of this study were (i) to stabilize cellulose nanocrystals (CNC) by adsorption of different molecular weight of polyethylene glycol

(PEG) and (ii) to understand the physicochemical behavior of phycobiliprotein loaded liposome stabilized through PEG adsorbed CNC during simulated gastrointestinal tract condition (GIT). Subsequent characterization includes particle size, zeta potential, stability PEG adsorbed CNC, the stability of PBPs loaded liposome, hydrolysis of the liposome, and release of PBP.

3.2. Materials and Methods

3.2.1. Chemicals and reagents

Dulse (Palmaria palmata) was purchased from Maine Sea Farm, Franklin, Maine,

United States. Sulfuric acid hydrolyzed cellulose nanocrystals (CNC: contained up to

- 2% sulfate ion (O-SO3 ), 11.5-12.5% solid, 5-20 nm wide and 150-200 nm long) was procured from the Process Development Center, University of Maine. PEG with different molecular weights, 400-4000 g/mol, xylanase (≥2500 units/g), α-amylase

(Aspergillus oryzae) pepsin from porcine gastric mucosa (~2500 units/mg), lipase type

II from porcine pancreas, mucin-type III from the porcine stomach, bile extracts porcine, uric acid ≥99% crystalline, phospholipid (soy lecithin), cellulose membrane dialysis tube (43 mm in diameter), maxi dialysis tube (3 mL capacity) with molecular cut-off 12-14 kDa, HDPE 20 μm porosity bed supports column with a dimension of 2.5 x 170 cm (volume capacity 230 mL) and Sephadex G-100 were purchased from Sigma-

Aldrich, USA. Phospholipid containing 17-24% phosphatidylinositol, 6% phosphatidic

38 acid, 18% phosphatidylethanolamine and 35% phosphatidylcholine and other analytical grades chemical & reagents were purchased from Fisher Scientific Products,

USA.

3.2.2. Purification and characterization of phycobiliproteins

The extraction of PBP was performed following the procedure of Dumey et al.

(2013) with some modifications. Briefly, 20 g of dried dulse was placed into 500 mL of acetate buffer (50 mM @ 5.0 pH) and 0.33 g of xylanase enzyme (16.5g/kg of the sample, ≥2500 units/g) was added. The reaction mixture was stirred at 200 rpm for 3 hours at the incubation temperature of 40°C. After hydrolysis, the reaction mixture was centrifuged 21,800x g for 20 minutes at 4 °C using a super speed centrifuge (Beckman

Coulter Avanti J-E Floor, CA, USA). The supernatant was collected and subsequently dialyzed against distilled water for 12 hours at 4 °C using a 12-14 KDa dialysis tube with intermittent replacement of distilled water to remove the soluble salts, minerals, and carbohydrates. Dialyzed samples were further purified using size exclusion column containing Sephadex G-100 as a stationary bed medium (Fig. 3.1). Purified fraction was freeze-dried (SP Scientific 2000 Virtis 35EL, Freeze Dryer, USA) at -40 °C under a vacuum of 250 millitorrs for three days in the absence of light and the dried PBP was stored in a deep freezer (-20 °C) for further experimental investigations.

Fig. 3.1. Size exclusion-based column separation of phycobiliproteins.

3.2.3. Quantification of phycobiliproteins

Total PBP in the purified extract was analyzed following the method of Rastogi et al. (2015). In brief, 200 µL of purified PBP was loaded in multi plate reader UV-Vis spectrophotometer (BioTek Eon, Vermont, U.S.A) and the absorbance was measured over 250–750 nm range. The equations of Bennett and Bogorad (1973) were used to calculate the amount of PBP:

−1 푃퐶 (푚푔 푚퐿 ) = [퐴615 − 0.474 (퐴652)]/5.34

−1 퐴푃퐶 (푚푔 푚퐿 ) = [퐴652 − 0.208 (퐴615)]/5.09

−1 푃퐸 (푚푔 푚퐿 ) = [퐴562 − 2.41 (푃퐶) − 0.849 (퐴푃퐶)]/9.62

푃퐵푃 (푚푔 푚퐿−1) = 푃퐶 + 퐴푃퐶 + 푃퐸 where PC is phycocyanin, APC is allophycocyanin, PE is phycoerythrin, A is the absorbance at a given wavelength.

3.2.4. Gel electrophoresis analysis of phycobiliproteins

The extracted PBP was analyzed for its molecular analysis using sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Sample was prepared according to the method of Lasekan and Nayak (2016). PBP was diluted with

Laemmli buffer (1:1) containing 5% of 2-mercaptoethanol. Diluted PBP was immediately heated at 100 ºC for 5 minutes. After cooling at room temperature for 20 minutes 7.5 µL sample was loaded into 12% precast gel cassette (Mini-Protean TGX

Precast Gel, Bio-Rad, Hercules, CA., USA). The electrophoretic separation was then performed at 170 V into Mini-Protean Tetra Tank (Bio-Rad, Hercules, CA., USA).

After 10 minutes of the electrophoresis period, the gel cassette was removed from the tank and the cassette was gently removed from the gel. The obtained gel was then washed with distilled water gently followed by staining into 50 mL of GelCode Blue

Stain Reagent (Pierce Biotechnology Inc, Rockford, IL, USA). Macroscopic visuals of the separated protein bands were recorded, and the molecular weights of each band were compared with the marker named Precision Plus Protein Dual Color Standard

(Lasekan and Nayak, 2016).

3.2.5. Adsorption of PEG with CNC

The CNC produced through sulfuric acid hydrolysis contained ~2% of sulfate

- ions (OSO3 ). Studies show that more active sites on the surface of CNC can be created through the desulfation of sulfate ions (Yang et al., 2013 and Cheng et al., 2015). CNC was desulfated to remove the sulfonates from the CNC surface using 0.85 mM NaOH.

Desulfated CNC (coded as DCs) were dialyzed against distilled water for two days and then freeze-dried (SP Scientific 2000 Virtis 35EL, Freeze Dryer, USA) for further PEG adsorption. PEG adsorption onto the desulfated CNC was performed following the procedure of Cheng et al. (2015) with slight modification. Briefly, freeze-dried desulfated CNC (3 g) was placed into a 250 mL Erlenmeyer flask and distilled water was added to make a concentration of 3% DCs. The mixture was stirred at 1200 rpm for 5 minutes followed by ultrasonication at 500 watts for 5 minutes using ultrasonic dismembrator (Fisherbrand, Ultrasonic Liquide Processor, USA). Then, 3.33 g of PEG was added into the flask with continuous stirring at 200 rpm for 3 hours at room temperature. The reaction was subsequently dialyzed for three days against distilled water through a 12-14 KDa molecular weight cutoff dialysis tube. Dialyzed sample was then freeze-dried (SP Scientific 2000 Virtis 35EL, Freeze Dryer, USA) and coded as desulfated PEG adsorbed CNC (DPCs).

3.2.6. Preparation and stabilization of phycobiliproteins loaded liposomes

The PBP was extracted and purified using the method optimized by Dumay et al. (2013). Purified PBP was freeze-dried using a freeze dryer (SP Scientific 2000 Virtis

35EL, Freeze Dryer, USA) and stored in a deep freezer. The liposomes were prepared

42 according to the method described by Gao et al., (2013) with slight adjustment. In brief,

600 mg of purified soy lecithin was dissolved into 200 mL of ethyl acetate and chloroform (60:40) solution. The solution was subsequently passed through a 0.22 µm filter paper to remove undissolved lecithin or any foreign particles, followed by rotary evaporation and thin layer formation in the round bottom flask. The flask with a thin layer of lecithin was placed under a vacuum dryer at room temperature overnight to remove any traces of solvent. To encapsulate the purified PBP, the thin layer was further rehydrated with PBP solution (300 mg/50 mL) followed by the addition of 100 mL of phosphate buffer saline (7.4 pH). The resulting multilamellar liposomes were sonicated for 1.2 minutes at 40% amplitude (at the interval of 15 seconds ON and 5 seconds OFF-pulse time) using an ultrasonic dismembrator (Fisherbrand, Ultrasonic

Liquid Processor, USA) to obtain unilamellar vesicle (SUV) liposomes. This liposome was coded as phycobiliprotein loaded liposome (PL). Loaded liposome was subsequently stabilized by the addition of 100 mL suspension/solution (1% w/v) of desulfated cellulose nanocrystals (DCs), or polyethylene glycol (PEG-400-4000 g/mol) or polyethylene glycol adsorbed onto cellulose nanocrystals (DCs-2000 or DCs-4000).

These stabilized liposomes are coded as PLDCs, PLP-2000, PLP-4000, PLDCs-2000 and PLDCs-4000. Stabilized liposomes were dialyzed in the absence of illumination over 48 hours at 4 ℃ using 50 kDa cellulose dialysis tubing against distilled water to remove the excess PBP.

3.2.7. In vitro gastrointestinal tract (GIT) study

The stability of liposome and PBP during GIT study was investigated following an in vitro GIT model proposed by Winuprasith et al. (2018) with slight adjustments. The

43 simulated digestion media were prepared for the in vitro study are given in Table 3.1.

Briefly, 100 mL volumetric flask (quantity 24) was washed properly with distilled water, dried under vacuum and wrapped with an aluminum sheet to avoid light degradation of PBP during the study. Initially, 40 mL of PBP loaded stabilized liposomes was preheated in a shaking water bath (JULABO, SW22, Julabo Inc., USA) at 37 °C for 2 minutes with a shaking speed of 100 rpm and was considered as initial sample of the GIT study. After minutes, 20 mL aliquot was transferred into the next flask for further GIT phase (mouth phase - SSJ) study. SSJ (20 mL) added to the reaction mixture previously, was incubated at 37 °C and adjusted to a pH of 6.8 using

0.5M NaOH/HCl solution. The reaction was allowed to occur for 5 minutes with continuous stirring at 100 rpm at 37 °C. The gastric phase investigation was performed by transferring 20 mL of SSJ phase sample into the next 100 mL volumetric flask.

Twenty mL of gastric juice (SGJ) was added into a flask and pH was maintained at 2.0 using 0.5M HCl. The mixture was then incubated for 2 hours at 37 °C with continuous stirring at 100 rpm. After completion of the SGJ phase, pH was neutralized using 0.5

M NaOH solution. The small-intestinal (SIJ) phase was performed by transferring 20 mL aliquot of the SGJ phase sample into the next 100 mL volumetric flask and adding

1.5 mL SIJ (CaCl2+NaCl solution) followed by 3.5 mL of bile salt and 2.5 mL of lipase solution. The reaction mixture was maintained at pH 7.0 and incubated at 37 °C with stirring at 100 rpm for 2 hours of the reaction period.

The sample fractions of each GIT phases were collected separately and stored under refrigeration condition until qualitative and quantitative characterizations.

However, particle size and the zeta-potential study were performed immediately.

Table 3.1. Compositions of chemicals for preparation of simulated gastrointestinal tract (GIT) medium for in vitro study.

Chemical Quantity Vol. (mL) pH name

Na2HPO4 10.11 g Initial 500 6.8 ± 0.2 NaH2PO4 1.70 g KCl 4.48 g KSCN 1.00 g

NaH2PO4 4.44 g

NaSO4 2.87 g Simulated saliva NaCl 8.77 g juice (SSJ): 500 6.8 ± 0.2 NaHCO3 8.47 g Mouth phase Urea 1.00 g Uric acid 75 mg Mucin 25 mg α-amylase 290 mg Simulated gastric HCl 3.5 mL juice (SGJ): NaCl 1.00 g 500 2.0 ± 0.2 Stomach phase Pepsin 1.60 g

Simulated CaCl2 1.39 g 50 intestinal juice NaCl 10.95 g 7.0 ± 0.2 (SIJ): Small Bile salt 250 mg 50 intestine phase Lipase 80 mg 50

Note: All prepared chemicals and reagents were prepared freshly and incubated at 37 °C prior in vitro study.

3.2.8. Determination of particle size and zeta potential

The samples collected from different GIT phases were further diluted using a phosphate buffer (6.8 pH) to standardize for CNC-loaded PBP concentration. Particle size and ζ-potential were performed on a zeta-sizer (Zetasizer Malvern 3000 HSA).

Means and standard deviations of ten automatic replications (by instrument) of each sample (triplicate of each sample) were recorded. Similarly, the particle size and ζ- potential of fresh DPCs sample were performed by dispersing 0.2 g of sample into 10 mL of distilled water by stirring at 100 rpm for 5 minutes.

3.2.9. Spectrofluorometer analysis of phycobiliproteins

Effects of the GIT phase on the stability of PBP, particularly fluorescence intensity was recorded using a spectrofluorometer (Jobin Yvon Fluorolog-3-21). 5 mL of samples from the GIT phases were mixed with 500 µL of a 100 mM Tween-20 solution and were vortexed for the 2 minutes followed by centrifugation at 4,550x g for

10 minutes on a rotor centrifuge (Beckman Coulter Avanti J-E Floor, CA, USA). The fluorescence intensity of the collected supernatant was measured using a spectrofluorometer (Jobin Yvon Fluorolog-3-21). Excitation and emission spectra of

545 nm 575 nm of the PBP were used to record the fluorescence intensity (Lage-Yusty, et al., 2013; Caracciolo, 2015).

3.2.10. FTIR analysis

Infrared spectroscopic analysis of the samples was recorded on an FTIR spectrometer (ABB FTLA 2000). The samples (fresh and GIT) were placed onto the top of a crystal diamond and spectra were recorded between the wavelengths of interest

46 ranging from 400 to 4000 cm-1. The results were recorded in percent transmittance

(%T). Obtained FTIR %T was then converted to the absorbance following the equation developed by Toft et al. (1993).

퐴푏푠표푟푏푎푛푐푒 = 2 − 푙표푔 (%푇) The adsorption of PEG onto the desulfated CNC surface was confirmed by exposing freeze-dried desulfated CNC and DPCs samples through this analysis. Similarly, PBP loaded liposome, liposomes stabilized by desulfated CNC and DPCs samples were directly exposed for FTIR analysis. An additional FTIR analysis was performed to understand the desorption behavior of PEG from the surface of CNC under the SGJ phase. The samples collected from the SGJ phase were dialyzed through a 3 mL maxi dialysis tube for 24 hours to remove the desorbed PEG, followed by freeze-drying (SP

Scientific 2000 Virtis 35EL, Freeze Dryer, USA). However, to study the liposomes digestion during the GIT phase, only SIJ phase samples were exposed for FTIR analysis

(to find a trend in the total digestion throughout the GIT phase). To extract undigested liposomes and free fatty acids, 5 mL of samples collected from the SIJ phase were mixed with 2 mL of ethyl acetate: chloroform solution (60:40). Samples were vortexed for 2 min and allowed for phase separation. After a clear phase separation of the upper ethyl acetate, the chloroform layer was collected gently with a 5 mL syringe. The lower portion was again subjected to the same solvent separation process. Collected upper layer samples were pooled and evaporated under vacuum and investigated for FTIR characterization.

3.2.11. Confocal laser scanning microscopy

The stability and coalescence behavior of stabilized liposomes under the GIT phase was performed on a Leica TCS SP laser scanning confocal microscope (LSCM).

50 µL of samples were loaded on a microscope slide and gently covered with a coverslip. A fluorescence characteristic of PBP (phycoerythrin) excitation at 545 nm and emission at 575 nm was used. A 100X magnification lens was used to acquire

LSCM results with a specimen scanning depth of 1.5 µm.

3.2.12. Statistical analysis

The experiments for the particle size, zeta potential, and PBP release were performed in triplicate. One-way ANOVA was applied to determine the significant effect at the level of p≤0.05, while the Tukey’s honestly significant difference (HSD) post hoc test was performed to identify the significant differences between the means of the characteristic parameters at different GIT phases.

3.3. Results and Discussion

3.3.1. Phycobiliproteins content in the extract

The macroscopic image of the purified PBP is given in Fig. 3.2. PBP in the extract accounted for 1.36 mg/mL for the extract without enzymatic hydrolysis; however,

14.99 mg/mL was obtained for the extract assisted with xylanases enzymatic hydrolysis. The major PBP in the extract was phycoerythrin (PE) followed by allophycocyanin (APC) and phycocyanin (PC) (Table 3.2).

Table 3.2. PBP contents (mg/mL) of Palmaria palmata (Dulse)

PE PC APC Total PBP

Fresh 0.74±0.07 0.30±0.02 0.32±0.02 1.36

Hydrolyzed 9.55±0.35 2.19±0.07 3.25±0.15 14.99

Note: Results are the mean ± standard deviation of triplicate for each sample (n=3). PE: phycoerythrin, PC: phycocyanin and APC: allophycocyanin.

Fig. 3.2. Macroscopic images of purified PBP (A) and freeze-dried PBP (B).

A B

3.3.2. Gel electrophoresis of purified phycobiliproteins

SDS-PAGE analysis of extracted PBP from the dulse revealed that there three bands, which comprised at 16-18 kDa of alpha and beta subunit of phycoerythrin and at 34 kDa of the gamma subunit of phycoerythrin. Furuta et al. (2016) reported that the molecular weight of the phycoerythrin extracted from the dulse molecular weight around 20 kDa (Fig. 3.3).

Fig 3.3. SDS-PAGE of phycobiliproteins. Molecular weight marker is in far left.

3.3.3. Adsorption of PEG with CNC

A schematic diagram of adsorption of polyethylene glycol (PEG) onto the desulfated CNC (DCs) surface with an O-H bond is given in Fig. 3.4a. From the FTIR spectra in the range of 2800 to 2900 cm-1, the C-H stretching of the alkane group of

PEG showed the most noticeable differences between DCs and PEG adsorbed DCs

(DPCs). DPCs with a higher molecular weight of PEG had higher C-H stretching vibration, which could be due to the long-chain length of the higher molecular weight of PEG. Similar to the C-H stretching vibration of the alkane group, characteristic peaks of PEG adsorption were C-H scissoring and bending vibration of alkane between 1290 to 1440 cm-1, C-O stretching vibration of an alcohol group at 1245 cm-1, C-O-C peak of ether group between 1190 to 987 cm-1 and a characteristic peak of PEG crystallinity at 834 cm-1 (Sun, Zhang, & Chu, 2008). These characteristics peak of the PEG in the

DPCs sample confirmed the successful adsorption of PEG on the surface of DCs (Fig.

3.4b).

Fig. 3.4. Schematic of polyethylene glycol adsorption (a) and FTIR spectra (b).

a b

Note: Desulfated CNC (DCs) Polyethylene glycol (PEG) of different molecular weight viz., 400, 600, 1000, 1500 and 4000 g/mol, PED adsorbed desulfated CNC (DPCs)

3.3.4. Effect of PEG’s molecular weight on average particle size and zeta potential of desulfated cellulose nanocrystals (DCs)

The particle sizes of DCs and PEG adsorbed DCs (DPCs) samples were analyzed to understand the effect of different molecular weights of PEG adsorption on the stability of DCs in an aqueous medium. An average particle size (APS) and zeta potential of DCs and DPCs are given in Fig. 3.5. Purchased CNC contained an initial average particle size of 150-200 nm; however, DCs displayed an average particle size of 6922.3 nm. The results implied that the desulfation might reduce the electrostatic repulsion among CNC by removing the negatively charged sulfate ions (Yang et al.,

2013). The desulfation may lead to the formation of CNC intermolecular O-H bridge causing CNC to agglomerate (Habibi, Lucia, & Rojas, 2010 and Cheng et al., 2015).

Adsorption of PEG had an effect on the re-dispersion of DCs as well. The average

Fig. 3.5. The average particle size of PEG-modified CNC

800 a a 700 600 500 400 300 b b c c,d c

Particle size Particle (nm) 200 100 0 DCs*10 DPCs-400 DPCs-600 DPCs-1000 DPCs-1500 DPCs-4000 Sample type

Sample type b DCs DPCs-400 DPCs-600 DPCs-1000 DPCs-1500 DPCs-4000 0

-5

-10

-15 c

-20 b b Zeta (mV) potential Zeta a a -25 a

-30

Note: Desulfated CNC (DCs) and PEG (molecular weight 400, 600, 1000, 1500 and 4000 g/mol) adsorbed DCs (DPCs) (a). Zeta potential (b). DCs*10 represents the particle size (nm) is 10 times higher for DCs sample. Different letters above columns indicate significant differences (Tukey’s HSD, p≤0.05).

52 particle size of DPCs samples was much lower (≤ 244.64 nm) than the DCs (6922.3 nm). From our observation, the adsorption of PEG on the surface of DCs might interfere with intermolecular attraction among DCs (Yadav et al., 2011; Guo, Xie, &

Luo, 2013). This adsorption of PEG onto DC’s may create a steric hindrance causing significant redispersion of desulfated CNC (Cheng et al., 2015). It was also observed that the adsorption of polyethylene glycol at the higher MW decrease in the average particle size ranging from 244.64 nm to 198.53 nm. This trend could be due to the longer chain length of higher MW PEG, which may result in more surface steric hindrances and subsequently limit the formation of O-H bond during the drying process

(Cheng et al., 2015). Desulfated CNC had a total surface charge of -21.74 mV, which implies that the surface charge was adequate to create a temporary electrostatic stabilization (Clogston and Patri, 2011).

