Responsive chitosan-based microgels

Responsive Chitosan-Based Microgels

Von der Fakultät für Mathematik, Informatik und Naturwissenschaften der RWTH

Aachen University zur Erlangung des akademischen Grades einer Doktorin der

Naturwissenschaften genehmigte Dissertation

vorgelegt von

M. Sc. Helin Li

aus Shaanxi, China

Berichter:

Prof. Dr. Andrij Pich

Prof. Dr. Felix A. Plamper

Tag der mündlichen Prüfung: 07. 05. 2021

Diese Dissertation ist auf den Internetseiten der Universitätsbibliothek online verfügbar.

II

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For My Family

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V

Content Summary ..................................................................................................................... 1

Zusammenfassung ....................................................................................................... 5

List of Abbreviations ................................................................................................... 9

1. Introduction ........................................................................................................... 15

1.1 An Overview of Functional Microgels ............................................................. 16

1.1.1 Biopolymer-Based Microgels .................................................................... 16

1.1.2 Conductive Polymer-Based Microgels ....................................................... 22

1.2 Properties of Functional Microgels ................................................................... 27

1.2.1 pH-Sensitive Properties of Microgels ........................................................ 27

1.2.2 Redox-Active Properties of Microgels ....................................................... 28

1.3 Applications of Chitosan-Based Microgels ....................................................... 29

1.3.1 Drug Delivery ........................................................................................... 29

1.3.2 Functional Coatings .................................................................................. 36

1.3.3 Tissue Regeneration .................................................................................. 38

1.3.4 Filtration and Purification .......................................................................... 40

1.4 Aim and Motivation ......................................................................................... 43

1.5 Scope of the Thesis .......................................................................................... 44

1.6 References and Notes ....................................................................................... 45

2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels ................ 59

2.1 Introduction ..................................................................................................... 60