However, results from the particle size depicted that the particles were not stable. It is assumed that the agglomeration might be due to the higher forces of attractions between surface O-H groups as compared to electrostatic repulsive force

(Yadav et al., 2011 and Guo et al., 2013). Adsorption of lower MW PEG (400 and 600 g/mol) resulted in an insignificant (p>0.05) change in the zeta potential value.

However, desulfated CNC adsorbed with higher MW of PEG (1000, 1500 and 4000 g/mol) decreased the zeta potential value slightly, which might be attributed to the surface coating of desulfated CNC by a long chain of higher MW of polyethylene glycol (Chibowski & Paszkiewicz, 1999). Despite the slight decrease in zeta potential from -21.74 to -15.44 mV with an increase in the MW of PEG, stabilization of CNC

53 was noticed. The stability of CNC implied that the particles were stabilized through steric hindrance rather than electrostatic stabilization in the aqueous medium.

3.3.5. Effect of gastrointestinal tract (GIT) phases on quality characteristics of phycobiliprotein (PBP) loaded liposomes

Liposomes are commonly used in pharmaceutical, cosmeceutical and nutraceutical industries as loading materials/carriers for hydrophilic and hydrophobic bioactive compounds. However, its physical and chemical stability under different environmental conditions is a concern. Various studies have been conducted so far to stabilize the liposomes (Parise et al., 2015; Mahmud, Piwoni, Filiczak, Janicka, &

Gubernator, 2016; Jang et al., 2018 and Raja et al., 2018). PBPs loaded small unilamellar vesicle liposome contains an average particle size of 319±78.61 nm and average zeta potential of -31±8.82 mV. To understand the physicochemical stability of liposomes, we investigated the agglomeration behavior and enzymatic digestibility of

PBP loaded liposome under the GIT phase, the effect of the GIT phase on the particle size, zeta potential, and liposomes digestion and release behavior of PBP’s were investigated and illustrated in the following sub-headings.

3.3.5.1. Effect on average particle size

The average particle size (APS) of the samples was investigated under different

GIT phases. PBP loaded liposomes stabilized with Desulfated CNC and DPCs into phosphate buffer (pH 6.8) was considered as an initial phase. From Fig. 3.6, it was observed that the sample stabilized with desulfated CNC had a much higher value of the average particle size (5392.6 nm) as compared to the samples stabilized with

Fig. 3.6. Effect of the GIT phases on the average particle size of samples.

800 DCs*10 DPCs-400 DPCs-600 Aa DPCs-1000 DPCs-1500 DPCs-4000 700 Ab Ab Ab Ab 600 Bb Ab BbBb CcCc 500 Cc CbCb 400 Cb Ba Ba BaBa 300 Ba Ca Ca Ba Ca Ca BaBaBa Ca Ca

200 Particle size Particle (nm) 100

0 PEG-DCs Initial SSJ SGJ*10 SIJ*10 Treatments Note: Desulfated CNC (DCs), PEG adsorbed DCs (DPCs). DCs*10 = values are ten times higher for DCs sample under however, SGJ*10 and SIJ*10 represent the values are ten times higher for all samples in gastric (SGJ) and intestinal phase (SIJ). The upper-case letters denote the significant difference (p ≤0.05) among sample types; however, small letters denote significant differences (p≤0.05) among gastrointestinal phases. different DPCs (DPCs 400 – 4000: 308.72 to 207.33 nm). Two possible reasons were presumed for these findings: first, it could be due to the poor re-dispersible and agglomeration of desulfated CNC as described in section 3.2, and second, the coalescence of liposomes due to the absence of steric hindrance among intermolecular of liposomes (Yadav et al., 2011). The confocal laser scanning microscopy (CLSM) images of the initial samples (Fig. 3.7) showed that liposomes were stabilized by different DPCs, which supports the earlier statement that the DPCs may create the steric hindrance among liposomes and thereby stabilize liposomes. The effect of simulated saliva juice (SSJ) phase onto the average particle size of samples was insignificant (p

> 0.05), except the sample stabilized through desulfated CNC. It was noticed that the average particle size of the desulfated CNC stabilized sample increased from 5392.6 to

6042.5 nm after exposure to SSJ phase (Fig. 3.6). It was inferred that as time passed, due to no stabilization effect (DCs was precipitated), the continuous coalescence of liposomes occurred with the progression of the GIT phase that causes an increase in the average particle size. The confocal microscopic images also confirmed an increase in the liposome’s particle size (Fig. 3.7), which may be due to a continuous coalescence of liposomes by the Van-der-Waals forces of attraction (Yadav et al., 2011 and Guo et al., 2013). The confocal microscopic images of the samples stabilized through DPCs showed good stability of liposomes under the saliva phase (SSJ) of the GIT. It was presumed that the stability of liposomes might be due to the steric behavior of DPCs, preventing the coalescence of liposomes (Yadav et al., 2011). It was observed that the average particle size of the sample increased up to 10 times higher than its SSJ phase except the sample stabilized through DCs (Fig. 3.6). The average particle size of desulfated CNC sample was increased slightly from 6042.5 to 6193.5 nm, which may be due to the further coalescence of liposomes during the gastric phase of GIT.

However, the average particle size of DPCs samples increased to a maximum value of

5526 nm for DPCs-400 followed by 5487 nm (DPCs-600), 4308 nm (DPCs-1000),

4182 nm (DPCs-1500) and 3718 nm (DPCs-4000). These findings could have happened due to the following reasons. Firstly, the instability of O-H bond between desulfated CNC and polyethylene glycol at lower pH (Habibi et al., 2010; Way et al.,

2012 and Yang et al., 2013) causing desorption of polyethylene glycol followed by

Fig. 3.7. Confocal laser scanning microscopic images of stabilized liposomes.

Note: Images are the effect of gastrointestinal tract (GIT) phase on the physical stability of stabilized liposomes, where desulfated CNC is DCs, PEG (400-4000 g/mol) adsorbed DCs is DPCs. Scale bars for the size of the liposomes are 1.0 µm. SSJ – simulated saliva juice, SGJ – simulated gastric juice, SIJ – simulated small-intestinal juice.

57 agglomeration of desulfated CNC. Secondly, the coalescence of liposomes is due to its physical instability (Yadav et al., 2011).

Besides, the confocal microscopic images also support the findings for an increase in average particle size (Fig. 3.7). The FTIR analysis of the gastric juice (SGJ) samples confirmed the desorption of PEG from the surface of the desulfated CNC. As it can be seen from Fig. 3.8 (FTIR spectra) the characteristics peak of the PEG (C-H stretching vibration alkane between 2800 to 2900 cm-1; C-H scissoring and bending of alkanes between 1290 to 1440 cm-1, C-O stretching of an alcohol group at 1245 of, C-

O-C peak of ether group between 1120 to 1010 cm-1 and a characteristic peak of crystallinity at 834 cm-1) of higher molecular weight of polyethylene glycols have disappeared. It was observed that the average particle size of lower molecular weight of polyethylene glycol’s samples (DPCs-400 and DPCs-600) increased more significantly (p≤0.05) than the DPCs-1000, DPCs-1500, and DPCs-4000, which was confirmed due to the anti-coalescence effect of longer chain length of higher MW PEG remaining in solution (Fig. 3.7). The results were in agreement with the finding of

Allen, Hansen, Martin, Redemann, and Yau- Young (1991), who observed that the liposomes stabilized under the higher molecular weight of polyethylene glycol from

1900 to 5000 g/mol were more stable than the liposomes stabilized under the lower molecular weight of polyethylene glycol 120 to 750 g/mol. A similar observation of the long chain of polyethylene glycol due to the entangling effect on the stability of liposomes was also reported by Hashizaki et al. (2003).

Fig. 3.8. FTIR spectra of CNC before and after the gastric phase of gastrointestinal tract.

a b

Note: PEG (molecular weights 400, 600, 1000, 1500 and 4000 g/mol) adsorbed desulfated CNC (DPCs) before (a) and after gastric (SGJ) phase (b). Highlighted spectra depict the adsorption of PEG adsorbed with DCs.

After exposing the samples into the next GIT phase, the small-intestine phase

(SIJ), the average particle size of the samples significantly increased with respect to

SGJ phase results. It was hypothesized that the increase might be due to a reduction in the number of lower particle size liposomes via lipase hydrolysis (Cheng et al., 2017;

Winuprasith et al., 2018). Similar to the SGJ phase, the desulfated CNC stabilized sample had larger average particle size compared to DPCs stabilized samples. Confocal microscopic images showed the cause of the increase in overall particle size with respect to the SGJ phase. It can be seen from Fig. 3.7 that the desulfated CNC samples have bigger liposomes that may have significantly contributed to the larger particle size. However, DPCs samples with a higher molecular weight of polyethylene glycol

59 even after desorption showed better stabilization of liposomes as compared to the DPCs with lower MW of PEG, thus the average particle size compared to desulfated CNC.

3.3.5.2. Effect on zeta potential

The effect of an electrostatic behavior of the samples throughout the GIT phases was recorded in the form of average zeta potential (AZP) (Fig. 3.9). It was also confirmed that the desulfated CNC sample showed coalescence of liposomes even at the initial phase (Fig. 3.7), which may have resulted in an increase in the size of the liposomes and subsequently lowering the surface area and zeta potential (Sahu et al.,

2020). Saliva phase of GIT had a slight effect on the average zeta potential value of the samples (Fig. 3.9), which could be due to the cationic (K+, P+, and Na+) and anionic

(mucin) compounds of saliva fluid (Pfeiffe et al., 2014 and Menchicchi et al., 2015) influencing the anionic behavior of liposomes and desulfated CNC (Wang & Wang,

2008). A decrease in the zeta potential of vitamin D3 loaded emulsion due to electrostatic screening effect caused by counter ions of the simulated saliva juice and mucin after exposure in the mouth phase of GIT was reported by Winuprasith et al.

(2018). From Fig. 3.9, it was observed that the average zeta potential of the samples increased after exposure to gastric juice (pH 2.0). An overall increase in the average zeta potential that might be due to a strong anionic characteristic of phospholipid at a lower pH (Pfeiffe et al., 2014). In contrast to the effect of GIT on the average particle size. An increase in the molecular weight of polyethylene glycol had an increase in the average zeta potential value. DPCs-400 and DPCs-600 displayed much lower average particle size value, ranging from -38.2 to -40.64 mV compared to DPCs-1000 (-48.4 mV), DPCs-1500 (-53.63 mV) and DPCs-4000 (-64.38 mV). Following the

Fig. 3.9. Effect of gastrointestinal (GIT) phase on zeta-potential of samples.

Simulated GIT phase PEG-DCs Initial SSJ SGJ SIJ 0 -10

Ab -20 Aa Aa Aa Aa Aa -30 Ba Ba Ba BbBb BbBbBb Ba BbBb -40 BbBbBbBbBb CbCb Cc Cc -50 Dc

Zeta (mV) potential Zeta Ed -60 Dd

-70 Fe DCs DPCs-400 DPCs-600 DPCs-1000 DPCs-1500 DPCs-4000 Note: PBPs is phycobiiprotein, DCs is desulfated CNC, DPCs is PEG (molecular weight 400, 600, 1000, 1500 and 4000 g/mol), SSJ is simulated saliva juice, SGJ is simulated gastric juice, SIJ is simulated small-intestinal juice. The upper-case letters denote the significant difference (p ≤ 0.05) among sample types; however, small letters denote the significant differences (p≤ 0.05) among gastrointestinal phases. antagonistic effect of SGJ phase on the average particle size (higher molecular weight of PEG resulted in a decrease in particle size), it was inferred that lower average particle size might have resulted in a higher surface charge. DPCs-4000 had the most significant increase in the average zeta potential value (p≤0.05), which could be due to the entangling effect of long-chain length of PEG-4000 that sterically hindered the coalescence of liposomes after desorption fromdesulfated CNC (Fig. 3.7) (Allen et al.,

1991; Hashizaki et al., 2003). The desorption effect of the gastric phase (SGJ) also exposed the desulfated CNC surface charge that might have contributed to increases in the zeta potential (Cheng et al., 2015). The electrostatic behavior of samples under the simulated intestinal phase (SIJ) displayed similar effects as resulted in the SGJ phase.

3.3.5.3. Stability of phycobiliproteins

The fluorescence intensity of the encapsulated PBP is given in Fig. 3.10. Results from the figure depict the significant effect of the GIT phases on the fluorescence intensity of the PBP. The most noticeable results were observed at the gastric phase, where the liposomes stabilized with modified CNC (DPCs) had higher intensity as compared to the liposomes with unmodified CNC (DCs). It was also noticed that the liposomes stabilized by the modified CNC with higher molecular weights of PEG (DPCs-4000) had higher fluorescence intensity, which was assumed due the physical stability of the liposomes. The physically stable liposomes may have avoided the formation of giant liposomes, thus less leakage of the loaded PBP and degradation, hence higher the fluorescence intensity.

Fig. 3.10. Stability of PBP during gastrointestinal tract phases.

45 40 35 30 DCs 25 DPCs-400 DPCs-600 20 DPCs-1000 15 DPCs-1500

10 DPCs-4000 Fluorescence intesnsity(CPS) Fluorescence 5 Initial SSJ SGJ SIJ Sample types Note: Fluorescence intensity of PBP during gastrointestinal (GIT) phase. SSJ - saliva juice, SGJ - gastric juice, SIJ - small-intestinal juice. De-sulfated CNC (DCs) and PEG (400-4000 g/mol) adsorbed DCs (DPCs).

3.4. Conclusion

The simulated gastrointestinal tract (GIT) phase significantly affected the stability of phycobiliprotein (PBP)-loaded liposomes stabilized through polyethylene glycol (PEG) adsorbed cellulose nanocrystals (CNC). Before the GIT experiments, an effort was made to re-disperse the freeze-dried desulfated cellulose nanocrystals (DCs) into the aqueous medium through adsorption of different MW of PEG viz., 400, 600,

1000, 1500 and 4000 (coded as DPCs). The effect of the same GIT phase on the characteristics of liposomes stabilized through desulfated CNC displayed an effective coalescence of liposomes. Lower pH of the gastric phase caused effective desorption of PEG from the surface of desulfated CNC, which resulted in precipitation of desulfated CNC, thereby causing coalescence of liposomes. In summary, we concluded that the adsorption of PEG on the desulfated CNC surface was appropriate to stabilize

PBP loaded liposomes initially and in the saliva environment. However, the desorbed gastric condition led to an effective coalescence of PBPs loaded liposomes. It was inferred from the investigation that further modification of CNC is needed for the stabilization of phycobiliprotein during encapsulation.

CHAPTER 4

IMPROVED STABILITY OF PHYCOBILIPROTEIN WITHIN LIPOSOMES

STABILIZED BY POLYETHYLENE GLYCOL ADSORBED CELLULOSE

NANOCRYSTALS

4.1. Introduction

Phycobiliproteins (PBPs) are a class of proteins found in different macroalgae

(seaweed) microalgae and cyanobacteria. These proteins are colored and fluorescence, and are subcategorized into three major groups of phycoerythrin, phycocyanin and allophycocyanin. PBP's have been commercially distinguished as pharmaceuticals and nutraceuticals for their potential application as anti-cancer, anti-cardiac, anti-aging, anti-inflammation, neuroprotective and hepatoprotective agents in the food industry

(Riss et al., 2007; Futura et al., 2016; Lee et al., 2017). In seaweed, PBP is embedded in the cell walls and bound to polysaccharides. Due to the conjugation with polysaccharides, the extraction of PBP from seaweed is difficult. Moreover, the lack of enzymes in the human digestion system for the specific type of polysaccharides in seaweed, microalgae, and cyanobacteria also limits their digestibility, thereby limiting the bioavailability of PBP (Dumey et al., 2013; Maehre et al., 2016; Martin).

Furthermore, the physicochemical stability of PBPs is strongly influenced by several stress factors such as temperature, illumination, oxidative stress and acidic/basic conditions (Munier et al., 2014; Rastogi et al., 2015). Studies show that

PBP's are unstable when exposed to temperatures above 40 °C, pH below 4.0 and above

8.0, resulting in the loss of 50 to 60% of phycobiliprotein (Rastogi et al., 2015).

Research also shows that exposure to the illumination, even at refrigeration at 4 °C for

64 a day, can significantly reduce the PBP content by 60%. On the other hand, exposure to oxidative stress has comparatively less effect on PBP content, reducing to the amount from 25 to 60% depending upon types of PBP (Liu et al., 2019; Munier et al., 2014). A few studies have been conducted on encapsulating the PBP for control release and targeted delivery (Chen et al., 1996; Hu et al., 2008; Manconia et al., 2009; Castangia et al., 2016; Suzery et al., 2017). However, no studies have been reported regarding the stabilization of PBP under the effects of different food processing and storage parameters such as temperature, pH, illumination and oxidative stress. Li et al. (2012) reported improved stability by incorporating the PBP within a polyelectrolyte multilayered microcapsule. The study concluded that the microcapsule, however, a further increase or decrease in the pH adversely affected the stability and encapsulation efficacy. Yan et al. (2014) microencapsulated (1–3 mm capsule size) phycoerythrin within alginate and chitosan as a carrier material using an extrusion process. Results showed that a combination of alginate and chitosan (2.5 and 2.0%) significantly retained the phycoerythrin content of up to 93% at 50 °C and up to 88% under illumination for 40 days in storage. The ability to obtain good stability through the effective distribution and dispersion of encapsulated bioactive compounds into different food matrices, targeted delivery and control release into the human digestion has resulted in major attention (Espitia et al., 2019; Guo et al., 2019)

Due to advancements in the science and technology of nanoencapsulation, this has become a promising area of research to enhance the encapsulation efficiency, targeted delivery and control release of encapsulated compounds (Aditya et al., 2017; Bai et al.,

2019b; Bao et al., 2019; Fu et al., 2019). Various nanoencapsulation techniques such

65 as molecular complex, ionic gelation, multiple emulsion, multiple layered emulsion, biopolymer complex colloid systems and solid fat particle have been tested

(McClements, 2015; Dhakane et al., 2017). These encapsulation techniques have several disadvantages, such as leakage of loaded materials, instability during ionic reflux, agglomeration and precipitation at lower pH, limited loading capacity, less bioavailability and production and handling cost.

Liposome encapsulation system, which provides higher loading capacity, superior bioavailability and less production cost, offers a multifaceted application for encapsulation of bioactive and unstable compounds (Couvreur et al., 2001;

McClements, 2015; Eloy et al., 2014). However, liposomes have a few physicochemical limitations of 1) progressive coalescence due to the Van-der-Waals attraction, which leads to the formation of giant liposomes, thereby leading to the leakage of the loaded compound (Taylor et al., 2003; Laye et al., 2008; Haider et al.,

2017; Tan et al., 2016) and 2) hydrolysis of liposome structure, that depends upon the composition of the phospholipid used in liposome preparation (Rasti et al., 2012;

Vafabakhsh et al., 2013; Peng et al., 2018; Peng et al., 2019). Stabilization of liposome using polyethylene glycol (PEG), chitosan and other polymers either covalently or noncovalently bound to the liposome have been reported, as they greatly improve the physicochemical stability of the liposome during storage (Laye et al., 2008; Tan et al.,

2016; Kono et al., 2018; Shen et al., 2018; Zalipsky, 2018).

Nano biopolymers for the liposomes in encapsulation technology have attracted many researcher's attention. Among them, cellulose nanocrystals (CNC) have been investigated for the stabilization of drug-loaded liposomes due to its unique properties

66 such as their dimension (150–200 nm long and 5–15 nm wide) and non-toxicity

(Roman et al., 2009). As mentioned in chapter 3, it was found that the adsorption of polyethylene glycol (PEG) on the surface of desulfated CNC (zeta potential −25 mV) helped in the stabilization of phycobiliprotein loaded liposome at neutral pH.

Commercially produced CNC via sulfuric acid hydrolysis has a significant amount of sulfate anions with a zeta potential of approximately −55 mV (Cheng et al., 2015).

Anionic property of the CNC and liposomes, anionic at neutral pH, creates a strong electrostatic repulsion limiting the adsorption of CNC on the surface of liposomes

(Navon et al., 2017). Although a few researchers have reported the adsorption of similar charged polymers and liposomes (Seki and Tirrell, 1984 and Liu et al., 2013), a reduced electrostatic repulsion can also assist the adsorption of CNC on the surface of liposome

(Navon et al., 2017).

As reported in chapter 3, it was noticed that a reduced electric property of the

CNC-liposome system can cause the agglomeration or precipitation to the CNC and then liposome. Therefore, a modification to the CNC was suggested. In addition to surface modification, understanding of the effects of different environmental stressors such as temperature, pH and illumination on the stability of liposome and the loaded compound is critical. Thus, the present study was designed to investigate the effect of pH viz., 1, 3, 5, 7, 9 and 11; temperature 4, 20, 40, 60, 80 and 100 °C and exposure to illumination (4, 8, 14, 24, 48 and 72 h) on the stability of the PBP, and the physical stability of liposome stabilized with polyethylene glycol adsorbed cellulose nanocrystals. It was hypothesized that the high surface area and hydrophilic surface properties of CNC could stabilize the PBP loaded liposome by creating steric

67 hindrance, preventing the leakage of the PBP. This study includes analysis of average particle size, zeta potential, confocal laser scanning microscopic imaging of liposome

(to measure their physical stability), and the optical microscopic imaging of PEG adsorbed cellulose nanocrystals (to measure dispersibility and agglomeration behavior). Moreover, the release behavior of PBP from the liposome, total retained and percent loss of loaded PBP during exposure to different stressors are measured.

4.2. Materials and Methods

Fresh dulse (Palmaria palmata) for the extraction of phycobiliprotein was procured from Maine Fresh Sea Farms, Maine, USA. Cellulose nanocrystals suspension (11.5-12.5% of total solid) with a dimension of 5-20 nm wide and 150-200 nm long was purchased from the Process Development Center of the University of

Maine. Polyethylene glycol of molecular weights viz., 1000, 2000, 4000 and 6000 g/mol, purified soy lecithin (>99%) containing 17-24% phosphatidylinositol, 6% phosphatidic acid, 18% phosphatidylethanolamine and 35% phosphatidylcholine were purchased from Fisher Scientific Products, USA. Cellulose dialysis tube MWCO 12-

14 kDa and 50 kDa, Sephadex G-100, HDPE column (2.5 x 170 cm) with a 20 μm porosity bed support was purchased from Sigma-Aldrich, USA.

4.2.1. Adsorption of PEG with CNC

Polyethylene glycol of molecular weight 2000 and 4000 g/mol were adsorbed onto desulfated cellulose nanocrystals (DCs) following the previous method described in Chapter 3. Briefly, 3 percent of CNC suspension was placed into a 250 mL

Erlenmeyer flask and 3.33 g of PEG was added. The mixture was stirred at 200 rpm at room temperature for 3 hours. The mixture was subsequently dialyzed for three days

68 against distilled water using a 14 KDa MW cutoff dialysis tube. Dialyzed CNC containing adsorbed PEG of molecular weights 2000 and 4000 g/mole were freeze- dried using a freeze dryer (SP Scientific 2000 Virtis 35EL, Freeze Dryer, USA) and are coded as DCs-2000 and DCs-4000, respectively. CNC without PEG was labeled as

DCs. The zeta potential and size were measured using a Malvern 3000 Zetasizer HSA and infrared spectra were recorded on an ABB FTLA 2000 FTIR as mentioned in

Chapter 3.

4.2.2. Encapsulation and stabilization of phycobiliproteins

The PBP was extracted and purified using the method optimized by Dumay et al. (2013). Purified PBP was freeze-dried using a freeze dryer (SP Scientific 2000 Virtis

35EL, Freeze Dryer, USA) and stored in a deep freezer. The liposomes were prepared according to the method described in chapter 3. In brief, 600 mg of purified soy lecithin was dissolved into 200 mL of ethyl acetate and chloroform (60:40) solution. The solution was subsequently passed through a 0.22 µm filter paper to remove undissolved lecithin or any foreign particles, followed by rotary evaporation and thin layer formation in the round bottom flask. The flask with a thin layer of lecithin was placed under a vacuum dryer at room temperature overnight to remove any traces of solvent.

To encapsulate the purified PBP, the thin layer was further rehydrated with PBP solution (300 mg/50 mL) followed by the addition of 100 mL of saline phosphate buffer

(7.4 pH). The resulting multilamellar liposomes were sonicated for 1.2 minutes at 40% amplitude (at the interval of 15 seconds ON and 5 seconds OFF-pulse time) using an ultrasonic dismembrator (Fisherbrand, Ultrasonic Liquid Processor, USA) to obtain unilamellar vesicle (SUV) liposomes. This liposome was coded as phycobiliprotein

69 loaded liposome (PL). Loaded liposome was subsequently stabilized by the addition of

100 mL suspension/solution (1% w/v) of desulfated cellulose nanocrystals (DCs), or polyethylene glycol (PEG-2000 or PEG-4000) or polyethylene glycol adsorbed onto cellulose nanocrystals (DCs-2000 or DCs-4000). These stabilized liposomes are coded as PLDCs, PLP-2000, PLP-4000, PLDCs-2000 and PLDCs-4000. Stabilized liposomes were dialyzed in the absence of illumination over 48 hours at 4 ℃ using 50 kDa cellulose dialysis tubing against distilled water to remove the excess PBP.

4.2.3. Encapsulation efficiency of phycobiliproteins

The encapsulation efficiency was determined by the addition of 500 µL of a 100 mM Tween-20 solution into 5 mL of the liposome dispersion followed by centrifugation at 4,500x g for 10 minutes. The fluorescence spectrum of the supernatant was measured on a spectrofluorometer (Jobin Yvon Fluorolog-3-21) using excitation at 545 nm and emission at 575 nm (Lage-Yusty et al., 2013; Caracciolo, 2015). The measured fluorescence was due to PBP encapsulated (within liposome) and PBP adsorbed on the surface of the liposome (probably due to the protein corona mechanism between the outer surface of the liposome and PBP) (Riviere et al., 2013). An estimate of the fluorescence from the adsorbed PBP was determined by adsorption of PBP on empty liposomes prepared using the same phospholipid layer rehydration method (see chapter 3). Briefly, the dried phospholipid layer was rehydrated with 10 mL of phosphate buffer followed by sonication. Additionally, the 5 mL of PBP solution (300 mg/50 mL) was added to the liposome dispersion. The mixture was allowed 3 hours at

4 ºC to adsorb the PBP on the outer surface of liposome with a gentle stirring speed of

100 rpm (Palchetti et al., 2016). Subsequently, the mixture was dialyzed against

70 distilled water over 48 hours at 4 ℃ using 50 kDa cellulose dialysis tubing to remove the non-adsorbed PBP. 5 mL of PBP adsorbed liposome was then dissolved with 500

µL Tween-20 solutions (100 mM) followed by centrifugation at 4,500x g for 10 minutes. The fluorescence intensity of the supernatant was considered as the adsorbed

PBP. The following equation was used to determine the encapsulation efficiency.

퐴 퐸푛푐푎푝푠푢푙푎푡푖표푛 푒푓푓푖푐푖푒푛푐푦 (%) = × 100 퐵

Where A is the fluorescence intensity of PBP within liposome and B is the fluorescence intensity of total added PBP.

4.2.4. Stability analysis of phycobiliproteins

The stability of the encapsulated PBP was determined by exposing the liposomes to different stressors such as pH, temperature and illumination, as described by Munier et al. (2014) and Rastogi et al. (2015). For each experiment, a 100 mL of stabilized

PBP loaded liposome suspension was placed into 250 mL bottles (DURAN® PURE bottles) and properly covered with aluminum foil.

A 100 mM solution of HCl and NaOH was used to maintain the pH of stabilized liposome samples using phosphate buffer solution at 1, 3, 5, 7, 9 and 11. Treated samples were placed into the refrigeration chamber at 4 ℃ for 60 minutes. The stability toward temperature was determined by exposing the samples to different treatment temperatures of 4, 20, 40, 60, 80 and 100 ℃ for 60 minutes at pH 7.0. The stability of

PBP toward illumination was evaluated in a 1.0-meter square cardboard box covered with a black polyethylene sheet. The box was placed in a cooling chamber (4 to 7 ℃) and a 3.5 W bulb (16.1 µmol m-2 s-1) was placed inside the top portion of the box.

Samples in 250 mL transparent bottles were placed into the center of the preinstalled carton box and the illumination test was conducted for different time intervals of 4, 8,

14, 24, 48 and 72 hrs. The fluorescence spectra of the PBP samples were recorded on a spectrofluorometer and the total loss of PBP was calculated using the following equation:

퐷 퐿표푠푠 표푓 푃퐵푃 (%) = 100 − ( × 100) 퐸 where D is the fluorescence intensity of retained PBP during the process and E is the initial fluorescence intensity of entrapped PBP.

4.2.5. Particle size and zeta potential analysis

Average liposome size and zeta potential of the samples were performed using a Malvern Zetasizer 3000 HSA. Our preliminary investigation and observation on the effects of temperature (4 – 100 °C) and illumination intensity (16.1 µmol m-2 s-1) did not show any significant effect on the liposome size and zeta potential (preliminary observation). However, adjusting pH did affect the average liposome size and zeta potential. A 10 mL sample was taken into 25 mL of the test tube and different pH viz.,

1, 3, 5, 7, 9 and 11 was maintained for 60 minutes at 4 ℃. Afterward, samples were immediately analyzed for average liposome size. A similar procedure was followed separately for the investigation of the effect of pH on zeta potential.

4.2.6. Microscopic analysis

A 5 mL aliquot of each sample was taken into a 25 mL dark, amber-colored

Pyrex test tube and a pH of 1.0 was maintained using 0.1M HCl solution. After 60 minutes at pH 1.0, 50 µL of the sample was loaded on a separate glass slide and covered

72 with a thin glass coverslip and images were recorded on a laser scanning confocal microscopy (Leica TCS SP instrument). An excitation and emission spectra of 545 and

575 nm was used for scanning using a 100X magnification lens.

An optical microscope (AmScope Polarized Light Microscope) was used to measure the agglomeration behavior of desulfated CNC (DCs) adsorbed with polyethylene glycol (DCs-2000 and DCs-4000) at different pH. A 10 mL sample of the f PLDCs or PLDCs-2000 or DCs-4000 was placed into 25 mL test tubes. Subsequently, the pH was adjusted using 100 mM solutions of HCl and NaOH. After 60 minutes of storage at 4 ℃, 100 µL of the suspensions were placed onto glass slides and covered with coverslips. Images were processed using a 20X magnification lens.

4.2.7. Statistical analysis

Data were collected in triplicate for the average liposome size, zeta potential, and fluorescence intensities. The data were analyzed using SAS software version 9.4.

One-way ANOVA was used to determine the significant effect at the level of p≤0.05, while the Tukey’s (HSD) post-hoc test was performed to identify the significant differences among the means values.

4.3. Results and Discussion

4.3.1. Adsorption of polyethylene glycol with desulfated CNC

FTIR analysis revealed the successful adsorption of polyethylene glycol (PEG) on the DCs (Fig. 4.1). Characteristic peak of the alkane group of PEG was recognized between 2800-2900 cm-1, (stretching of C-H group) and between 1290-1440 cm-1

(scissoring and bending of C-H group). The peak at 1245 cm-1 was identified as the

73 stretching vibration C-O of alcohol group and between 1190 to 987 cm-1 as C-O-C vibration of ether group. A characteristic peak of the crystallinity of PEG was confirmed at 834 cm-1 (Sun, Zhang, & Chu, 2008).

Fig. 4.1. FTIR spectra of modified CNC.

Note: Highlighted peaks confirm the adsorption of polyethylene glycol (2000 and 4000) with desulfated cellulose nanocrystals (DCs).

4.3.2. Effect of pH on average particle size

The average size of the stabilized phycobiliprotein loaded liposome samples at different pH is shown in Fig. 4.2. At neutral pH (7.0±0.05) the PL sample had an initial average liposome size of 177±3 nm. A gradual decrease in the pH from 7.0 to 5.0, 3.0 and 1.0 resulted in a significant (p≤0.05) increase in the particle size of 424±3 nm,

1141±202 nm and 1306±80 nm, respectively. Stability of PBP loaded liposome at pH

7.0 may have been due to an anionic surface property of the of a phosphate group (zeta potential: -34±3 mV) of a phospholipid, shows that charge stabilization does not lead to the coalescing of the liposomes (see the section 4.3.3.). However, a decrease in the

+ -3 pH to 3 led to protonation (H ) of an anionic phosphate group (PO4 to PO4) of the liposome, causing a reduced zeta potential and hence to the coalescence of the liposomes into larger aggregates (Aoki et al., 2015 and Chiang, Lyu, Wen, and Lo,

2018). Conversely, an increase in the pH of the liposome dispersion medium up to pH

9.0 had no significant effect (p>0.05) on the coalescence of the liposomes as the zeta potential did not change. However, a further increase in the pH up to 11.0 led to a slight increase (p≤0.05) in the average size from 177±3 nm to 200± 4 nm.

Control experiments were performed using liposomes containing adsorbed PEG

2000 and 4000 g/mol (PLP-2000 and PLP-4000) and compared to those liposomes stabilized with CNC that contain the same adsorbed PEG (PLDCs-2000 and PLDCs-

4000). The average particle size PLP-2000 and PLP-4000 increased from 161±3 and

197±2 nm to 938±113 and 1024±45 nm, respectively, with a decrease in the pH from

7.0 to 1.0 (Fig. 4.2). In contrast, an increase in the pH from 7.0 to 11.0 resulted in a minor increase in the particle size to 193±2 and 221±3 nm for PLP-2000 and PLP-

4000, respectively. Again, the coalescence at a pH of 1.0 is the result of lower zeta potential and hence lower electrostatic stabilization of the system. No significant

(p>0.05) difference between the average particle size of the stabilized liposome with polyethylene glycol 2000 and 4000 (2000 g/mol) was obtained at neutral pH (7.0±0.05)

(Fig. 4.2). The freeze-dried desulfated CNC (DCs) and desulfated CNC adsorbed with polyethylene glycol 2000 and 4000 g/mol (DCs-2000 and DCs-4000)

Fig. 4.2. Effect of pH range on the average particle size of stabilized liposomes.

9000 pH 1.0 pH 3.0 Ba Ba pH 5.0 pH 7.0 Bb 7500 pH 9.0 pH 11.0 Ca Bc Ca

Bc 6000 Bd Cb

Cb 4500

3000 Cc Average (nm) particle size Average Ce Ce Aa 1500 Aa Aa Aa Aa Aa Ab Ab Ab Cd Cd AcAc Ac Ac Cd Cd Ad Ac Ad Ac Ad 0 PL PLP-2000 PLP-4000 PLDCs PLDCs-2000 PLDCs-4000 Sample types

Note: Phycobiliprotein loaded liposomes (PL), PBP loaded liposome stabilized with polyethylene glycol (PLP-2000 and PLP-4000), PBP loaded liposome stabilized with desulfated cellulose nanocrystals (PLDCs) and desulfated cellulose nanocrystals adsorbed with polyethylene glycol (PLDCs-2000 and PLDCs-4000). Capital letters denote significant differences (p≤0.05) among samples and small letters denote significant differences (p≤0.05) among pH. Error bars represent the standard deviation of three independent experiments.

(without liposome) had an average particle size of 7304±639, 200±23 & 198±14 nm and zeta potential value of -22±3, -16±1 & -15±3 mV at pH 7. A considerably larger average particle size of 7304±639 for the desulfated CNC as compared to DCs-2000 and DCs-4000 (200±23 & 198±14 nm) was due to the irreversible agglomeration of a surface hydroxyl group (O-H) of the CNC during the freeze-drying process (Cheng et al., 2015). However, adsorption of polyethylene glycol on the surface of the desulfated

CNC led to steric stabilization of the CNC at a size of about 200 nm. (Cheng et al.,

2015). Both desulfated CNC and desulfated CNC containing adsorbed PEG (DCs-2000

76 and DCs-4000) were employed as a steric stabilizer for the PBP loaded liposome and the effect of different pH was studied after 60 minutes after adjusting the pH. At pH

7.0, the liposomes containing adsorbed desulfated CNC (PLDCs) had approximately thirty times higher average particle size (5505±1120) as compared to the PBP loaded liposome (PL) and PBP loaded liposome stabilized with polyethylene glycol (PLP-

2000 and PLP-4000) (Fig. 4.2). This is attributed to the higher initial particle size

(7304±639 nm) of the freeze-dried desulfated CNC (DCs). Furthermore, a decrease in the pH down to 1.0 led to an increase in the average particle size of the PLDCs from

5505±1120 nm to 7975±1273 nm and a charge reversal in zeta potential from -37±5 to

+15±2. The average particle size of the PBP loaded liposome stabilized by desulfated

CNC adsorbed with polyethylene glycol (PLDCs-2000 and PLDCs-4000) was

189.43±3.14 and 206.97±2.36 at pH 7.0. Results showed that the average particle size of PLDCs-2000 and PLDCs-4000 samples were significantly different compared to the particle size of the PBP loaded liposome (PL), PLP-2000 and PLP-4000. It was observed that the average particle size of the PLDCs-2000 and PLDCs-4000 samples increased significantly up to 2192.67±284.83 & 1959.50±201.73 nm at pH 5.0 and

3186.34±634.97 and 2728.10±618.63 nm at pH 3.0, followed by 6464.33±211.65 and

6698.55±317.53 nm at pH 1.0. It was assumed that the increase in the average particle size might be due to the desorption of polyethylene glycol from the surface of desulfated CNC at lower pH (see in chapter 3), that may have led to the agglomeration of desulfated CNC (Fig 4.3). This may also have caused the coalescence of PBP loaded liposome, thereby significant increase in the particle size. The following mechanisms were hypothesized for the agglomeration and coalescence of the samples. First the

Fig. 4.3. Schematic illustration of behavior of liposomes and CNC at lower pH.

Note: desulfated cellulose nanocrystals (DCs) as an effect of lower pH ( ) and their agglomeration. (A) Stabilized PBP loaded liposome, (B) molecular structure of liposome (phosphatidylcholine) showing anionic phosphate group and cationic choline + - group, (C) protonation (H ) of a phosphate group (PO4 ) of the liposome, (D,E) Agglomeration of liposome and DCs together due to entangling effect caused by the addition of cationic liposome with anionic DCs due to ionic interaction, (E) protonation + - (H ) of anionic surface group of DCs (SO4 ) and agglomeration, (F) Protonation of an anionic group of the desulfated CNC resulting in agglomeration and precipitation of DCs.

+ - protonation (H ) of phosphate (PO4 ) group of the liposome (Aoki et al., 2015 and Kono et al., 2018); second, the ionic interaction of protonated liposome (cationic at lower pH) with slight anionic desulfated cellulose nanocrystals (DCs) resulting entangling effect (Navon, Radavidson, Putaux, Jean, & Heux, 2017); third, the successive agglomeration of desulfated CNC at lower pH due to protonation of its residual sulfate group (SO-3) (Eyley et al., 2015) (Fig. 4.3).

4.3.3. Effect of pH on Zeta potential

The average zeta potential value of the samples is given in Fig. 4.4. PL contained a zeta potential value of -33±3 mV at pH 7.0. With a decrease in the pH from pH 7.0 to pH

5.0, the zeta potential value decreased to -12±2 mV. Further, a decrease in the pH down to 3.0 and 1.0, resulted in the zeta potential turning positive with values of +8±2.74 to

+18±3 mV. The positive zeta potential values were due to the positive choline group

+ -3 of the liposomes and the protonation (H ) of PO4 group of the liposome (Fan, Chen,

Huang, Zhang, & Lin, 2017) (Fig. 3). An increase in the average particle size of the samples may also have led to less surface area, reducing the charge density on the surface, and leading to aggregation. (Yadav et al., 2011). The trend in zeta potential with pH agrees with Peng et al. (2018), who reported that the zeta potential value of liposome prepared using Sunlipon-75 (a type of phospholipids) decreased down to zero when the pH was reduced to 3.0 and afterword gradually changed into positive values at pH 2.0. Furthermore, no significant change in the zeta potential values of the PBP loaded liposome (PL) was observed when the pH was increased to 9.0 and 11.0 and this is in agreement with Peng et al. (2018). A similar trend in zeta potential with pH for PL was obtained for PBP loaded liposome with polyethylene glycol 2000 and 4000 g/mol (PLP-2000 and PLP-4000). Results also displayed that the zeta potential value of PLP-2000 and PLP-4000 were slightly lower compared to PBP loaded liposome

(PL) at pH 7.0, 9.0 and 11.0 (Fig. 4.4). Studies show that the polyethylene glycol molecules can lower the zeta potentials value through the surface coating of the colloidal system (Cheng et al., 2015).

Fig. 4.4. Effect of pH range on the zeta potential value of stabilized liposomes.