2.2 Experimental Section ....................................................................................... 64

2.2.1 Materials ................................................................................................... 64

2.2.2 Synthesis of Microgels .............................................................................. 64

2.2.3 Degradation of Microgels .......................................................................... 65

2.2.4 Drug Loading and Release Studies ............................................................ 67

2.2.5 Electrochemical Assay .............................................................................. 68

2.2.6 XTT Assay................................................................................................ 69

2.2.7 Characterization Methods .......................................................................... 70

2.3 Results and Discussion..................................................................................... 71

2.3.1 Synthesis of Microgels via Oxidative Polymerization ................................ 71

2.3.2 Chemical Composition of Microgels ......................................................... 73

2.3.3 Influence of pH on Microgel Size and Electrophoretic Mobility ................. 75

2.3.4 Colloidal Stability of Microgels ................................................................. 78

2.3.5 Electrochemical Properties of Microgels ................................................... 79

2.3.6 Degradation of Microgels ....................................................................... 84

VI

2.3.7 Drug Loading and Release Studies ............................................................ 93

2.3.8 Cytotoxicity Evaluation ............................................................................. 94

2.4 Conclusions ..................................................................................................... 95

2.5 References and Notes ....................................................................................... 96

3. Polyaniline-Chitosan Microgels ........................................................................... 101

3.1 Introduction ................................................................................................... 102

3.2 Experimental Section ..................................................................................... 107

3.2.1 Materials ................................................................................................. 107

3.2.2 Synthesis of Chitosan-Grafted-Polyaniline (CH-g-PANI) Copolymers..... 108

3.2.3 Synthesis of Microgels (W/O miniemulsion) ........................................... 109

3.2.4 Characterization ...................................................................................... 110

3.2.5 Enzymatic Degradation of Microgels....................................................... 111

3.3 Results and Discussion................................................................................... 112

3.3.1 Synthesis of Microgels ............................................................................ 112

3.3.2 FTIR Spectra of Microgels ...................................................................... 116

3.3.3 Influence of pH on Microgel Size and Electrophoretic Mobility ............... 118

3.3.4 Electrochemical Properties ...................................................................... 120

3.3.5 Degradation of Microgels ........................................................................ 122

3.4 Conclusion..................................................................................................... 124

3.5 References and Notes ..................................................................................... 125

4. Dual-Degradable Dextran-Chitosan Microgels ..................................................... 131

4.1 Introduction ................................................................................................... 132

4.2 Experimental Section ..................................................................................... 136

4.2.1 Materials ................................................................................................. 136

4.2.2 Synthesis of 3-Azidopropyl Carbonylimidazole ....................................... 137

4.2.3 Synthesis of Azide Modified Dextran (Dextran-Azidopropylcarbonate) ... 138

4.2.4 Synthesis of Alkyne Modified Chitosan (Alkyne-Pendant Chitosan) ........ 138

4.2.5 Synthesis of Microgels via Click Cross-linking Reactions ....................... 139

4.2.6 Characterization Methods ........................................................................ 141

4.2.7 Alkaline-Induced Degradation ................................................................. 141

4.2.8 Enzymatic Degradation ........................................................................... 142

4.2.9 Drug Loading and Release Studies .......................................................... 143

4.2.10 Cytotoxicity Study in Vitro .................................................................... 144

4.2.11 Statistical Analysis ................................................................................ 144

4.3 Results and Discussion................................................................................... 145

4.3.1 Chemical Structure of Microgels ............................................................. 145

VII

4.3.2 Influence of pH on Microgels Size and Electrophoretic Mobility ............. 149

4.3.3 Degradation of Microgels ........................................................................ 151

4.3.4 Cytotoxicity Evaluation ........................................................................... 158

4.3.5 Drug Loading and Release Studies .......................................................... 159

4.4 Conclusion..................................................................................................... 162


5. Conclusion and Outlook ...................................................................................... 167


6. Acknowledgement ............................................................................................... 171

7. List of Publications .............................................................................................. 173

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Summary/Zussamenfassung

1

Summary

Regarding the development of stimuli-responsive microgels as drug delivery

systems for cancer therapies, improvements in biocompatibility, stability, and

controlled release are the major challenges due to the generally limited dosages

of anticancer drugs capable of being loaded, poor drug bioavailability, and non-

specialized drug administration. To overcome those challenges, this Thesis

presents various pH-sensitive biopolymer-based microgel systems which exhibit

good biocompatibility and biodegradability (whilst producing non-toxic

degradation by-products), thus demonstrating the great potential for the

incorporation of various active agents including drugs and biologics. Based on

these properties, microgels can be utilized as drug delivery vehicles for stimulus-

triggered degradation and controlled drug delivery, thus suggesting that the

presented microgel systems are good candidates for site-specific cancer

therapies.

This Thesis focuses on conductive polymer-based, as well as biopolymer-

based microgels, for use in drug delivery systems. Chapter 1 provides an

overview of different functional microgels. These microgels exhibit good

biocompatibility, biodegradability, non-toxicity, pH-sensitivity, redox-activity,

and adjustable chemical and mechanical properties. These properties endow

them with a wide variety of applications, such as drug encapsulation, which

facilitates their use as delivery systems, electrical sensors, and functional


2

coatings, as well as their application in areas such as tissue regeneration and

wastewater filtration.

Chapter 2 introduces a controlled drug release system: drug-loaded

biopolymer-based microgels. Due to the present problems faced with its use,

such as insufficient cellular uptake as well as the numerous drug resistance

mechanisms in cells, an anticancer drug, doxorubicin (DOX), has been

developed to be capable of being encapsulated into nanocarriers. Moreover, this

drug can also be released under the control of the microenvironment, most

notably in tumor tissues. The Thesis details the preparation of cross-linked

chitosan-poly(hydroquinone) (CHHQ) microgels with pH and redox sensitivity.

Due to their pH-sensitivity, redox-activity, and biodegradability, CHHQ

microgels have previously been exploited to load and release DOX. The loading

of the active ingredient is achieved by means of physical entrapment of both π-

π stacking and hydrogen bonding between chitosan, poly(hydroquinone), and

DOX. The drug loading profiles were investigated and an encapsulation

efficiency of 80.9% was observed. The drug release profiles show that

approximately 43% of DOX is released over one hour at pH 6; contrastingly,

very little DOX release is observed over the same time period at pH 7.4. These

results suggest that CHHQ microgels are a promising anti-tumor drug carrier for

anticancer drug delivery systems.

Chapter 3 describes the development of chitosan-poly(aniline) (CH-PANI)

microgels. These microgels exhibit both pH-sensitivity and redox-activity. The

CH-PANI microgels are composed of chitosan and poly(aniline), using

glutaraldehyde as the cross-linker. The degradation results show that CH-PANI

microgels can be degraded in an acidic environment, in the presence of lysozyme.