Note: PBP loaded liposome (PL), PBP loaded liposome stabilized with polyethylene glycol (PLP-2000 and PLP-4000), PBP loaded liposome stabilized with desulfated cellulose nanocrystals (PLDCs) and desulfated cellulose nanocrystals adsorbed with polyethylene glycol (PLDCs-2000 and PLDCs-4000). Capital letters denote significant differences (p≤0.05) among samples and small letters denote significant differences (p≤0.05) among pH. Error bars represent the standard deviation of three independent experiments.

The zeta potential values of PBP loaded liposome (PL), PL-2000 and PLP-4000 at pH 7.0 and above (9.0 and 11.0); were slightly lower than obtained for PBP loaded liposome with desulfated CNC (PLDCs), PLDCs-2000 and PLDCs-4000 (Fig. 4.4).

This difference could be due to the contribution from the lower average zeta potential of desulfated CNC (-22±3 mV). Furthermore, higher zeta potential values were recorded at pH of 5.0 and 3.0 for the PLDCs, PLDCs-2000 and PLDCs-4000 (-11±3 and -2±0.7, -19±0.4 and -3±0.3 and -18±2 and -4±0.7 mV, respectively) compared to

PLP-2000 and PLP-4000. Again, these higher values may be due to the contribution from the lower isoelectric point of the cellulose nanocrystals (around 2.5) (Eyley et al.,

2015). Further decrease in the pH to 1.0 resulted in a positive zeta potential values of

+15±3, +16±3 and +14±1 mV, respectively, which may be due to neutralization of the anionic group of the CNC below its isoelectric point (Eyley et al., 2015; Bai et al.,

2019a).

4.3.4. Effect on the coalescence and agglomeration of behavior of liposomes and cellulose nanocrystals

To confirm the increase in the average particle size and decrease in the zeta potential value, we conducted the confocal laser scanning microscopic analysis of PBP loaded liposome followed by optical microscopic observation of desulfated CNC with polyethylene glycol. Fig. 4.5 shows the confocal image of PBP loaded liposomes with different types of stabilizers at pH 7.0 and 1.0. The images show that the liposome particles were unevenly dispersed into the aqueous medium for all of the samples; however, a decrease in the pH led to effective coalescence. This coalescence behavior

Fig. 4.5. Confocal laser scanning microscopic images of stabilized liposomes.

Note: Agglomeration behavior phycobiliprotein loaded liposomes. pH 7.0 - first row and pH 1.0 - second row. The scale bar is 2 µm. PBP loaded liposome (PL), PBP loaded liposome stabilized with polyethylene glycol (PLP-2000 and PLP-4000), PBP loaded liposome stabilized with desulfated cellulose nanocrystals (PLDCs) and desulfated cellulose nanocrystals adsorbed with polyethylene glycol (PLDCs-2000 and PLDCs- 4000). 81 of the liposome supports our hypothesis that an increase in the average particle size of the samples was due to the coalescence of liposomes at lower pH (Fig. 4.3). Optical microscopic images of the desulfated CNC (DCs) and polyethylene glycol (2000 &

4000 g/mol) adsorbed desulfated CNC with phycobiliprotein loaded liposomes are given in Fig. 4.6. It was observed that the desulfated CNC (DCs) with phycobiliprotein loaded liposome clustered at all pH values. However, PLDCs-2000 and PLDCs-4000

(Fig. 4.6) showed no observable cluster at the pH condition of 7.0, 9.0 and 11.0 and some tiny clusters of approximately 10 µm at pH 5.0. Further, a decrease in the pH from 5.0 to 3.0 showed a group of clusters of about 20 to 30 to 50 µm in diameter; however, further, decrease in pH to 1.0 showed a complete agglomeration of desulfated

CNC resulting a cluster of similar to optical microscopic images of PLDCs (Fig. 4.6).

The agglomeration of PLDCs-2000 and PLDCs-4000 was hypothesized to be due to the desorption of polyethylene glycol at lower and higher pH levels (chapter 3).

Fig. 4.6. Optical microscopic image of stabilized liposomes samples at different pH.

Note: Polyethylene glycol adsorbed desulfated CNC (PLDCs-2000 and PLDCs-4000). Scale bar 10 µm.

4.3.5. Effect of pH on the stability of phycobiliproteins

Fluorescence intensity values (centipoise - CPS) of the PBP for the samples viz., PBPs, PL, PLP-2000, PLP-4000, PLDCs, PLDCs-2000 and PLDCs-4000 are shown in Fig. 4.7. There is a substantial decrease in the fluorescence intensity for the

PBP in solution (PBPs) with a gradual decrease in the pH from 7.0 to 1.0, resulting in a significant (p≤0.05) loss of 91±2%. In contrast, raising the pH from 7.0 to 11.0 had comparatively less effect on the fluorescence intensity resulting in a total loss of 32±2%

(Fig. 4.8). A similar effect of the pH on the fluorescence intensity of the PBP extracted from Phormidium rubidium was observed by Rastogi et al. (2015). They found a total loss of 30.19 and 99.58% after an exposure period of 60 minutes at pH 4.0 and pH 2.0.

Considering the effect of basic pH environment on the stability of extracted PBP,

Fig. 4.7. Effect of stressors viz., pH (a), temperature (b) and illumination (c) on the fluorescence intensity of PBP.

Note: Values are given in CPS are 1000 times higher of the given values. Error bars represent the standard deviation of three independent experiments. PBP loaded liposome (PL), PBP loaded liposome stabilized with polyethylene glycol (PLP-2000 and PLP-4000), PBP loaded liposome stabilized with desulfated cellulose nanocrystals (PLDCs) and desulfated cellulose nanocrystals adsorbed with polyethylene glycol (PLDCs-2000 and PLDCs-4000). Error bars represent the standard deviation of three independent experiments.

Munier et al. (2014) found a total loss of up to 97% of the fluorescence intensity of the

PBP at pH 12.0, which was considerably higher compared to the value of 32% obtained in this study at pH 11.0. The loss of fluorescence intensity of PBP was attributed to the dissociation or denaturation of the subunits of PBP into monomeric units such as α-, β- and γ- etc., of the PBP (Liu et al., 2009). An extreme change in the pH of the PBP’s environment alters the chromophore structure by disturbing its electrostatic property and O-H bond of protein structure (Szalontai, Gombos, Csizmadia, & Lutz, 1987).

These properties vary with the sources of PBP and types of monomeric subunits (Liu et al., 2009; Munier et al., 2014 and Rastogi et al., 2015).

For the samples containing PBP in liposomes (PL, PLP-2000, PLP-4000,

PLDCs, PLDCs-2000, and PLDCs-4000), a slight decrease in the pH from 7.0 to 5.0 had no significant (p>0.05) effect on the fluorescence intensity (Fig. 4.7). Liposome bilayers have good stability at pH above 5.0 and show slight chemical shifts of polar head-groups of the phospholipid and mild protonation of phosphoric groups

(Sułkowski et al., 2005 and Ding et al 2009).

A further decrease in the pH from 7.0 to 1.0 reduced the fluorescence intensity of the PBP significantly (p≤0.5) (Fig. 7), accounting for a total loss of 28±2.1, 9±3.8,

13±2.12, 25±1.7, 19±0.7 and 17±0.3% for PL, PLP-2000, PLP-4000, PLDCs, PLDCs-

2000, and PLDCs-4000, respectively (Fig. 4.8). These losses may be due to destabilization of the liposome’s bilayer at lower pH that increased the permeability of the bilayer resulting more release of the PBP and causing the loss of the fluorescence intensity (Ding et al., 2009; Yadav et al., 2011 and Fan et al., 2017). Results showed that the PLP-2000 and PLP-4000 retained approximately 15% more fluorescence

85 intensity as compared to the PL and PLDCs and approximately 7% more than PLDCs-

2000 and PLDCs-4000. As illustrated in Figure 3, a lower pH caused the agglomeration of the CNC, resulting in less steric hindrance for the protonated liposomes. This might have caused the coalescence of liposomes and loose liposome bilayers, resulting in the release of PBP.

4.3.6. Effect of temperatures on the stability of phycobiliproteins

The temperature had the most noticeable effect on the fluorescence intensity of the PBP in solution (PBPs), which decreased drastically with a gradual increase in the exposure temperatures from 4 ℃ to 20, 40 and 60 ℃ (Fig. 4.7). This resulted in a total loss of 8.4±2.9, 39.5±2.6 and 70.7±1.4, respectively (Fig. 4.8). Further increase in the temperature above 60 ℃, resulted in a complete loss of the fluorescence intensity.

Munier et al. (2014) and Rastogi et al. (2015) reported a similar trend for the loss of fluorescence intensity of the PBP isolated from Porphyridium cruentum and

Phormidium rubidium. Our results demonstrate the extreme instability of PBP above

40 ℃. PBP has a distinguishing characteristic of fluorescence due to its linear tetrapyrrole bilin chromophores attached to a protein molecule (Glazer and Cohen-

Bazire, 197; Sidler, 1994). Likewise, the deleterious effect of the lower pH of 3.0 and

1.0 on fluorescence intensity of the PBP, higher temperature also might have damaged the double covalent bond between tetrapyrrole bilin chromophores and protein conjugates, causing the loss of the fluorescence intensity (Glazer et al., 1994).

Considering the stability of the fluorescence intensity for the PL, PLP-2000,

PLP-4000, PLDCs, PLDCs-2000 and PLDCs-4000, no significant (p>0.05) loss was observed at the exposure temperature of 20 ℃ as compared to those stored at 4 ℃ (Fig.

Fig. 4.8. Effect of stressors viz., pH (a), temperature (b) and illumination (c) on the % loss of the fluorescence intensity of PBP.

100 PBPs PL PLP-2000 PLP-4000 a 80 PLDCs PLDCs-2000 PLDCs-4000

Fluorescence intensity (% loss) loss) (% intensity Fluorescence 20

0 1 3 5 7 9 11 -20 pH levels

120 PBPs PL b PLP-2000 PLP-4000 100 PLDCs PLDCs-2000 PLDCs-4000 80

20 Fluorescence intensity(% loss) Fluorescence 0 4 ℃ 20 ℃ 40 ℃ 60 ℃ 80 ℃ 100 ℃ -20 Temperature (ºC)

100 PBPs PL c PLP-2000 PLP-4000 80 PLDCs PLDCs-2000 PLDCs-4000

20 Fluorescence intensity(% loss) Fluorescence 0 4 8 14 24 48 72 Time (hours) -20

Note: Error bars represent the standard deviation of three independent experiments. PBP loaded liposome (PL), PBP loaded liposome stabilized with polyethylene glycol (PLP-2000 and PLP-4000), PBP loaded liposome stabilized with desulfated cellulose nanocrystals (PLDCs) and desulfated cellulose nanocrystals adsorbed with polyethylene glycol (PLDCs-2000 and PLDCs-4000). Error bars represent the standard deviation of three independent experiments.

4.7 and Fig. 4.8). Further increase in the temperature from 20 ℃ to 40 ℃, PL, PLP-

2000, PLP-4000 and PLDCs samples showed lower fluorescence intensity recorded a total loss of up to 35% as compared to the PLDCs-2000 and PLDCs-4000, which retained approximately 20% more fluorescence intensity. It was presumed that the PEG adsorbed desulfated CNC (DCs-2000 and DCs-4000) might have provided the steric stability to the liposomes by coating the surface resulting in less motile phospholipid bilayers thereby less leakage and denaturation of the PBP. Further, increase in the temperature from 40 ℃ to 60 ℃, the total loss difference between the samples with

(PLDCs-2000 and PLDCs-4000) and without (PL, PLP-2000, PLP-4000 and PLDCs)

PEG adsorbed desulfated CNC decreased from 20% to approximately 7%. A complete loss of the fluorescence intensity was recorded when temperature increased from 60 to

80 ℃. A similar effect on the activity of flavorsome loaded within liposome was reported by Vafabakhsh et al. (2013) at different temperatures, who found a complete loss of the fluorescence activity when exposed at 85℃.

4.3.7. Effects of illumination on the stability of phycobiliproteins

Illumination had the least significant effect on the fluorescence intensity of the

PBP as compared to the effect of pH and temperature, resulting in an insignificant

(p>0.05) loss over 4 hours of storage period. (Fig. 4.7). Further increase in the storage period from 4 to 8 hours significantly (p≤0.05) reduced the fluorescence intensity for the PBP in solution (PBPs), PL, PLP-2000, PLP-4000, and PLDCs, resulting in a total loss of 13±1.1, 14±2.1, 13±3.9, 11±2.7 and 16±2.7% There was an insignificant difference among the total loss for PLP-2000, PLP-4000, and PLDCs, which inferred that PEG-2000 and PEG-4000 did not show any preserving activity against illumination for the PBP despite ensuring the physicochemical stability of the liposome. Because the liposome’s bilayers are transparent, illumination could have easily penetrated the bilayer over time, effecting the fluorescence intensity of the PBP.

Further progression in the storage period from 8 to 14 hours, a gradual decrease in the fluorescence intensity for the PBP in solution (PBPs), PL, PLP-2000, PLP-4000 and

PLDCs was recorded, resulting in a total loss of 29±0.7, 23±1. 3, 24±2.5, 22±0.9 and

31±3.6%, respectively. In contrast to the effect of illumination on PLP-2000, PLP-

4000, and PLDCs, PLDCs-2000 and PLDCs-4000 significantly remained the fluorescence intensity resulting in zero loss over the storage period of 14 hours (Fig.

4.8). Throughout a total storage period of 72 hours, PLDCs-2000 and PLDCs-4000 displayed promising stabilizing activity for the fluorescence intensity of PBP in liposome against illumination, accounted only a total loss of 32±4.4 and 34±1.0%, respectively. This shows that the DCs-2000 and DCs-4000 had a significant stabilizing activity for the fluorescence intensity of PBP against illumination, which may be due to the partial UV-visible shielding activity of cellulose nanocrystals (Sirviö, Visanko,

Heiskanen, & Liimatainen, 2016). However, a total loss of 84±1.3, 78±2.8, 74±2.9,

74±1.7 and 75±2.3% were recorded for the PBP in solution (PBPs), PL, PLP-2000,

PLP-4000, and PLDCs. At illumination intensity of 33.57 μmol m−2 s−1 Munier et al.

(2014) reported a total loss of fluorescence intensity of PBP of up to 70% after a storage period of 24 hours at 4 ℃. In agreement with our results, Rossano et al. (2003) has reported that the exposure of illumination causes instability of R-phycoerythrin, a type of PBP due to the loss of tetrapyrrole bilin chromophores.

4.4. Conclusion

An effective re-dispersion of desulfated cellulose nanocrystals into the aqueous medium was achieved through adsorption of polyethylene glycol (2000 and 4000 g/mol). FTIR analysis confirmed the adsorption of PEG on the surface of the desulfated

CNC. Desulfated CNC adsorbed with polyethylene glycol was used as a stabilizing material for the PBP loaded liposome. Results displayed that the DCs-2000 and DCs-

4000 retained 20% more fluorescence intensity at 40 ℃ as compared to PBP in solution

(PBPs), PL, PLP-2000, PLP-4000 and PLDCs. However, at the higher temperature of above 80 ℃, DCs-2000 and DCs-4000 had an insignificant (p>0.05) effect on the stability of fluorescence intensity, resulting in a complete loss by heat. Similarly, DCs-

2000 and DCs-4000 retained the fluorescence intensity of PBP up to 65% against illumination after a storage period of 72 hours; however, the total fluorescence intensity of up to 25% was retained for PL, PLP-2000, PLP-4000, PLDCs and 15% for the PBP in solution (PBPs). pH affected the average particle size and zeta potential values of the samples along with fluorescence intensity, resulted in a significant (p≤0.05) increase in the average particle size due to agglomeration of PEG adsorbed desulfated

CNC and coalescence of liposomes. Lower pH of below 5.0 also caused the charge reversal of liposomes resulting in positive zeta potential. Confocal laser scanning microscopic and optical microscopic analysis confirmed the coalescence of PBP loaded liposome and the agglomeration of PEG adsorbed DCs at pH 1.0. The study concluded that the encapsulated PBP had a good stability below 60 °C, above pH of 5.0 and against illumination. However, further modification of DCs is needed to enhance its stabilization at a lower pH for delivering sensitive bioactive compounds.

CHAPTER 5

PHYSICOCHEMICAL STABILITY OF O/W PICKERING EMULSION

STABILIZED WITH LAURIC ACID ADSORBED CELLULOSE

NANOCRYSTALS AS AFFECTED BY DILUTION, pH AND STORAGE

PERIOD

5.1. Introduction Pickering emulsion is a well-known type of emulsion system in which a physically unstable dispersal phase is stabilized by solid colloid sized particles (Ramsden, 1904 and Pickering, 1907). In the past, a variety of colloidal particles and polymers have been utilized to ensure the physicochemical stability and dispersity of the oil in water

(O/W) type emulsion, which includes synthetic polymers, surfactants and nano metals

(Midmore, 1998; Ashby and Binks, 2000; Tikekar et al., 2013; Sharma et al., 2015;

Shah et al., 2016). However, an increased sustainability concern and the demand of food-grade stabilizing materials have pulled the researcher’s interest towards natural polymers and colloidal particles such as solid lipids (Dieng et al., 2019a; Dieng et al.,

2019b), proteins (Dhakane et al., 2017 and Dai et al., 2018) and polysaccharides (Shao et al., 2018; Ye et al., 2018; Fu et al., 2019). Furthermore, the effort to replace commercial synthetic stabilizing materials has led researchers to explore for more sustainable and novel sources, including cellulose nanocrystals, also known as crystalline cellulose or cellulose nanowhiskers (Abiaziem et al., 2019), cellulose nanofibers (Winuprasith et al., 2018) and TEMPO-oxidized cellulose (Goi et al., 2019).

Cellulose nanocrystals (CNC) possess several unique features such as nano-sized dimension (150-200 nm long and 5-15 nm wide) (Cheng et al., 2015), non-toxic

(Seabra et al., 2018), dispersible in aqueous medium but insoluble (Cheng et al., 2015), cheaply produced from wood and other underutilized bioresources (Trache et al., 2017;

Grishkewich et al., 2017), and contains a large number of surface O-H groups (Tang et al., 2017).

CNC is produced via different methods such as acid hydrolysis (includes H2SO4,

HCl, HNO3, Citric acid, H3PO4 etc.), alkali hydrolysis (NaOH) and oxidation (Sharma et al., 2019). However, several studies have reported that the CNC produced through sulfuric acid hydrolysis has shown better emulsion stabilizing activity due to its high electrical property contributed by surface sulfate groups (Cheng et al., 2015; Bai et al.,

2019a; Zhang et al., 2019). CNC helps to create a stable Pickering emulsion through providing 1) a strong electrostatic repulsion and 2) steric hindrance (Bai et al., 2019a;

- Zhang et al., 2019). Higher the sulfate groups (-SO3 ) on the surface of CNC, more stable the Pickering emulsion is, due to stronger electrostatic repulsion created among the oil droplets (Zhang et al., 2019). Moreover, CNC adsorbed on interfaces of the oil droplets create steric hindrance (Cherhal et al., 2016; Bai et al., 2018). Bergfreund &

Fischer (2019) and Bertsch et al. (2019) have reported that the efficacy of the CNC can be improved by adding salt that can improve the rate of adsorption of CNC on the interface of oil droplets due to increased surface pressure. Bai et al. (2019a) found that addition of salt to the O/W Pickering emulsion can affect the electrostatic interactions property of CNC such as (i) altered adsorption of CNC to the interface, (ii) altered packing density of CNC at the interface and (iii) altered CNC-CNC interactions in the aqueous phase. An increased CNC-CNC interaction can enhance the stability of an emulsion by altering rheological properties, e.g., increasing gel strength of aqueous

93 phase that limits the motion of the oil droplets into a continuous phase, thereby less interaction among oil droplets thus stable emulsion. However, it has been reported that the stability of these types of emulsion may significantly be affected by dilution and addition of opposite ions (Wooster, 2006; Maphosa and Jideani, 2018). Dilution can lead to a segregative phase separation to the stable Pickering emulsion due to the following reasons: (a) desorption of the polymer from the oil droplet interface tumbling steric or electrostatic repulsion among droplets, (b) weaker gel strength of aqueous medium, thereby increased droplets interaction thus coalescence/Ostwald ripening and

(c) reduced surface pressure on the oil droplets due to decreased salt concentration desorption of adsorbed polymers (Wooster, 2006; Maphosa and Jideani, 2018; Bai et al., 2019a).

It has been reported that the CNC with hydrophobic or amphiphilic surface property can be an effective way to create stable Pickering emulsion (Zoppe et al., 2012;

Girouard et al., 2016; Saidane et al., 2016). Chemically modified CNC shows good stability against dilution and ionic stress due to its stronger hydrophobic-hydrophobic interaction between oil and CNC interface as compared to emulsion stabilized through unmodified CNC. However, the chemical modification process is complex and requires a large number of organic chemicals and other reagents, which may not be a food-grade and sustainable. In this work, a hypothesis was made, modifying the surface of CNC through adsorption of an amphiphilic molecule such as fatty acid to create a hydrophobic surface, which can provide a better lipophilic-lipophobic balance between oil droplets and CNC, creating a stable Pickering emulsion.

Thus, a stable beta-carotene O/W Pickering emulsion was created using different concentrations (0.5, 1.0, 1.5 and 2.0%) of lauric acid adsorbed CNC (CL) and neat

CNC separately. Effect of different dilution factors (1-10x), pH (1-11) and storage period (1-30 days) on the physicochemical stability of the Pickering emulsion was analyzed. Following quality attributes, including average particle size, zeta potential, optical microscopic image, creaming index (CI), lipid oxidation (TBARS assay), and beta-carotene stability, were recorded during the experiment.

5.2. Materials and Methods

Sulfuric acid-hydrolyzed CNC (11.5 to 12.5%) with a dimension of 150-200 nm long and 5-20 nm wide was purchased from the Process Development Center (PDC) of the University of Maine. Canola oil (Great value, Walmart, Bangor, Maine) was purchased from the local supermarket of Bangor, Maine. Lauric acid flake (>99% pure), 2-thiobarbituric acid (TBA), trichloroacetic acid (TCA), butylated hydroxytoluene (BHT), 1,1,3,3-tetraethoxypropane (TEP) and other analytical grade solvents were purchased from Acros Organics/Fisher Scientific, USA. Beta-carotene

(standard grade) was purchased from Sigma Aldrich, USA.

5.2.1. Adsorption of lauric acid with CNC

A 20 g quantity of lauric acid powder was dissolved in 400 mL of ethanol; subsequently, a 100 g of CNC (11.5 to 12.5%) was added to the solution. Mixture was then stirred at 500 rpm until the CNC was distributed homogeneously, then the mixture was consistently stirred for 24 hours at 100 rpm at room temperature. The adsorption took place in an airtight one-liter bottle to prevent the evaporation of the solvent.

Mixture was then centrifuged at 21,800x g for 10 minutes at room temperature using a

95 super speed centrifuge (Beckman Coulter Avanti J-E Floor, CA, USA). The pellet was collected and washed repeatedly with ethanol. The pellet was later vacuum dried at 60

°C overnight and stored into a Ziploc bag at room temperature for further use. Later the adsorption of lauric acid was confirmed by FTIR (ABB, FTLA 2000) analysis and the average particle size and zeta potential were measured using zeta-sizer (Zetasizer

Malvern 3000 HAS). The amount of the adsorbed lauric acid was quantified, preparing a blend of the nine different concentrations of lauric acid (10, 20, 30, 50, 75, 100, 150,

200 and 250 mg/500 mg of dried CNC). FTIR % transmittance was recorded, and the

% transmittance value was converted to the absorbance (Fig. 5.1) following the equation developed by Toft et al. (1993):

퐴푏푠표푟푏푎푛푐푒 = 2 − 푙표푔 (%푇)

Obtained absorbance values were plotted to a calibration curve to calculate the amount of lauric acid adsorbed (Fig. 5.1).

Fig. 5.1. FTIR spectra of the lauric acid mixed CNC (a) and calibration curve of lauric acids with different concentrations (b).

Note: Different concentrations of lauric acid (10-250 mg) mixed with CNC (500 mg) to plot a calibration curve to quantify the amount of adsorbed lauric acids on the surface of CNC.

5.2.2. Preparation of beta-carotene O/W emulsion and stabilization

Different concentration of lauric acid adsorbed CNC (CL) dispersion (0.5, 1.0,

1.5 and 2.0%) was prepared by re-dispersing dried CL into distilled water stirring at

400 rpm for one hour and coded as CL-0.5, CL-1.0, CL-1.5 and CL-2.0, respectively.

Since the adsorption of lauric acid decreased the pH of the disperse solution to 4.1±0.2, pH was neutralized to 7.0 using 0.1 N NaCl. Similarly, a dispersion of neat CNC (never dried CNC without lauric acid) of the same concentrations (0.5, 1.0, 1.5 and 2.0%) were prepared by dilution with distilled water and coded as C-0.5, C-1.0, C-1.5, and C-2.0.

10 mL of beta-carotene dissolved canola oil (1 mg/100 mL) was added to the 90 mL of

CL and CNC dispersed solution (a continuous medium) and a coarse emulsion was prepared using an Ultra-Turrax (IKA S 18 N - 19 G Dispersing Element) for 2 minutes at 10,000 rpm. In addition to the emulsion prepared into neat CNC dispersed continuous medium, a 0.06 g of NaCl was added to facilitate the stabilization activity of CNC as described by Bai et al. (2018). A fine emulsion was prepared by sonicating the coarse emulsion for 5 minutes at 4-6 ℃ using an ultrasonic dismembrator (500W, 20 kHz,

Fisherbrand™ Model 505, USA) facilitated with a ¾ inch titanium alloy microtip probe. A 40 percent amplitude of the sonication at the interval of 15 seconds ON and 5 seconds OFF-pulse time was used during the process (Mehmood et al., 2017).

5.2.3. Stability study of emulsions

Physicochemical stability of the beta-carotene O/W emulsion stabilized with CL and CNC was investigated, exposing the emulsion to different stressing factors such as dilution, pH and storage period to predict the approximate stability of emulsion during food processing and consumption.

5.2.3.1. Effect of dilutions

Processed foods encounter different dilutions or addition of different solvent mediums during processing and consumption. An emulsion system must be stable when added to a different food matrix and subjected to the digestion process. To understand the effect of dilution on the stability of created Pickering emulsions, a series of dilution viz., initial (1x), 1:1 (2x), 1:3 (4x), 1:5 (6x), 1:7 times (8x) and 1:9 (10x) with distilled water was performed. In brief, a dilution level of 6x was prepared by mixing 1 mL of emulsion with 5 mL of distilled water. Diluted emulsion samples were kept in the dark at room temperature for 4 hours and the characterization was performed subsequently. For each type of attribute analysis, fresh samples were prepared, and the effect of dilution was investigated in triplicate with simultaneous duplication of the process.

5.2.3.2. Effect of pH

Since pH is the major factor causing the electrostatic screening of the electrostatically stable emulsion system, the stabilized emulsion was exposed to different pH of 1, 3, 5, 7, 9 and 11, respectively. A 10 mL of the emulsion was placed into an amber-colored Erlenmeyer flask and the respective pH was maintained using

0.1 N HCl and 0.1 N NaOH. The samples were stored in the dark at room temperature for 2 hours of reaction period and subsequently, samples were analyzed.

5.2.3.3. Effect of storage periods

A 100 mL of prepared emulsions were placed into 250 mL amber-colored

Erlenmeyer flask and stored in the dark at room temperature (Tikekar et al., 2013). The

99 quality attributes of the stabilized emulsion such as average particle size, creaming index, TBARS assay and beta-carotene were recorded after 0, 1, 2, 3, 5, 10, 15, 20 and

30 days of storage period.

5.2.4. Particle size and zeta potential analysis

Average particle size and zeta potential of the stabilized emulsion were performed on dynamic light scattering zeta-sizer (Zetasizer Malvern 3000 HSA) as described by Zhang et al. (2019). The samples were diluted with distilled water to prevent the multiple scattering effects during analysis. The refractive index for the canola oil was fixed at 1.465 (Przybylski, 2001) and 1.330 for water (Zhang et al.,

2019).

5.2.5. Optical microscopic analysis

Optical microscopic images of the stabilized emulsion were performed on

AmScope Polarized Light Microscope. A 100 µL of emulsion samples from each experiment were placed on a glass slide and images were acquired simultaneously at

20x magnification of the lens (Zhang et al., 2019).

5.2.6. Creaming index

Creaming index of the samples was recorded following the method suggested by Varka et al. (2010) with some modifications. Instead of measuring the height of the separated oil phase and the continuous phase, the amount of the beta-carotene was measured between both phases using a multi-plate reader UV-Vis spectrophotometer

(BioTek Eon, Vermont, U.S.A) at 452 nm. In brief, 1 mL of sample was withdrawn from the separated phase and the continuous phase separately and dissolved into 10 mL

100 of n-hexane by vortexing for 1 minute. The solution was stored at 4 ℃ until a clear layer of the aqueous phase and n-hexane appeared. A 200 µl of fraction from the top layer was placed into 96 well microplates and absorbance was recorded at 452 nm. The percent creaming index was calculated using the following equation:

퐴푏푠푡표푝 × 100 퐴푏푠푡표푡푎푙

퐴푏푠푡표푡푎푙 = 퐴푏푠푡표푝 + 퐴푏푠푐표푛푡𝑖푛푢표푢푠 푝ℎ푎푠푒

Where Abstop is the absorbance of beta-carotene in the separated oil phase.

Additionally, the UV-Vis spectra of the beta-carotene present of the Pickering emulsion was analyzed, withdrawing 1 mL of emulsion from the center of the experimental vessel and 10 mL of n-hexane was added. Mixture was vortexed for 1 minute then stored at 4℃ until a clear layer of the aqueous phase and n-hexane appeared. A 200 µl of fraction from the top layer was placed into 96 well microplates and spectral scanning was performed between 330 nm to 550 nm.

5.2.7. TBARS analysis

Thiobarbituric acid reactive substances assay (TBARS) analysis of the

Pickering emulsion stored at ambient temperature in the dark was recorded over 30 days of storage periods. Malondialdehyde present in the emulsion was measured using the TBARS assay following the method of Alamed et al. (2006). One liter of TBA reagent was prepared by dissolving 3.75 g of TBA, 150 g of TCA and 2% BHT into ethanol, then into 750 mL of 0.25 M HCl solution. 0.5 mL of emulsion sample was taken into the screw cap glass test tube and 9.5 mL of TBA reagent was added. The test tube was placed into a preheated water bath at 90 ℃ for 15 minutes, followed by 101 immediate cooling by placing it into the refrigerator for 10 minutes. The reaction medium was then centrifuged at 3,480x g for 15 minutes at room temperature using a centrifuge model Beckman Coulter Avanti J-E Floor, CA, USA. 200 µL of the supernatant was withdrawn and placed into 96 well microplates, and absorbance was recorded at 532 nm. A calibration curve of the different concentrations of TEP was used as a standard to quantify the TBARS values (Fig. 5.2). TBARS analysis of the control emulsion sample was prepared following the protocol of Bai et al. (2019b). A

10 mL of beta-carotene dissolved canola oil (1 mg/100 mL) was added to the 90 mL of

1% solution of Gum Arabic. A fine emulsion was prepared following the stapes discussed in sub-section 2.2.

Fig. 5.2. Calibration curve of the different concentrations of 1,1,3,3- tetraethoxypropane (TEP) standard.

4 y = 0.0682x + 0.4473 3.5 R² = 0.9989

2.5

2 Absorbance 1.5

0.5

0 0 10 20 30 40 50

TEP (µmol)

102

5.2.8. Beta-carotene stability

Beta-carotene stability during the storage period of up to 30 days was recorded at room temperature under dark conditions. Quantification was performed, taking 5 mL of an emulsion into 20 mL of n-hexane and vortexing for 1 minute and stored at 4 ℃ until a clear layer of the water and n-hexane was formed. A 200 µl of fraction from the yellow portion was placed into 96 well microplates and absorbance was recorded at

452 nm. A standard curve of the beta-carotene was plotted to calculate the amount of beta-carotene (Fig. 5.3).

Fig. 5.3. Calibration curve of the different amounts of beta-carotene standard.

3.5 y = 0.1441x + 0.3191 R² = 0.9975 3

2.5

1.5 Absorbance

0.5

0 0 5 10 15 20

Beta-carotene (mg)

5.2.9. Mineral analysis

The mineral content of the CNC sample was analyzed after getting the spike in lipid oxidation of emulsion stabilized with CNC as compared to reference emulsion (no

103

CNC rather a gum arabic). Assuming that it may be due to the presence of some transition metals in the CNC causing increased lipid oxidation, the elemental analysis was performed on an Inductively Coupled Plasma Mass Spectrometry (ICP-MS) device

(Finnigan Element2, Thermo Fisher, Waltham, MA). A 10 g of neat CNC was dried overnight at 80 °C to remove the moisture content. Dried CNC was exposed to a high temperature of 400 ℃ for 5 hrs. The resulting ash was dissolved into 20 mL of 1% nitric acid solution and the sample was loaded on an autosampler (CETAC ASX-100).

Distilled water and ICP-MS lab blank were also analyzed for the mineral content to account for possible contamination during sample preparation.

5.2.10. Statistical analysis

Each experiment was conducted in triplicate. A Tukey’s (HSD) post hoc test was performed on SAS® Studio (Version 9.4, University edition) to identify the significant differences among the mean values. Analysis of Variance (ANOVA) was used to determine the significant effect among treatments at the level of p≤0.05.

5.3. Results and Discussion

5.3.1. Characterization of lauric acid adsorbed CNC (CL)

The adsorption of the lauric acid on the CNC was confirmed by FTIR analysis

(Fig. 5.4). A strong characteristic IR peak for the CL between 2800 to 2960 cm-1 is due to the stretching vibration of C-H group of lauric acid. However, peak at 1694 cm-1 is due to the stretching vibration of C=O (Shifu et al., 1989). After ten repeated washes, the presence of these characteristic peaks validates the successful adsorption of the lauric acid. A calibration curve was used to quantify the amount of adsorbed lauric acid

104 on the surface of CNC (Fig. 5.1), which recorded 78.62 mg of lauric acid per 500 mg of dried CNC. The average particle size and analysis and zeta potential value for the

CL were found be 224±11 nm and -54±5 mV; however, it was 176±7 nm and -52±3 mV for neat CNC (fresh CNC).

Fig. 5.4. FTIR spectra of cellulose nanocrystal and lauric adsorbed CNC.

Note: Cellulose nanocrystal (CNC), lauric acid (LA) and lauric acid adsorbed CNC (CL). Highlighted peaks confirm the adsorption of lauric acid on CNC.

105

5.3.2. Stability of Pickering emulsion

5.3.2.1. Effect of dilutions on average particle size

The average particle size of the Pickering emulsions was recorded against different dilution factors, pH and storage period. Since the food matrix passes through different stages such as processing conditions, serving styles, and digestion, it is vital assuring the key attributes of the foods are maintained. A complex food system such as

Pickering emulsion, which is a physicochemically stable emulsion system; must retain its quality during processing, storage, serving as well as during initial and stomach phases of the digestion system (Wooster, 2006; Maphosa and Jideani, 2018; Bai et al.,

2019b). To understand the physicochemical stability of the Pickering emulsion, dilution was considered as one of the major factors. Dilution is a widespread practice during serving the food; for example, dilution of concentrated beverages can result in phase separation of the lipid phase. Moreover, secretion of digestive juices can weaken gel strength of the viscous foods and can also result in a phase separation of oil phase from the aqueous phase (Winuprasith et al., 2018; Bai et al., 2020b).

Results from particle size analysis revealed that the neat CNC of all the different concentrations had an average particle size of 2.23±0.14, 2.18±0.27, 2.16±0.08 and

2.21±0.11 µm for the C-0.5, C-1.0, C-1.5 and C-2.0, respectively. However, emulsion stabilized with CNC adsorbed lauric acid (CL) had significantly lower (p≤0.05) average particle size of 1.53±0.17, 1.33±0.05 and 1.23±0.09 µm (CL-1.0, CL-1.5 and

CL-2.0) except for the CL-0.5 (3.41±0.26 µm) as compared to the emulsion stabilized with neat CNC. It was assumed that the larger average particle size for the emulsion with neat CNC was due to the following two possible reasons: i) increased packing

106 density of CNC at the oil droplet interface due to increased surface pressure in the presence of NaCl, leading to increased particle diameter and ii) NaCl can increase

CNC-CNC interactions in the aqueous phase due to presence of Na+ cations resulting in bulkier and aggregated CNC particles (Bergfreund & Fischer, 2019; Bertsch et al.,

2019; Bai et al., 2019a). In contrast to the emulsion stabilized with neat CNC, smaller average particle size for the emulsion with CL may be due to a thinner packing density of CL at oil droplets interface triggered by hydrophobic-hydrophobic interaction rather

NaCl induced interaction and adsorption of CNC (Zoppe et al., 2012; Girouard et al.,

2016; Saidane et al., 2016). Results are shown in Figure 5.5, indicated that a serial dilution had a significant effect (p≤0.05) on the average particle size of the C-0.5 C-

1.0, C-1.5 and C-2.0, resulted in a gradual increase in the average particle size from

4.11±0.39 to 10.36±0.08, 2.66±0.2 to 10.27±0.06, 2.38±0.11 to 8.22±0.28, 2.21±0.07 to 7.02±0.34 µm. It was also observed that the higher dilution factors of eight (8x) and ten (10x) resulted in a significantly (p≤0.05) smaller average particle size as compared to the dilution factor of six (6x). It has been reported that a dilution of a Pickering emulsion can lead to a segregative phase separation due to the following reasons: (a) desorption of the polymer from the oil droplet interface reducing steric or electrostatic repulsion among droplets, (b) weaker gel strength of aqueous medium thereby increased droplets interaction thus coalescence/Ostwald ripening and (c) reduced surface pressure on the oil droplets due to decreased salt concentration desorption of adsorbed polymers (Wooster, 2006; Maphosa and Jideani, 2018; Bai et al., 2019a). Bai et al. (2019b) recorded a significant increase in the droplet size of the NaCl-CNC stabilized O/W emulsion during the in vitro gastrointestinal study, which may be due

107

Fig. 5.5. Effect of the dilution factors on the average particle size of emulsions.

Ad Ad Ae Af 10 Be Bf 1x Ac Ce Bc Cd 8 Cf 2x Dc 4x Dd 6 Cc De 6x

Ab Db Db Ab 8x Eb Eb Eb 4 Ea 10x Ec

Paricle (µm) size Paricle Cb Aa Aa Aa AaAa AaAaFd FbFc 2 Fa Gb GaGa GaGaGa GaGaFaEaGaGa

0 C-0.5 C-1.0 C-1.5 C-2.0 CL-0.5 CL-1.0 CL-1.5 CL-2.0 Sample types

Note: Unmodified CNC or neat CNC (C), lauric acid adsorbed CNC (CL), C-0.5 is the emulsion stabilized with 0.5% of unmodified CNC. Similarly, given numbers are the different concentrations viz., 0.5, 1.0, 1.5 and 2.0% of CNC and CL (solid basis). The upper-case letters denote significant differences (p ≤0.05) among sample types; however, small letters denote significant differences (p≤0.05) among dilutions (1x, 2x, 4x 6x 8x, and 10x), where “x” represents the number of dilutions. to the addition of a large amount of the simulated digestive juices (~60 mL/20 mL of

2% O/W emulsion) leading to the dilution of stabilized emulsion. In contrary to the effect of the dilution on C-0.5 C-1.0, C-1.5 and C-2.0, it was found that the dilution has less or no effect on the average particle size of the emulsion stabilized with CL (CL-

1.0, CL-1.5 and CL-2.0) except for the CL-0.5. It was hypothesized that the lower concentration of CL, i.e., for the CL-0.5, might not have covered the interface of oil droplets resulting in simultaneous coalescence and Ostwald ripening, thus larger average particle size. Finding inferred that the lauric acid adsorbed on the CNC facilitated the partial hydrophobicity to the CNC, resulting in a stable emulsion. Fig.

108

Fig. 5.6. Optical microscopic image of the stabilized emulsion.

Note: 1x is no dilution and 10x is ten times diluted emulsion. 0.5, 1.0, 1.5 and 2.0 are the concentration of CNC (C) and lauric acid adsorbed CNC (CL). Scale bar is 10 µm.

5.6 displays the optical microscopic images of the emulsion before (1x) and after ten dilutions (10x). It was noticed that the dilution led the coalescence/Ostwald ripening of oil droplets. This further support the findings of the effect of dilution on the increased average particle size of the O/W emulsion stabilized with NaCl-CNC. As described earlier, the dilution can disrupt the gel packing/network of a NaCl-CNC Pickering emulsion and can trigger the coalescence/Ostwald ripening of the oil droplets (Fig.

5.7). However, dilution may not have a similar effect for the emulsion stabilized with

109

Fig. 5.7. Schematic illustration (not to scale) of the stabilization of emulsion.

Note: Emulsion stabilized with unmodified CNC + NaCl (a) and lauric acid adsorbed CNC (b). An effect of dilution on the stability of emulsion stabilized with NaCl-CNC (a). Dilution breaks the CNC gel packing/network causing the coalescence/Ostwald ripening of the oil droplets.

Fig. 5.8. Macroscopic image of stabilized emulsions.