The results suggest that the prepared CH-PANI microgels hold great potential

as drug delivery carriers for the selective delivery of therapeutics to acidic

tissues, such as tumors.


3

Chapter 4 details and explores how novel pH-sensitive dual-degradable

dextran-chitosan (DE-CH) microgels are suitable as drug carriers for the

efficient, targeted delivery of drugs to the colon. A series of DE-CH microgels

were synthesized by cross-linking two modified biopolymers, alkyne-modified

chitosan, and azide-modified dextran with varying azide:alkyne molar ratios

from 1:0.5, 1:1, 1:1.5 to 1:2. The microgels were cross-linked via copper(II)-

catalyzed azide-alkyne cycloaddition (CuAAC) without a cross-linker. By

conducting dynamic light scattering (DLS) and electrophoretic mobility studies,

it was demonstrated that the microgels were pH-sensitive. Under slightly acidic

conditions, the microgels can be degraded in the presence of dextranase, an

enzyme present in the colon. In addition, the prepared DE-CH microgels are

capable of loading vancomycin hydrochloride (VM), an antibiotic effective

against many gram-positive bacteria. The results showed an encapsulation

efficacy of up to 93.7%, indicating a possible application for the microgels as an

effective platform for site-specific targeted drug delivery (e.g., to the colon).


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5

Zusammenfassung

Die größten Herausforderungen bei der Entwicklung stimuli-responsiver

Mikrogele als Wirkstofffreisetzungssysteme (engl. drug-delivery systems) für

die Krebstherapie sind die Biokompatibilität, die Stabilität und die kontrollierte

Freisetzung. Diese Limitierungen basieren auf der bisher im allgemeinen

begrenzten Dosierung zur Beladung vorgesehener Krebsmedikamente,

schlechter Bioverfügbarkeit der Medikamente und nicht spezialisierter

Medikamentenverabreichung. Zur Bewältigung dieser Herausforderungen

werden in dieser Dissertation verschiedene pH-empfindliche Mikrogelsysteme

auf Biopolymer-Basis vorgestellt, die eine gute Biokompatibilität sowie

biologische Abbaubarkeit mit nicht-toxischen Abbaunebenprodukten aufweisen

und somit großes Potenzial für die Einarbeitung von Wirkstoffen, einschließlich

verschiedener Arzneimittel und Biologika, aufweisen. Basierend auf diesen

Eigenschaften können sie als Vehikel für den Wirkstofftransport, für Stimulus-

induzierten Abbau und kontrollierte Wirkstofffreisetzung eingesetzt werden,

was darauf hindeutet, dass die vorgestellten Mikrogelsysteme gute Kandidaten

für die ortsspezifische Krebstherapie darstellen.

Diese Dissertation konzentriert sich auf Mikrogele für die Anwendung als

Wirkstofffreisetzungssysteme. Die dabei eingesetzten Mikrogele sind sowohl

auf Basis leitfähiger Polymere als auch auf Biopolymer-Basis. Kapitel 1 gibt

einen Überblick über verschiedene funktionale Mikrogele. Diese Mikrogele

weisen eine gute Biokompatibilität, biologische Abbaubarkeit, Ungiftigkeit, pH-

Empfindlichkeit, Redox-Aktivität und einstellbare chemische und mechanische

Eigenschaften auf. Diese Eigenschaften verleihen ihnen eine Vielzahl von


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Anwendungen, wie die Verkapselung von Medikamenten und damit den Einsatz

als Transport- und Freisetzungssysteme oder als elektrische Sensoren und

funktionelle Beschichtungen, sowie in Bereichen der Geweberegeneration und

Abwasserfiltration.

In Kapitel 2 wird anhand von Mikrogelen auf Biopolymer-Basis, die mit

Medikamenten beladen sind, ein System zur kontrollierten

Medikamentenfreisetzung vorgestellt. Aufgrund der derzeitigen

Einschränkungen bei der Anwendung, wie z. B. der unzureichenden zellulären

Aufnahme sowie der zahlreichen Resistenzmechanismen in den Zellen, wurde

ein Krebsmedikament, Doxorubicin (DOX), entwickelt, das in Nanocarrier

verkapselt werden kann. Darüber hinaus kann dieses Krebsmedikament unter

der Kontrolle der Mikroumgebung, insbesondere in Tumorgewebe, freigesetzt

werden. In dieser Dissertation wird die Herstellung von vernetzten Chitosan-

Poly(hydrochinon) (CHHQ)-Mikrogelen mit pH- und Redox-Empfindlichkeit

gezeigt. Aufgrund ihrer pH-Empfindlichkeit, Redox-Aktivität und biologischen

Abbaubarkeit wurden CHHQ-Mikrogele bereits zuvor zur Beladung und

Freisetzung von DOX genutzt. Die Wirkstoffbeladung erfolgt durch

physikalischen Einschluss sowohl durch π-π-Wechselwirkungen als auch durch

Wasserstoffbrückenbindungen zwischen Chitosan, Poly(hydrochinon) und

DOX. Die Wirkstoffbeladungsprofile wurden untersucht und zeigen, dass die

Einkapselungseffizienz 80,9% beträgt. Die Wirkstofffreisetzungsprofile zeigen,

dass bei einem pH-Wert von 6 innerhalb von einer Stunde etwa 43% DOX

freigesetzt werden, während bei einem pH-Wert von 7,4 über den gleichen

Zeitraum nur eine geringe DOX-Freisetzung zu beobachten ist. Diese

Ergebnisse deuten darauf hin, dass die CHHQ-Mikrogele einen

vielversprechenden Antitumor-Wirkstoffträger für Antikrebs-

Wirkstofffreisetzungssysteme darstellen.


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In Kapitel 3 wird die Entwicklung von Chitosan-Poly(anilin) (CH-PANI)-

Mikrogelen beschrieben. Diese Mikrogele weisen sowohl pH-Empfindlichkeit

als auch Redox-Aktivität auf. Die CH-PANI-Mikrogele bestehen aus Chitosan

und Poly(anilin) und wurden unter Verwendung von Glutaraldehyd als

Vernetzer synthetisiert. Die Degradationsergebnisse zeigen, dass CH-PANI-

Mikrogele in Gegenwart von Lysozym in saurem Milieu abgebaut werden

können. Die Ergebnisse deuten darauf hin, dass die hergestellten CH-PANI-

Mikrogele ein großes Potenzial als Wirkstoffträger für die selektive Abgabe von

Therapeutika an saures Gewebe, wie z.B. Tumore, besitzen.

In Kapitel 4 werden neue pH-sensitive, dual abbaubare Dextran-Chitosan

(DE-CH)-Mikrogele beschrieben, die als Wirkstoffträger für die effiziente und

gezielte Freisetzung von Medikamenten in den Dickdarm geeignet sind. Eine

Reihe von DE-CH-Mikrogelen wurde durch Vernetzung von zwei modifizierten

Biopolymeren, alkinmodifiziertem Chitosan und azidmodifiziertem Dextran,

mit unterschiedlichen Molverhältnissen von Azid zu Alkin von 1:0,5, 1:1, 1:1,5

bis 1:2 synthetisiert. Die Mikrogele wurden durch eine Kupfer(II)-katalysierte

Azid-Alkin-Cycloaddition (CuAAC) ohne einen Vernetzer vernetzt.

Dynamische Lichtstreuung (DLS) und elektrophoretische Mobilitätsstudien

zeigen die pH-Sensitivität dieser Mikrogele. Unter leicht sauren Bedingungen

können die Mikrogele in Gegenwart von Dextranase, einem im Dickdarm

vorkommenden Enzym, abgebaut werden. Darüber hinaus können die

hergestellten DE-CH-Mikrogele Vancomycin Hydrochlorid (VM) aufnehmen,

ein Antibiotikum, das gegen viele grampositive Bakterien wirksam ist. Die

Verkapselungseffizienz beträgt bis zu 93,7%, was darauf hindeutet, dass die

Mikrogele möglicherweise als wirksame Plattform für eine ortsspezifische und

gezielte Wirkstoffabgabe (z.B. im Dickdarm) eingesetzt werden können.