Note: Creaming stability of emulsion after ten times (10x) of dilution and a storage period of after 4 hours of storage period. Unmodified CNC (C) and lauric acid adsorbed CNC (CL) with different concentrations of 0.5, 1.0, 1.5 and 2.0% (solid basis).

110

CL due to a hydrophobic-hydrophobic interaction between oil droplets and CL.

Macroscopic images of the beta-carotene O/W emulsions are shown in Fig. 5.8 exhibit the effect of ten times dilution (10x) on the creaming stability of oil droplets after 4 hrs.

A clear phase separation for the C-0.5, C-1.0, C-1.5, C-2.0 and CL-0.5 can be seen, except for the CL-1.0, CL-1.5 and CL-2.0 inferred that the CL above 1.0 % can be a potential candidate for stable Pickering emulsion.

5.3.2.2. Effect of pH on average particle size

The effect of pH ranging from 1 to 11 on the particle size and zeta potential of the Pickering emulsions are given in Fig. 5.9a. It was observed that the pH level of above 3.0 had no significant (p>0.05) effect on the average particle size of the Pickering emulsion for both types of samples; however, a significant (p≤0.05) increase in the average particle size was recorded at the pH level 1.0. Since an extremely lower pH

+ - can lead to the protonation (H ) of the sulfate anions (-OSO3 ) present on the surface of the CNC, leading to the agglomeration of the CNC (Varanasi et al., 2018). It was also presumed that the protonation of CNC led the reduced electrostatic repulsion causing agglomeration of the CNC adsorbed oil droplets, thus flocculation. Bai et al.

(2019a) reported a significant increase in the particle size of the CNC stabilized O/W

Pickering emulsion below the pH level of 2.0. The study concluded that the increase in the particle size was caused by flocculation rather coalescence/Ostwald ripening, as confirmed by confocal laser scanning microscopic analysis of the samples. As it was hypothesized that the lower pH decreased electrostatic repulsion among the oil droplets due to protonation of the CNC adsorbed oil droplets. Results shown in Fig. 5.9b displayed a significant change in the zeta potential values of the Pickering emulsions

111

Fig. 5.9. Effect of pH on the average particle size of stabilized emulsion.

6 1 3 5 7 9 11 (a)

Ba 5

Aa Aa 4 Aa Aa Aa Aa Aa AaAa

Ab 3 Ab Ab Ab Ab Ab Ab Ab Ab Ab Ab Ab Ab Ab AbAb Ab Ab Ab Ab Ab AbAb

Particle size Particle (µm) 2 Cb Cb Cb CbCb Cb Cb Cb Cb Cb Cb Cb CbCbCb 1

0 C-0.5 C-1.0 C-1.5 C-2.0 CL-0.5 CL-1.0 CL-1.5 CL-2.0 Sample types

10 pH levels (b) 0 1 3 5 7 9 11 -10

-20 C-2.0 CL-2.0 -30

Zeta (mV) potential Zeta -40

-50

-60 Note: Emulsion stabilized with unmodified CNC (C) and lauric acid adsorbed CNC (CL) with different concentrations of 0.5, 1.0, 1.5 and 2.0% (a). The upper-case letters denote the significant difference (p ≤0.05) among sample types; however, small letters denote the significant difference (p≤0.05) among pH (1, 3, 5, 7, 9 and 11). Effect of pH on the zeta potential of the emulsion stabilized with CNC 2.0% (C-2.0) and lauric acid adsorbed CNC 2.0% (CL-2.0).

112 as affected by a decrease in pH; however, a pH of above 7.0 had no significant effect

(p>0.05). A consecutive less negative zeta potential values were recorded with a gradual decrease in the pH due to protonation of the sulfate ions, causing a charge inversion to the particles at the pH level of 1.0. This resulted in a positive zeta potential values of 9.64±0.78 mV and 10.84±1.69 mV below the isoelectric point of the CNC

(pKa = 1.9, Varanasi et al., 2018). The charge inversion of a particle either from negative to positive or positive to negative is a phenomenon of altering the electric behavior of a particle with the addition of an excessive number of the opposite ions, which leads to the overcharge, also known as charge reversal (Schubert et al., 2019).

5.3.2.3. Effect of storage on average particle size

Fig. 5.10 shows the average particle size of the Pickering emulsion stored at ambient temperature after 1, 2, 3, 5, 10, 15 and 30 days of preparation. Result displayed that the particle size for the emulsion stabilized with neat CNC (C-0.5, C-1.0, C-1.5 and C-2.0) was constant over the storage period. An insignificant (p>0.05) change in average particle size for the emulsion with neat CNC can be explained due to the following reasons: (i) NaCl increased adsorption of CNC to the oil droplet interface due to increased surface pressure, (ii) increased packing density of CNC at the oil droplet interface, and (iii) NaCl increased CNC-CNC interactions in the aqueous phase

(Bergfreund & Fischer, 2019; Bertsch et al., 2019; Bai et al., 2020a). Presence of NaCl increased the interaction of CNC-CNC due to Na+ cations in the continuous phase, increasing the gel strength of the aqueous phase. The increased gel strength of the aqueous phase can limit the mobility of the oil droplets resulting in less interaction among oil droplets, thus stable emulsion. A similar result was recorded for the emulsion

113

Fig. 5.10. Effect of storage periods on the average particle size of stabilized emulsions.

12 C-0.5 C-1.0 C-1.5 10 C-2.0 CL-0.5 8 CL-1.0

4 Particle size (µm) size Particle 2

0 0 1 2 3 5 10 15 30 Storage period (Days) Note: Emulsions stabilized with unmodified CNC (C) and lauric acid adsorbed CNC (CL) with different concentrations of 0.5, 1.0, 1.5 and 2.0% (solid basis). stabilized with CL except for the CL-0.5 and CL-1.0. It was observed that the average particle size for both samples (CL-0.5 and CL-1.0) increased significantly (p≤0.05) over the storage period. It can be reasonably assumed that the coalescence/Ostwald ripening of the partially coated or unentrapped oil droplets was caused by the Van der

Waals attraction that increased the average particle size (Bai et al., 2019a).

5.3.2.4. Effect on creaming index

The creaming index (CI) of the Pickering emulsion was estimated to analyze beta-carotene content present in the top layer of the emulsion and the total beta-carotene in the emulsion. Results shown in Figure 5.11a, displayed that the dilution had a significant (p≤0.05) increase in the CI for the emulsion stabilized with neat CNC (C-

0.5, C-1.0, C-1.5 and C-2.0), resulting in up to 90% of the CI at a dilution factor of 10x

(Fig. 5.11a). Reasonably it was due to weakening or complete loss of the gel strength

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Fig. 5.11. Creaming index of the stabilized emulsions.

100 Ab Ab Ab (a) Ab 80 Cb 1x 10x 60 Ca

Db Aa Creaming (%) index Creaming 20 Aa Ba Ba Ba Ba Ba Ba Ba 0 C-0.5 C-1.0 C-1.5 C-2.0 CL-0.5 CL-1.0 CL-1.5 CL-2.0 Sample types

100 C-0.5 C-1.0 C-1.5 C-2.0 (b) CL-0.5 CL-1.0 CL-1.5 CL-2.0 85

25 Creaming (%) index Creaming

-5 Intial 1 5 15 30 Storage period (Days)

Note: Effect of dilutions (1x and 10x) after 4 hours (a) and storage periods (b) on the creaming index. The upper-case letters denote significant differences (p≤0.05) among sample types; however, small letters denote significant differences (p≤0.05) between dilutions (1x and 10x).

115 of the continuous phase resulting coalescence/Ostwald ripening of the oil droplets led by Van der Waals force of attraction (Bai et al., 2019b). It was also presumed that the dilution caused the depletion of any adsorbed CNC from the surface of the oil droplets altering the steric and electrostatic stability of the Pickering emulsion. In contrast to the effect of dilution on C-0.5, C-1.0, C-1.5 and C-2.0, it was noticed that the dilution did not affect the CI of Pickering emulsion stabilized with CL except for the CL-0.5 and

CL-1.0. An increase in the CI for the CL-0.5 and CL-1.0 from 51.98 to 69.81% and

17.13 to 24.04% was attributed due to coalescence/Ostwald ripening/flocculation of the uncoated/partially coated oil droplets. It can be reasonably assumed due to lower concentration of the CL for the CL-0.5 and CL-1.0 as compared to the CL-1.5 and CL-

2.0. It was observed that an increase in the concentration of the CL in the emulsion gradually decreased the CI of the Pickering emulsion and so the effect of dilution (Fig

5.11a). UV-Vis spectrum of the beta-carotene from the Pickering emulsion shown in

Fig. 5.12 displayed an indistinguishable change in the absorbance of the beta-carotene of the samples except for the CL-0.5 and CL-1.0 (Fig 5.12a-b). However, a dilution factor of 10x resulted in an intense decrease in the absorbance at 452 nm for the C-0.5,

C-1.0, C-1.5 and C-2.0 Pickering emulsions (Fig 5.12c). A decreased absorbance of the beta-carotene in the emulsion indicated the creaming of the disperse phase (oil phase) from the continuous phase (aqueous phase), resulting in less beta-carotene in the continuous phase thus lower absorbance. CI for the Pickering emulsion was recorded after 1, 5, 15 and 30 days of storage period are shown in Fig 5.11b. Results indicated that the CI of the samples were not significantly different (p>0.05) over the

30 days of storage period except for the CL-0.5 and CL1.0. A sharp increase in the CI

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Fig. 5.12. UV-Vis spectra of the beta-carotene of stabilized emulsions.

3 C-0.5 C-1.0 (a) 2.5 C-1.5 C-2.0 CL-0.5 CL-1.0 2 CL-1.5 CL-2.0

1.5

Absorbance 1

0.5

0 330 380 430 480 530 Wavelength (nm)

3 C-0.5 C-1.0 (b) 2.5 C-1.5 C-2.0 CL-0.5 CL-1.0 2 CL-1.5 CL-2.0

1.5

Absorbance 1

0.5

0 330 380 430 480 530 Wavelength (nm)

117

3 C-0.5 C-1.0 (c) 2.5 C-1.5 C-2.0 CL-0.5 CL-1.0 CL-1.5 CL-2.0 2

1.5

Absorbance 1

0.5

0 330 380 430 480 530 Wavelength (nm)

3 C-0.5 C-1.0 (d) C-1.5 C-2.0 2.5 CL0.5 CL1.0 CL1.5 CL2.0 2

1.5 Absorbance 1

0.5

0 330 380 430 480 530 Wavelength (nm)

Note: Spectra of the beta-carotene from stabilized emulsions after 0 hour (a), 4 hours (b), 10 times (10x) diluted emulsion after 4 hours (c) and undiluted emulsions after 30 days of storage (d).

118 of 66.33% was recorded on the first day of storage for the CL-0.5; however, a further increase in the storage period had an insignificant (p>0.05) effect on the CI of the CL-

0.5. Since the concentration of the CL was the lowest, a quick increase in the CI for the

CL-0.5 was due to coalescence/Ostwald ripening or flocculation of uncoated oil droplets resulting in creaming and phase separation. However, an increase in the CL concentration in the emulsion had significantly (p≤0.05) lower CI over the storage periods. UV-Vis spectrum of the beta-carotene from the Pickering emulsion after 30 days of storage is given in Fig. 5.12d. Results displayed that the absorbance of the beta- carotene at 452 nm decreased drastically for the CL-0.5 after 30 days of storage.

However, as the concentration of the CL increased above 1.0 %, the absorbance of the beta-carotene in the continuous phase was unchanged. Bai et al. (2018) found that the

Pickering emulsion stabilized with a lower concentration of CNF had higher CI during the storage period.

5.3.3. Oxidative stability of emulsions

The oxidative stability of the O/W emulsion is greatly influenced due to the presence of lipid hydroperoxides and the transition metals in the emulsion system.

Lipid hydroperoxides on the surface of the oil droplets and the presence of transition metals in the continuous phase are the major cause of the lipid oxidation (Alamed et al., 2006). Malondialdehyde, an end product of the deterioration of the lipid hydroperoxides and other primary and secondary lipid peroxidation products (Janero,

1990) is measured by TBARS analysis (Alamed et al., 2006). The oxidative stability of the Pickering emulsion at ambient temperature over the storage period of 1, 2, 3, 5,

10, 15, 20 and 30 days is shown in Fig. 5.13. Results indicated that the TBARS

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Fig. 5.13. Thiobarbituric acid reactive substances (TBARS) values of emulsions.

1 Control C-2.0 CL-2.0 (a)

0.8

0.6

0.4

0.2 TBARS (µmol/mL emulsion) (µmol/mL TBARS 0 0 1 2 3 4 5 Storage periods (Days)

35 Control (b) ) 30 C-2.0 25 CL-2.0

20 mL emulsion mL 15

5 TBARS (µmol/ TBARS 0 10 15 20 25 30 Storage periods (Days)

Note: Control emulsion contains gum Arabic, however, other emulsions contain unmodified CNC (C) and lauric acid adsorbed CNC (CL). The TBARS values were recorded over the storage periods of 1 to 5 days (a) and from 10 to 30 days (b). concentration for the Pickering emulsion stabilized with neat CNC (C-0.5, C-1.0, C-

1.5 and C-2.0) and CL (CL-0.5, CL-1.0, CL-1.5 and CL-2.0) increased gradually over

120 the storage period, recorded a significantly (p≤0.05) higher concentration of TBARS as compared to the control sample. As not anticipated, a higher TBARS concentration for the Pickering emulsions with CNC later conformed with mineral composition analysis of the CNC (Table 5.1). Mineral composition analysis of the CNC revealed the presence of a wide variety of transition metals. The higher concentration of the

TBARS for the Pickering emulsion stabilized with CNC may have been due to catalytic effect of the associated higher concentration of the transition metals that induced the oxidation/deterioration of hydroperoxides on the surface of the lipid droplets producing malondialdehyde thus higher TBARS value. Since most of the processing tools involved in the production of CNC are metallic (Nanocellulose Pilot Plant, NFL,

USDA). We assumed that the presence of transition metals in the CNC was the traces from the involved processing equipment.

Table 5.1. Mineral contents of cellulose nanocrystals (CNC). Superscripts are the unit values of minerals per gram of CNC.

Note: Results are mean ± standard deviation of triplicate (n=3). α – values in nanogram (ng/g); β – values are in microgram (µg/g) and γ – values in milligram (mg/g).

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5.3.4. Beta-carotene stability

Since beta-carotene possesses a large number of conjugated double bonds and is very susceptible to oxidative degradation. These conjugated bonds trigger the oxidative degradation of beta-carotene similar to transition metals, light, heat, singlet oxygen, etc. (Boon et al., 2010). The stability of beta-carotene during storage in the

Pickering emulsions is shown in Fig. 5.14. It was observed that beta-carotene content for the control emulsion was significantly higher as compared to the Pickering emulsion stabilized with neat CNC and CL. However, no significant (p≤0.05) difference was observed between emulsion stabilized with neat CNC and CL. Since the

CNC, CL and Gum Arabic were only differences among the composition of the emulsions. It was assumed that the loss of beta-carotene for the C-2.0 and CL-2.0 was simply assumed due to the presence transition metals such as Fe, which is a primary cause of the oxidative degradation of the beta-carotene, resulting formation of carotene cation radical and ferrous iron (Boon et al., 2010). Several studies have been conducted to ensure the stability of beta-carotene O/W emulsion using a vast number of natural and synthetic antioxidants. Among them, the metal-chelating antioxidant, i.e., EDTA, has shown very promising results preventing the degradation of beta-carotene (Boon et al., 2010; Qian et al., 2012 and Liu et al., 2015).

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Fig. 5.14. Stability of beta-carotene during storage periods.

9 carotene (mg/100 mL) carotene

- 7 Beta

Control C-2.0 CL-2.0

3 0 1 2 3 5 10 15 20 30

Storage periods (Days)

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5.4. Conclusion

Adsorption of lauric acid with cellulose nanocrystals (CL) resulted in a physically stable beta-carotene O/W Pickering emulsion at different dilution factors, pH levels, and storage periods. However, neat CNC (no lauric acid) maintained the physical stability of the emulsion at different pH and during storage nonetheless resulted in a significant (p≤0.05) increase in average particle size of the while diluted.

Results displayed that a gradual increase in the concentration of CL from 0.5 to 2.0 had better physical stabilization activity for the Pickering emulsion, resulting in an average particle size of 3.41±0.26, 1.53±0.17, 1.33±0.05 and 1.23±0.09 µm (CL-0.5, CL-1.0,

CL-1.5 and CL-2.0), respectively. Dilution resulted in a significant (p≤0.05) increase in average particle size of the C-0.5, C-1.0, C-1.5 and C-2.0; however no or a moderate increase was recorded for the CL-0.5, CL-1.0, CL-1.5 and CL-2.0. This was assumed due to better lipophilic-lipophilic balance between oil droplets and lauric acid present on the surface of CL, resulting in a stable emulsion. An intriguing result was noticed during the storage of the Pickering emulsions. Results indicated that emulsion stabilized with neat CNC and CL had significantly (p≤0.05) higher TBARS concentration 28.19 and 29.28 µmol (C-2.0 and CL-2.0) and lower beta-carotene 4.33 and 4.02 mg/100 mL (C-2.0 and CL-2.0) as compared to the control emulsion (9.28

µmol and 6.53 mg/100 mL) stabilized with Gum Arabic. This study thus demonstrates the potential application of adsorption of lauric acid modulating the stability of the O/W emulsion. The study also highlights the limitation of the CNC and its consequential cause to the lipid food system due to the presence of transition metals.

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CHAPTER 6

DEVELOPMENT OF O/W PICKERING EMULSIONS USING LAURIC ACID-

ADSORBED CNC: AN IN VITRO GASTROINTESTINAL TRACT STUDY

6.1. Introduction Cellulose nanocrystals (CNC), also known as crystalline nanocellulose, nanocrystalline cellulose, or cellulose nanowhiskers, is a derivative of cellulose, which is the most common type of polysaccharide available on the earth. CNC is a linear structure of anhydro D-glucose units condensed through a β-1,4 glycosidic bond

(García, Gandini, Labidi, Belgacem, & Bras, 2016). Morphologically, CNC is a rod or needle-shaped nanoparticle with a dimension of 150-200 nm long and 5-15 nm wide

(Cheng et al., 2015) and possesses up to 99% crystallinity (Abiaziem et al., 2019). The properties of CNC include nano-scaled dimension, high crystallinity, surface hydroxyl groups (O-H) (Natterodt et al., 2017), excellent dispersibility into an aqueous medium

- (Cheng et al., 2015), high charge potential due to sulfate ions (-SO3 ) (Prathapan et al.,

2016) and non-toxicity (Seabra et al., 2018), thus offering an excellent opportunity to explore its potential applications. There have been several studies utilizing CNC in foods (Hedjazi and Razavi, 2018; Aswathanarayan, and Vittal, 2019) and pharmaceuticals (Ngwabebhoh et al., 2018; Low et al., 2019).

In the last few decades, the market demands have shifted towards healthy and low- calorie foods due to increased public health concern and development of nutraceutical and functional foods (Siro et al., 2008; Sharif and Zahid, 2018). Developing nutraceuticals and functional foods through the Pickering emulsion process is one of

125 the more common techniques that are being used, hence the exploration of novel and unique coating/stabilizing materials has increased (Dhakane et al., 2017; Hedjazi and

Razavi, 2018; Sharif and Zahid, 2018; Winuprasith et al., 2018). A Pickering emulsion is a type of emulsion system that is stabilized by solid colloid particles by adsorbing them onto the interface of two immiscible phases (Pickering, 1907). Since CNC possesses unique characteristics of being an excellent stabilizer/emulsifier due to its aforesaid physicochemical properties (Kalashnikova et al., 2012; Zoppe et al., 2012;

Prathapan et al., 2016; Ujala et al., 2016; Meirelles et al., 2020), several researchers have investigated the effect of CNC on the physicochemical stability of an oil-in-water

(O/W) type emulsion (Rayner et al., 2014; Hedjazi and Razavi, 2018; Sharif and Zahid,

2018; Aswathanarayan, and Vittal, 2019). CNC in a Pickering emulsion system adsorbed onto the interface of oil droplets, creating either electrostatic (Prathapan et al., 2016) or steric repulsion (Zoppe et al., 2012) among the oil droplets that leads to a

- stable emulsion system. Electrostatic stability is due to the presence -SO3 groups on the surface of CNC; however, a steric repulsion is due to adsorbed CNC that limits the coalescence/Ostwald ripening among the droplets. Adsorption of CNC onto the interface of oil droplets may not be an easy target due to the hydrophilic nature of CNC due to the presence of a vast number of O-H groups. However, several researchers have found that the addition of a certain amount of NaCl (0.06%) facilitated the adsorption of CNC particles onto the interface of oil droplets due to increased surface pressure

(Bergfreund and Fischer 2019; and Bertsch et al., 2019). The addition of NaCl also increased the CNC-CNC interaction due to the counterion effect of sodium ions (Na+) altering the rheological properties, e.g., increasing gel strength of aqueous phase

126

(higher viscosity). The increased gel strength of the continuous phase restricts the mobility of the free oil droplets (no CNC on the interface), resulting in less interaction among oil droplets, thus forming a stable emulsion (Aaen et al., 2019).

A study conducted by Bai et al. (2019a) has reported a significant increase in the oil droplet size in the O/W Pickering emulsion stabilized with CNC while investigating the in vitro gastrointestinal tract (GIT) fate of the Pickering emulsion. Results showed an average increase of up to 30% in the diameter of the oil droplet at the simulated stomach phase; nonetheless, no agglomeration of oil droplets was observed by confocal laser scanning microscopic (CLSM) analysis. However, CLSM images recorded at the small intestinal phase of the study depicted a clear agglomeration of the oil droplets, thus increasing the diameter of the oil droplets. It is assumed that agglomeration of oil droplets could have been due to a gradual dilution of the Pickering emulsion with simulated digestive juices and the addition of counterions (Wooster, 2006; Maphosa and Jideani, 2018). It has been reported that dilution can lead to a phase separation of the stable Pickering emulsion due to the following reasons: (a) reduced surface pressure, thus desorption of polymers from the interface of oil droplet plunging steric or electrostatic repulsion among droplets and (b) weaker gel strength of the aqueous medium thereby increasing droplets’ interaction leading to coalescence/Ostwald ripening (Wooster, 2006; Maphosa and Jideani, 2018; Yokota et al., 2019).

CNC with hydrophobic or amphiphilic surface properties can be effectively utilized in creating a stable Pickering emulsion (Zoppe et al., 2012; Girouard et al.,

2016; Saidane et al., 2016). However, the sustainability of the chemical modification of CNC utilizing non-food grade organic chemicals is a serious concern (Chemat et al.,

127

2019; Spinelli et al., 2019). Prior work suggested that adsorption of amphiphilic molecules on the interface of hydrophilic molecules can easily be achieved, resulting in a hydrophobic surface. The adsorption of amphiphilic molecules on the interface of the hydrophilic surface takes place due to the ligand exchange phenomenon in which the polar group of the amphiphilic molecules replaces the aqueous molecules from the interface of hydrophilic molecules and facilitating the adsorption (Paleos, 1969 and

Ulrich et al., 1988).

Additionally, the developed Pickering emulsion can be used for the stabilization, fortification, and delivery of a wide variety of lipophilic nutraceuticals

(Dhakane et al., 2017; Hedjazi and Razavi, 2018; Sharif and Zahid, 2018; Winuprasith et al., 2018). However, it is crucial to understand the physicochemical stability and fate of the digestion of developed nutraceutical-loaded Pickering emulsions for their potential applications.

Thus, in this work, a medium-chain saturated fatty acid named lauric acid

(C12:0) was adsorbed onto the surface of CNC and lauric acid adsorbed CNC (CL) was used as an emulsifier to create O/W Pickering emulsion. Beta-carotene was used as model nutraceutical. Developed Pickering emulsion was investigated for the static in vitro GIT study. Different concentrations of dried CL viz., 0.5, 1.0, 1.5 and 2.0% (w/v) was utilized as emulsifier. Stabilized Pickering emulsion was then passed through various stages of the GIT such as simulated mouth phase, stomach, and small intestinal phase. Characteristics of the Pickering emulsion such as FTIR, TEM, average particle size, zeta potential, CLSM analysis, creaming index lipid digestion, and beta-carotene bioaccessibility were recorded.

128

6.2. Materials and Methods

CNC with a total solid content of 11.5 to 12.5% was procured from the PDC,

University of Maine. The obtained CNC had an average particle length of 150-200 nm and 5-20 nm wide and contained up to two percent of the surface sulfate group. Cooking grade canola oil was purchased from the local supermarket of Bangor, Maine (Great

Value, Walmart). Lauric acid flakes (>99% pure) were purchased from Acros

Organics/Fisher Scientific, USA. Enzymes used for in vitro gastrointestinal study such as α-amylase (from Aspergillus oryzae), pepsin (from porcine gastric mucosa: ~2500 units/mg), lipase type II (from the porcine pancreas), mucin-type III (from the porcine stomach), porcine bile extracts, uric acid (≥99% crystalline), beta-carotene (>99%) and

Nile red (a fluorescence marker) were purchased from Sigma Aldrich, USA. Other analytical grade solvents were purchased from Acros Organics/Fisher Scientific, USA.

6.2.1. Adsorption of lauric acid with CNC

Adsorption of lauric acid with CNC was performed dissolving 20 grams of lauric acid flakes into 400 mL of absolute ethanol. Subsequently, 100 grams of CNC suspension (11.5 to 12.5%) was mixed to the solution stirring at 500 rpm on a digital magnetic stirrer plate (Isotemp™, Fisherbrand, USA). The mixture was stirred continuously overnight at 100 rpm followed by ten times centrifugation at 21,800x g for 10 minutes at room temperature using a super speed centrifuge (Beckman Coulter

Avanti J-E Floor, CA, USA). Obtained pallet was washed several times with the subsequent addition of ethanol to remove the unabsorbed lauric acid. Washed pallet was then dried under vacuum at 60 °C overnight and was considered to be lauric acid-

129 adsorbed CNC (CL). CL was later ground to a fine powder manually using mortar and pestle and was stored into the airtight LDPE Ziplock bag at room temperature for further use. The amount of adsorbed lauric acid was subsequently quantified on FTIR

(ABB, FTLA 2000) using a blend of different concentrations of lauric acid with dried

CNC viz., 10, 20, 30, 50, 75, 100, 150, 200 and 250 mg of lauric acid per 500 mg of

CNC (dry basis). The FTIR spectra were recorded as relative percent transmittance

(%T), which was later converted into absorbance following the equation developed by

Toft et al. (1993):

퐴푏푠표푟푏푎푛푐푒 = 2 − 푙표푔 (%푇)

The obtained absorbance values were plotted to create a calibration curve to calculate the amount of lauric acid adsorbed.

The morphology of the unmodified CNC (neat CNC) and CL was characterized by transmission electron microscopy (TEM CM10, Philips/FEI, Hillsboro, Oregon,

USA). The 100x diluted samples were prepared using the drop-casting method on a negatively stained copper grid fixed on a carbon film. After 15 minutes, images were collected under a vacuum at 200 kV.

6.2.2. Preparation of emulsions

An oil-in-water (O/W) type Pickering emulsion was prepared by mixing 10 grams of beta-carotene dissolved canola oil (1 mg/100 mL) into 90 grams of the aqueous phase (CL dispersed water). The CL dispersed aqueous phase contained four different concentrations of CL viz., 0.5, 1.0, 1.5 and 2.0%. Resulting mixtures were coded as

CL-0.5, CL-1.0, CL-1.5 and CL-2.0. Similarly, 10 grams of beta-carotene dissolved

130 canola oil was mixed into another 90 grams of aqueous phase (neat CNC dispersed water). In this case aqueous phase contained four different concentration of neat CNC viz., 0.5, 1.0 1.5, and 2.0% and coded as C-0.5, C-1.0, C-1.5, and C-2.0. Neat CNC with different concentrations were used to compare the stabilization efficacy of CL on the stability Pickering emulsion.

Oil and aqueous phase as mentioned above were mixed using a homogenizer

(Ultra-Turrax, IKA S 18 N - 19 G Dispersing Element) at 10,000 rpm for 2 minutes and resulting mixture was considered as coarse emulsion. 0.06 g of NaCl was added to each C-0.5, C-1.0, C-1.5, and C-2.0 as described by Bai et al. (2018); however, no NaCl was added to the CL stabilized emulsion. To obtain a fine emulsion, resulting coarse emulsion was subsequently sonicated for 5 minutes using an ultrasonic dismembrator

(500W, 20 kHz, Fisherbrand™ Model 505, USA) with 0.75-inch titanium alloy microtip probe. Sonification was performed at 40 percent amplitude, maintaining the

ON and OFF-pulse time at the interval of 15 and 5 seconds (Mehmood et al., 2017).

Resulting Pickering emulsions were used for in vitro GIT study.

6.2.3. In vitro gastrointestinal tract (GIT) study

A static in vitro GIT of the Pickering emulsions was performed following the protocol of Bai et al. (2019a) with a small adjustment. All the simulated digestive solutions were prepared, as described in Chapter 3. Simulated digestion solutions were prepared and preheated at 37 °C just before use. The reaction was performed in a 100 ml amber-colored Erlenmeyer flask to avoid the effect of light on the degradation of beta-carotene.

131

6.2.3.1. Initial phase of the in vitro GIT

Before starting the in vitro GIT experiment, an initial sample was prepared by taking 40 mL of Pickering emulsion containing 2% oil (w/w) into the flask. Samples were incubated in a shaking water bath (JULABO, SW22, Julabo Inc., USA) at 37 °C for 2 minutes.

6.2.3.2. Simulated saliva juice (SSJ) phase of the in vitro GIT

The SSJ phase of the in vitro GIT, also known as simulated mouth phase, was performed by withdrawing 20 mL of a sample from the initial phase followed by adding

20 mL of preheated (37 °C) SSJ. The pH of the mixture was maintained at 6.8±0.2 using 0.1 N HCl or 0.1 M NaOH solutions. The reaction was allowed to process for 5 minutes at 37 °C with a continuous shaking speed of 100 rpm.

6.2.3.3. Simulated gastric juice (SGJ) phase of the in vitro GIT

Similar to the SSJ phase, the SGJ or simulated stomach phase of the in vitro

GIT was performed by withdrawing a 20 mL aliquot of the sample from the SSJ phase and mixing it with 20 mL of preheated SGJ. The pH of the reaction medium was maintained at 2.0 using 0.1 N HCl, and the digestion was allowed for 2 hours with continuous shaking at 100 rpm.

6.2.3.4. Small-intestinal juice (SIJ) phase of the in vitro GIT

After 2 hours of digestion, a 20 mL of sample was withdrawn from the SGJ phase into a separate flask, and pH was immediately brought to 7.0 ±0.2 using 0.1 M

NaOH. Subsequently, 1.5 mL of SIJ was added to the flask, followed by the addition of 3.5 mL of bile salt and 2.5 mL of lipase solutions (Bai et al., 2019a). Since a pH

132 titration method was used to measure the lipid digestion during the SIJ phase of the in vitro GIT, the pH of the reaction medium was checked carefully after the addition of reaction solutions. The reaction was further allowed to process for 2 hours at 37 °C with continuous shaking at 100 rpm. A replication of three for each sample was performed. Obtained samples were subsequently investigated for their attributes, as given below.

6.2.4. Particle size and zeta potential analysis

The average particle size and zeta potential of the Pickering emulsions were analyzed on a DLS zeta-sizer (Zetasizer Malvern 3000 HSA). Analysis was performed following the method of Zhang et al. (2019). In brief, samples from each phase of the in vitro GIT study were diluted with distilled water to prevent the multiple scattering effects during analysis. The refractive index for the canola oil was fixed at 1.465

(Przybylski, 2001) and 1.330 for water (Zhang et al., 2019).

6.2.5. Creaming index

Creaming index of the samples collected from different phases of the in vitro

GIT was analyzed following the method of Varka et al. (2010) with some modifications. In brief, the amount of the beta-carotene in the emulsion was measured using a multi-plate reader UV-Vis spectrophotometer (BioTek Eon, Vermont, USA) at

452 nm. One mL of sample was withdrawn from the separated phase (top layer) and the continuous phase (bottom phase) separately into 10 mL of n-hexane, then vortexed for 1 minute. The solutions were then stored at 4 ℃ until a clear layer of the aqueous phase and n-hexane appeared. A 200 µl of the fraction from the top layer was placed

133 into 96 well microplates and absorbance was recorded at 452 nm. The percent creaming index was calculated using the following equation:

퐴푏푠푡표푝 × 100 퐴푏푠푡표푡푎푙

퐴푏푠푡표푡푎푙 = 퐴푏푠푡표푝 + 퐴푏푠푐표푛푡𝑖푛푢표푢푠 푝ℎ푎푠푒

Where Abstop is the absorbance of beta-carotene in the separated oil phase, Abscontinuous phase is the absorbance of beta-carotene present in the bottom phase.

6.2.6. Confocal laser scanning microscopic (CLSM) analysis

CLSM analysis of the Pickering emulsion obtained each phase of the in vitro

GIT study was performed staining the oil droplets with the Nile red (1 mg/1mL ethanol) (Winuprasith et al., 2018). A 50 µL of stained emulsion sample was loaded on a glass slide (75 x 26 x 1 mm) and covered with a glass coverslip. Images were collected simultaneously on a high-resolution CLSM (Leica TCS SP instrument). An excitation and emission spectra of 545 and 575 nm were used for scanning using a

20X magnification lens.

6.2.7. Determination of lipid digestion

The digestion of oil droplets of the Pickering emulsion was determined by quantifying available free fatty acids (%) at the end of the SIJ phase of the in vitro GIT.

A pH titration method was used to quantify the free fatty acids (FFAs), as suggested by Winuprasith et al. (2018). The obtained sample was titrated with 0.1 M NaOH until a pH of 7.0 was achieved. Total FFAs (%) were then calculated following the number of moles of NaOH required (titer value) to neutralize the released FFAs.

134

푉푁푎푂퐻 × 푚푁푎푂퐻 × 푀퐿𝑖푝𝑖푑 퐹퐹퐴 (%) = 푊퐿𝑖푝𝑖푑 × 2

Where VNaOH is the consumed volume of NaOH (mL) during the titration, mNaOH is the molarity of the NaOH used (0.1 M), MLipid is the molecular weight of the canola oil

(~876.6 g/mole) and WLipid is the total weight of lipid initially present in the reaction vessel.

6.2.8. Beta-carotene bioaccessibility

The bioaccessibility of the beta-carotene in the Pickering emulsion was estimated by the method of Winuprasith et al. (2018). The bioaccessibility is defined as the percent of beta-carotene available in the form of mixed micelle as compared to the digesta (total digest resulting after the small intestine phase) for its absorption into the small intestine (Marze, Meynier, & Anton, 2013). A 5 mL sample of the Pickering emulsions collected from the SIJ phase was transferred into a 15 mL centrifuge tube and centrifuged at 1,135x g for 50 min at 20 ºC. The top clear yellowish layer from the centrifuge tube was considered as the mixed micelle from which an aliquot of 1 mL was mixed with 5 mL of isooctane and ethyl alcohol (1:3) to extract the beta-carotene.

Mixture was then centrifuged again for 15 minutes at 205x g. Absorbance of the beta- carotene from the supernatant was recorded at 452 nm on a multi-plate reader UV-Vis spectrophotometer ((BioTek Eon, Vermont, U.S.A)). The bioaccessibility was then calculated using the equations below:

퐶 퐵푖표푎푐푐푒푠푠푖푏푖푙푖푡푦 (%) = ( 푀𝑖푐푒푙푙푒 ) × 100 퐶퐷𝑖𝑔푒푠푡푎

135

Here, CMicelle is the beta-carotene in the mixed micelle phase; however, CDigesta is the total beta-carotene in the digesta obtained at the end of the SIJ phase of the in vitro

GIT.

6.2.9. Statistical analysis

Date collected in triplicate were analyzed using a Tukey’s (HSD) post hoc test on SAS® Studio (Version 9.4, University edition) to identify the significant differences among the mean values. Analysis of Variance (ANOVA) was used to determine a significant effect among the sample types and the different stages of the GIT at the level of p≤0.05.

6.3. Results and Discussion

6.3.1. Characterization of lauric acid adsorbed CNC (CL)

Fourier transforms infrared (FTIR) spectra shown in Fig. 6.1 confirmed the adsorption of lauric acid onto the surface of CNC. The characteristic peak of the C-H group of lauric acid between 2800 to 2960 cm-1 confirmed the adsorption of lauric acid.

Moreover, the peak at 1694 cm-1 is due to the stretching vibration of the C=O group of the lauric acid (Shifu et al., 1989). Using a calibration curve plotted against different concentrations of the lauric acid, the amount of lauric acid adsorbed onto the surface of CNC was quantified as 78.62 mg/500 mg of dried CNC.

136

Fig. 6.1. FTIR spectra of cellulose nanocrystal (CNC), lauric acid (LA) and lauric acid adsorbed CNC (CL).

6.3.2. Morphology of CNC and CL

The structure of CNC and CL was characterized by transmission electron microscopy (TEM). TEM images are shown in Fig. 6.2 clearly showed that the CNC particles had rod/spindle-shaped and were uniformly distributed. A previous study conducted by Bai et al. (2019a) has reported the rod-like structure of the neat CNC recorded on AFM analysis; however, it is important to understand that the morphology and diameter of the CNC may vary depending upon the type of analytical tools used such as TEM, AFM or SEM. Data collected on the Zeta-sizer showed the average

137

Fig. 6.2. Transmission Electron Microscopic (TEM) image of CNC (a) and lauric acid adsorbed CNC (b)

particle sizes for the CNC and CL were 176±7 and 224±11 nm; however, the zeta potential values were -52±3 and -54±5 mV. Larger average particle size for CL can be due to the adsorption of lauric acid on the surface of CNC, thereby widening the diameter.

6.3.4. In vitro GIT fate of emulsions

6.3.3.1. Initial phase

Initially, the Pickering emulsion was diluted to 2% and incubated for 2 minutes at 37 °C prior to investigating the fate of Pickering emulsion under GIT conditions.

Data shown in Fig. 6.3, revealed that the average particle size of the Pickering emulsion stabilized with CL i.e., CL-1.0, CL-1.5 and CL-2.0 had significantly (p≤0.05) smaller average particle sizes of 1.52±0.15, 1.31±0.03, and 1.21±0.22 µm except for the CL-

0.5 (3.82±0.17 µm) as compared to the emulsion stabilized with neat CNC, 3.14±0.09,

2.46±0.15, 2.18±0.22 and 2.21±0.08 µm (C-0.5, C-1.0, C-1.5 and C-2.0). It has been

138

Fig. 6.3. Effect of in vitro gastrointestinal tract (GIT) phases on average particle size of the emulsions.

Note: Emulsion stabilized with different concentration of unmodified CNC (i.e., C-0.5, where C stand for neat CNC and 0.5 is the concentration) and lauric acid adsorbed CNC (i.e., CL-0.5, where CL stand for lauric adsorbed CNC and 0.5 is the concentration). The upper-case letters denote significant differences (p ≤0.05) among sample types; however, small letters denote significant differences (p≤0.05) among in vitro gastrointestinal tract (GIT) phases. SSJ - simulated saliva juice, SGJ - simulated gastric juice and SIJ – simulated small-intestinal juice. mentioned in Chapter 5 that that larger mean particle size for the C-0.5, C-1.0, C-1.5 and C-2.0 was due to the following reasons: firstly, increased packing density of CNC at the oil droplet interface due to increased surface pressure in the presence of NaCl leading to increased particle diameter and secondly, NaCl can increases CNC-CNC interactions in the aqueous phase due to presence of Na+ cations resulting bulkier and aggregated CNC particles (Bergfreund & Fischer, 2019 and Bertsch et al., 2019).

However, smaller mean particle sizes for the CL-1.0, CL-1.5 and CL-2.0 can be due to

139 thinner packing density of CL at the oil droplets’ interface facilitated by hydrophobic- hydrophobic interaction rather NaCl-induced interaction and adsorption onto the oil interface (Zoppe et al., 2012; Girouard et al., 2016; Saidane et al. ,2016).

The data shown in Fig 6.4 are the average zeta potential values of the Pickering emulsion. Results revealed that the increase in the concentration of CNC and CL had no significant effect (p>0.05) on the zeta potential of the Pickering emulsion. However, a significant difference was observed between the Pickering emulsion with CNC and

CL. A comparatively higher zeta potential value of -53.53±5.39 to -50.29±2.11 mV was recorded for the Pickering emulsion with CL than for the zeta potential value of the Pickering emulsion with CNC, which ranged from -47.41±2.04 to -48.32±4.41 mV.

Values were comparable with a zeta potential of Pickering emulsion stabilized by CNC that were reported by Bai et al. (2019a). A higher zeta potential value for the Pickering emulsion with CL was presumed due to the presence of lauric acid on the surface of

CNC.

6.3.3.2. Simulated saliva juice (SSJ) phase

Incubated Pickering emulsions from the initial stage were exposed to the SSJ phase of the GIT. Average particle size data shown in Fig. 6.3 displayed an insignificant (p≥0.05) increase in the average particle size of Pickering emulsion with

CNC from the initial phase to the SSJ phase. A similar trend was observed for the

Pickering emulsion with CL, resulting in an insignificant increase in the average particle size. Results agreed with the findings of Bai et al. (2019a), who reported a negligible increase in the mean particle size of Pickering emulsion with CNC.

140

Fig. 6.4. Effect of in vitro gastrointestinal tract (GIT) phases on zeta-potential of emulsions.

GIT Phases Initial SSJ SGJ SIJ

10 Ac Ac Ac Ac Ac Ac Ac Ac 0

-10

-20 Bd Bd Bd -30 Bd Ab Ab Ab Ab Ab Ab Zeta (mV) potential Zeta -40 Ab Ab C-0.5 C-1.0 C-1.5 Bb Bb Bb C-2.0 CL-0.5 CL-1.0 Aa Aa Bb -50 Aa Ba Aa CL-1.5 CL-2.0 Ba Ba Ba -60

Note: Emulsions stabilized with CNC (i.e., C-0.5, where C stand for neat CNC and 0.5 is the concentration) and lauric acid adsorbed CNC (i.e., CL-0.5, where CL stand for lauric adsorbed CNC and 0.5 is the concentration). The upper-case letters denote significant differences (p ≤0.05) among sample types; however, small letters denote significant differences (p≤0.05) among in-vitro gastrointestinal tract (GIT) phases. SSJ - simulated saliva juice, SGJ - simulated gastric juice and SIJ – simulated small- intestinal juice.

However, Winuprasith et al. (2018) reported a slight increase in the mean particle size of the Pickering emulsion stabilized with whey protein isolates (WPI). An increase in the mean particle size was hypothesized due to the flocculation of lipid droplets instigated by deposition of anionic proteins of SSJ, i.e., mucin on the surface of lipid droplets initiating the bridge flocculation (Sarkar et al., 2009). It was observed that the

C-0.5 and CL-0.5 had a comparably larger average particle size (p≤0.05) than the other types of Pickering emulsions. CLSM images in Fig. 6.5 depicted that the larger particle

141

Fig. 6.5. Confocal laser scanning microscopic images of emulsions.

Note: Unmodified CNC and lauric acid adsorbed CNC stabilized (concentration = 0.5, 1.0, 1.5 and 2.0%) emulsions. Images shows the effect of in vitro gastrointestinal tract stages on the oil droplet behavior. Scale bar is 20 µm. SSJ - simulated saliva juice, SGJ - simulated gastric juice and SIJ – simulated small-intestinal juice. size for the C-0.5 and CL-0.5 could be due to partial flocculation of the oil droplets. A similar result was observed by Winuprasith et al. (2018) who reported flocculation of some WPI stabilized lipid droplets at the mouth stage of the GIT. Results shown in Fig.

6.4 displayed a significant (p≤0.05) slight decrease in the average zeta potential of the

Pickering emulsions. It was observed that the negative zeta potential of the particles, which was recorded -47.41±2.04 to -48.32±4.41 mV at the initial phase, dropped significantly (p≤0.05) in the range between to -34-42±3.18 to -37.42±2.05 mV for the

Pickering emulsion with neat CNC. A similar trend was observed for the Pickering emulsion with CL, which initially ranged from -53.53±5.39 to -50.29±2.11 mV declined to -44.16±1.85 to 46.42±5.21 mV. A decline in the zeta potential for the particle in the SSJ phase can be assumed due to the electrostatic screening effect of 142 counterions present in the saliva juice (Israelachvili, 2011). Additionally, adsorption of the anionic polymer from the saliva juice components i.e., mucin, can also decrease the zeta potential value by masking the surface potential of the particles (Singh & Sarkar,

2011; van Aken et al., 2011).

6.3.3.3. Simulated gastric juice (SGJ) phase

A considerable increase in the average particle size of the Pickering emulsion was recorded at the end of the SGJ phase. From Fig. 6.3, it can be seen that the average particle increased significantly (P≤0.05) by approximately 115% for the C-0.5 and by approximately 140% for the C-1.0, C-1.5 and C-2.0, resulting up to an average particle size of 7.26±0.23 (C-0.5), 6.23±0.32 (C-1.0), 6.34±0.14 (C-1.5) and 4.95± 0.08 nm (C-

2.0), respectively. A similar effect of the SGJ phase was observed on the average particle size of the Pickering emulsion stabilized with CL, which accounted for an increase by approximately 140% for the CL-0.5 and CL-1.0, however, by approximately 80% for the CL-1.5 and CL-2.0. Resulting an average particle size of

8.78±0.33 (CL-0.5), 4.04±0.05 (CL-1.0), 2.32±0.16 (CL-1.5) and 2.44±0.23 nm (CL-

2.0), respectively. Bai et al. (2019a) has reported a slight increase in the mean particle size of the Pickering emulsion stabilized with CNC at the stomach phase of the GIT, recorded an increase by approximately 30%; however, a study conducted by

Winuprasith et al. (2018) reported an insignificant effect of the stomach phase on the average particle size of the Pickering emulsion stabilized with CNF.

An increase in the average particle of the Pickering emulsion with CNC viz.,

C-0.5, C-1.0, C-1.5 and C-2.0 was hypothesized due to following three phenomena: firstly, a lower pH of the stomach phase can lead to the protonation (H+) of the sulfate

143

- anions (OSO3 ) present on the surface of the CNC leading to the agglomeration of the

CNC (Varanasi et al., 2018) thus increased average particle size. Secondly, the protonation of CNC present on the interface of lipid droplets can lead to a reduced electrostatic repulsion among the lipid droplets, thus causing flocculation of the lipid droplets. This effect can also be seen from the CLMS results of Bai et al. (2019b), who have reported the flocculation of lipid droplets stabilized with CNC at pH 2.0. The third reason for an increased average particle size of the Pickering emulsion is destabilization of gel strength (3D network) of the continuous phase (aqueous phase) due to gradual dilution, leading to the weaker gel strength. The weaker gel strength allows lipid droplets to interact with other lipid droplets with and coalesce or flocculate. It has been reported that the presence of NaCl in unmodified CNC stabilized Pickering emulsion increases the gel strength of the continuous phase, limiting the mobility and interaction of lipid droplets, therefore stable emulsion (Aaen et al., 2019). Indeed, CLMS images displayed a large amount of flocculation and coalescence of lipid droplets for the

Pickering emulsion with CNC, i.e., C-0.5, C-1.0, C-1.5 and C-2.0 at SGJ phase of the

GIT (Fig. 6.5). However, a similar effect of the SGJ phase on the Pickering emulsion with CL i.e., CL-0.5 and CL-1.0, was also noticed. From the CLSM images, it was noticed that a gradual increase in the concentration of CL from 0.5 to 2.0 maintained the stability of Pickering emulsion against the SGJ phase. This can simply be assumed that a higher concentration of CL can facilitate a complete coverage of a lipid droplet’s interface. Moreover, lauric acid adsorbed on CL developed an amphiphilic surface, creating a Pickering emulsion that was stable against dilution and lower pH (details in schematic diagram Fig. 6.6).

144

Fig. 6.6. Schematic illustration of fate of emulsions in small intestine (not to scale).

Results from the zeta potential analysis displayed a significant (p≤0.05) decline in the zeta potential of the Pickering emulsions after the SGJ phase of the GIT. Zeta potential values were changed from negative to slightly positive zeta potential (Fig.

6.4). A change in zeta potential from negative to positive can be due to the charge reversal phenomenon triggered by lower stomach pH (2.0), resulting in protonation of the sulfate ions present on the surface of CNC. The charge reversal or charge inversion, either from negative to positive or positive to negative, is a phenomenon of altering the electric behavior of a particle with the addition of an excessive number of the opposite ions, which leads to the overcharge (Schubert et al., 2019). Charge reversal occurs below or above the isoelectric point of the CNC (pKa = 1.9) (Varanasi et al., 2018).

145

Despite extremely low zeta potential, Pickering emulsions, i.e., CL-1.5 and CL-

2.0, had good stability against flocculation and coalescence as compared to the C-0.5,

C-1.0, C-1.5, C-2.0, CL-0.5 and CL-1.0. This finding confirmed that the stability of

CL-1.5 and CL-2.0 at the SGJ phase was due to steric repulsion rather electrostatic effects (Zoppe et al., 2012 and Meirelles et al., 2020).

Results from the creaming index showed a significant (p≤0.05) increase in the creaming index from 2-3% (SSJ) to 70-80% (SGJ) for the C-0.5, C-1.0, C-1.5, C-2.0 and CL-0.5. However, a creaming index of up to 23% for CL-1.0 and 5% for the CL-

1.5 and CL-2.0 was recorded (Fig 6.7). This finding again confirmed the strong stability of CL-01.5 and CL-2.0 against the SGJ phase of the GIT.

Figure 6.7. Effect of in vitro gastrointestinal tract (GIT) phases on creaming index of emulsions.

100 C-0.5 C-1.0 C-1.5 C-2.0 80 CL-0.5 CL-1.0

Creaming (%) index Creaming 20

0 Initial SSJ SGJ SIJ

GIT Phases Note: Emulsions stabilized with unmodified CNC (C) and lauric acid adsorbed CNC (CL) with concentrations of 0.5, 1.0, 1.5 and 2.0% (solid basis).

146

6.3.3.4. Simulated small-intestinal juice (SIJ) phase

Similar to the effect of SGJ phase on the Pickering emulsions, a gradual increase in the average particle size was recorded after the SIJ phase of GIT (Fig. 6.3).

An increase in the particle size for the Pickering emulsion with CNC viz., C-0.5, C-1.0,

C-1.5 and C-2.0 was simply assumed to be due to the effect of continuous dilution with simulated digestive juices (Winuprasith et al., 2018 and Bai et al., 2020a). As it is mentioned in the methods (sub-section 6.2.3), a 20 mL of Pickering emulsion mixed with 20 mL of SSJ and a fraction of 20 mL from the SSJ was further mixed with 20 mL of SGJ. Further, 20 mL from the SGJ phase was mixed with a total of 7.5 mL of

SIJ (1.5 SIJ + 3.5 mL bile juice + 2.5 lipase solution). The addition of this much digestive juice can cause substantial dilution of a Pickering emulsion leading to the loss of the initial arrangement of the CNC-CNC and CNC-lipid droplets (Fig. 6.6). CLSM images clearly displayed flocculation and coalescence of lipid droplets stabilized with

CNC at all the concentrations and lower concentrations of the CL. From these results, it can be inferred that the dilution of Pickering emulsion may have destabilized the complex structure of the Pickering structure leading to the flocculation/coalescence of droplets. In contrast to the above findings, the flocculation/coalescence of the lipid droplets for the Pickering emulsion stabilized with a higher concentration of CL i.e.,

CL-1.5 and CL-2.5 were quite stable against dilution (Fig. 6.5). A stable lipid droplet for the CL-1.5 and CL-2.0 against dilution can be due to adsorbed CL onto the interface of lipid droplets creating steric repulsion (Zoppe et al., 2012). Similarly, Bai et al.

(2019a) have reported a significant increase in the average particle size of the Pickering emulsion stabilized with CNC during the small intestinal phase of the GIT. An increase

147 in the average particle size was assumed to be due to the fusion of undigested lipid droplets, the formation of large, aggregated vesicles, and the association of CNC- micelle with present calcium soap. In contrast to these findings, Winuprasith et al.

(2018) have reported a significant decrease in the average particle size of the Pickering emulsion stabilized with CNF. A reduction in the average particle size at the small intestinal phase attributed to the digestion of lipid droplets reducing the diameter and the formation of a complex mixture of a micelle, small vesicle and calcium soap, which reflected the light scattering pattern of the samples (McClements et al., 2009).

The zeta potential of the Pickering emulsion reverted to negative when introduced into the SIJ phase of the GIT, resulting in the average zeta potential of up to -34.78±5.26 mV for the Pickering emulsion stabilized with CNC. However, up to -

25.71±4.22 mV occurred for the emulsion stabilized with CL (Fig. 6.4). The anionic zeta potential of the Pickering emulsion at the SIJ phase can be due to the following reasons: a) the digestion of lipid droplets released anionic molecules such as free fatty acids that contributed the negative zeta potential, b) the SIJ phase also contained several anionic molecules of bile extract such as bile salts and phospholipids can be absorbed on the lipid droplets, creating a negative zeta potential and c) neutralization of pH of a sample from SGJ phase from 2.0 to 7.0 can lead to the CNC becoming anionic again above its isoelectric point (pKa = 1.9) (Varanasi et al., 2018). Similar results were reported by Winuprasith et al. (2018) and Bai et al. (2019a) during in vitro GIT studies of O/W Pickering emulsion stabilized with CNF and CNC. Results also inferred that the lower concentration of the CNC or CL had higher zeta potential value (Fig. 6.4). It

148 was assumed that the lower concentration of CNC or CL was due to a rate of higher lipid digestion, thus a higher amount of released free fatty acids.

6.3.3.5. Lipid digestion

Results shown in Fig 6.8 displayed that the Pickering emulsion containing a lower amount of CNC or CL had increased lipid digestion, releasing a higher amount of free fatty acids (FFA) accounted for a total FFA of 73.35±3.29% for C-0.5, 67.27±3.26%.

However, no significant (p>0.05) difference was observed among the FFA values of

C-1.0, C-1.5 and C-2.0. Similarly, a total FFA of 68.27±2.41% for the CL-0.5 and

56.66±1.17% for the CL-1.0 was recorded and an insignificant (p>0.05) difference between CL-1.5 and CL-2.0 was observed. Results showed that the Pickering stabilized with CNC had slightly higher lipid digestion than the emulsion stabilized with CL. It was hypothesized that the higher lipid digestion for the Pickering emulsion stabilized with CNC can be due to higher lipase and lipid droplet interaction because dilution of a stable Pickering emulsion can result in a thinner/weaker gel strength of the continuous phase, disturbing the arrangement of CNC around the interface of lipid droplets, thus more interfaces available for the lipase activity (Fig. 6.6). However, dilution may not have affected the adsorbed CL onto the interface of lipid droplets less, thus limited lipase activity (Fig. 6.6). CL contained amphiphilic molecules on its surface that can facilitate better interfacial interaction as compared to the CNC (Paleos, 1969 and Ulrich et al., 1988). Digestibility of a vitamin D-loaded soybean oil Pickering emulsion stabilized with whey protein isolate (concentration 0.7%) and CNF (concentration 0.1-

0.7%) has been studied by Winuprasith et al. (2018), reported >90% and 75-85% release of FFA at the small intestinal phase of GIT. However, Bai et al. (2019a) found

149 a total FFA release that ranged from 40-75% for the Pickering emulsion stabilized with

CNC (concentration 0.1 to 1.0%) to >95% for the Pickering emulsion stabilized with gum arabic. Findings inferred that the CNC adsorbed onto the interface of lipid droplets limited the lipase activity, reducing the lipid digestion.

Fig. 6.8. Release of free fatty acids during the small intestinal phase.

100

80 a a b b b 60 c d d

40 Free acid fatty (%) Free

v 0 C-0.5 C-1.0 C-1.5 C-2.0 CL-0.5 CL-1.0 CL-1.5 CL-2.0 CL-2.0* Sample types

Note: Emulsion stabilized with neat CNC (C) and lauric acid adsorbed CNC (CL) with different concentrations of 0.5, 1.0, 1.5 and 2.0% (solid basis). 2% of lauric acid adsorbed CNC with no emulsion (CL-2.0*) was taken to observe the release of fatty acid from adsorbed CNC. The alphabets denote significant differences (p ≤0.05) among sample types.

6.3.3.6. Bioaccessibility of beta-carotene

Beta-carotene was used as a model nutraceutical to explore the influence of

CNC and CL on the bioaccessibility. Results shown in Fig. 6.8 and Fig. 6.9 displayed an apparent relationship between the degree of lipid digestion and beta-carotene

150 bioavailability. The Pickering emulsion with higher lipid digestion had a higher beta- carotene bioaccessibility, and so decreased lipid digestion had lower bioaccessibility.

It is well understood that higher the degree of lipid digestion, the more the mixed micelle formation, thus higher bioaccessibility (Winuprasith et al., 2018). The application of CNC or CL could significantly limit the bioaccessibility of nutraceuticals by arresting the lipid digestion process.

Fig. 6.9. Bioaccessibility of beta-carotene.

100

80 a b b b c d 60 e e

40 Bioaccessibility(%)

0 C-0.5 C-1.0 C-1.5 C-2.0 CL-0.5 CL-1.0 CL-1.5 CL-2.0

Sample types

Note: Alphabet letters denote significant differences (p ≤0.05) among emulsions.

151

6.4. Conclusion

An O/W type Pickering emulsion containing CNC or lauric acid adsorbed CNC

(CL) has good potential to stabilize lipid droplets by coating the interfacial area.

Adsorption of CNC or CL onto the interface of lipid droplets created a good electrostatic and steric repulsion maintaining the stability of stabilized Pickering emulsion during initial and mouth phases of the in vitro GIT study. However, a substantial flocculation/coalescence of the lipid droplets was observed for the Pickering emulsion stabilized with CNC during gastric and small intestinal phases of the GIT.

Flocculation or coalescence of the lipid droplets was confirmed with CLSM. In contrast to the flocculation/coalescence of lipid droplets for the Pickering emulsion stabilized with CNC, it was observed that CL with a concentration of above 1% displayed a potential maintaining the stability of lipid droplets through the GIT phases. CL adsorbed onto the interface of lipid droplets showed a strong steric repulsion at the gastric phase even though the zeta potential was below zero. Pickering emulsion with

CL had lower lipid digestion as compared to the Pickering emulsion stabilized with

CNC. Similarly, the bioaccessibility of beta-carotene was found to be lower for the emulsion containing CL as compared to the CNC as compared to the emulsion stabilized with CNC. Study concluded that lauric acid-adsorbed CNC be effectively utilized to create a stable emulsion through GIT limiting lipid digestion; however, it may not be a suitable emulsifier for developing Pickering emulsions to deliver a nutraceutical for absorption in the small intestine.

152

CHAPTER 7

OVERALL CONCLUSIONS AND RECOMMENDATIONS

This study inferred that the CNC as a biopolymer could significantly be utilized as an emulsifier or stabilizer for the lipid carriers. However, the surface modification of the

CNC by means of either chemical or physical is needed. A physical modification of

CNC by adsorbing a compatible molecule such as polyethylene glycol (PEG) or medium-chain fatty acids, i.e., lauric acid on CNC's surface, can add the value to cellulose nanomaterials industry. CNC modified via adsorption of PEG facilitated the re-dispersion of the dried CNC into an aqueous medium, and the stabilization of phycobiliprotein loaded liposomes. Likewise, adsorption of lauric acid on the surface of CNC (CL) created a stable O/W Pickering emulsion and impede the coalescence of the beta-carotene loaded lipid droplets. Unlike wise, the Pickering emulsion stabilized by neat CNC, CL preserved the original particle size of the emulsion during dilution and in vitro gastrointestinal digestion process. Moreover, it was observed that the CL impeded the lipolysis process of the beta-carotene loaded O/W emulsion resulting in a decreased bioaccessibility of the beta-carotene. It was concluded that the modified

CNC might not be a suitable candidate for the encapsulation and delivery of nutraceuticals due to its hypo-lipolysis activity. However, modified CNC can be used to extend the rate and degree of lipolysis into the small intestine, hence a sustained release of the loaded nutraceuticals. CL can also be utilized as a potential emulsifier in designing low-calorie functional foods. Since lauric acid is an amphiphilic, ethanol- soluble molecule, adsorption of ethanol-soluble antioxidants on CNC’s surface can

153 facilitate the hydrophobic surface with antioxidant property, which can enhance the oxidative stability of the emulsion.

These processes may be impractical to implement commercially due to cost and potential legal barriers. Future research on the adsorptive modification of CNC must focus on the use of food-grade solvents and other reagents that will be accepted by the

Food and Drug Administration. Additional research is needed in animal and human trials to verify the biocompatibility/toxicity of the modified CNC as well as the bioavailability and efficacy of nutraceuticals encapsulated by CNC versus other means of delivery.

154

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BIOGRAPHY OF THE AUTHOR

Avinash Singh Patel was born in Varanasi, Uttar Pradesh state of India on May 07,

1992. He graduated in Biology from Mahatma Gandhi Kashi Vidyapith, Varanasi,

India in June 2012 and attended Center Food Science and Technology at the Banaras

Hindu University, Varanasi, India in July 2012 for his postgraduation study. Avinash has worked for two and a half years as Senior Research Fellow at the Indian

Agricultural Research Institute, New Delhi, India. He is a recipient of the Netaji Subhas

ICAR Fellowship in 2016. He is a candidate for the Doctor of Philosophy degree in

Food and Nutrition Sciences from the University of Maine in December 2020.

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