8

List of Abbreviations

9


List of Chemicals

AG Agarose

Alg Alginate

AIBN 2, 2’-Azoisobutyronitrile

AP-CI 3-Azidopropyl carbonylimidazole

APS Ammonium persulfate

ASGP Asialoglycoprotein

BSA Bovine serum albumin

CAT Catalase

CDI 1-1’-Carbonyldiimidazole

CHHQ Chitosan-poly(hydroquinone)

CH-PANI Chitosan-polyaniline

ChS Chondroitin sulfate

CL Cellulose


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CMCs-CBA-Dox NPs Carboxymethyl chitosan-

carboxybenzaldehyde-doxorubicin

nanoparticles

CS Chitosan

CuS Copper sulphide

DA Dopamine

DCl Ceuterium chloride

Dex Cextran

DFO Deferoxamine

DMSO Cimethyl sulfoxide

DMF N,N-Dimethylformamide

D2O Ceuterium oxide

DOX Doxorubicin

EDC N-(3-dimethylaminopropyl)-N’-

ethylcarbodiimide hydrochloride

EM Emeraldine

FA Folic acid

Fc Ferrocene

Fe3O4 Iron(II,III) oxide

FR Folate receptor


11

GA Glutaraldehyde

Ga-DTPA Gadopentetate dimeglumine

Gal Galactose

GC Glassy carbon

GOx Glucose oxidase

GSH Glutathione

HA Hyaluronic acid

HCC Hepatocellular carcinoma

HCl Hydrochloric acid

HCS Hydrochloride chitosan

2-Hydroxyethyl methacrylate 2-Hydroxyethyl methacrylate

K2HPO4 Dibasic potassium phosphate

LA Lactobionic acid

LM Leucoemeraldine

MES 2-(N-morpholino)ethanesulfonic

acid

MPS Mononuclear phagocyte system

mPEG Methoxy poly(ethylene glycol)

MTT (3-(4,5-Dimethylthiazol-2-yl)-2,5-

diphenyltetrazolium bromide)


12

NA Nigraniline

NHS N-hydroxysuccinimide

NH3·H2O Ammonium hydroxide solution

NHS-PEG-NHS Succinimide-end polyethylene

glycol

NIPAAm-co-AA N-isopropylacrylamide-co-acrylic

acid

NMP methylpyrrolidone

PAA Poly(acrylic acid)

PANI Poly(aniline)

PA6 Polyamide 6

p-CBA P-Carboxybenzaldehyde

PDMAEMA Poly(N,N-dimethylaminoethyl

methacrylate)

PDPA-b-PEI Poly(2-(diisopropylamino

ethylmethacrylate)-block-

Poly(ethyleneimine)

PE Pectin

PEDOT Poly(3,4-ethylenedioxythiophene)

PEG Poly(ethylene glycol)

PENPs Polyelectrolyte nanoparticles


13

PFH Perfluorohexane

PHQ Poly(hydroquinone)

PLGA Poly(L-glutamic acid

PMAA Poly(methacrylic acid)

PNA Pernigraniline

PNIPAM Poly(N-isopropylacrylamide)

PPy Poly(pyrrole)

PSBMA Poly(sulfobetaine methacrylate)

PSS Poly(4-styrene sulfonate)

Pt Platinum

PuL Pullulan

PVCL Poly(N-vinylcaprolactam)

PVP Poly(4-vinylpyridine)

RBC Red blood cells

Si-QAC 3-(trimethoxysilyl)-

propyldimethyloctadecyl

ammonium chloride

Sr-GO Strontium-graphene oxide

TPP Tripolyphosphate

VM Vancomycin hydrochloride


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XTT Sodium 2,3-bis-(2-methoxy-4-nitro-

5-sulfophenyl)-5-[(phenylamino)-

carbonyl]-2H-tetrazolium

List of Instruments and Methods

ATR-FTIR Attenuated total reflectance Fourier

transform infrared spectroscopy

CuAAC Copper(II)-catalyzed azide-alkyne

click reaction

CV Cyclic voltammetry

DLS Dynamic light scattering

FTIR Fourier transmission infrared

spectroscopy

LCST Lower critical solution temperature

micro-CT Micro-computed tomography

MPS Mononuclear phagocyte system

clearance

MRI Magnetic resonance imaging

TEM Transmission electron microscopy

UV Ultraviolet

1. Introduction

15

1. Introduction

1. Introduction

16

1.1 An Overview of Functional Microgels

Micro- and nanogels are micro- and nanometer-sized cross-linked colloidal

polymer networks1, which have high water absorption capacities2, tunable

microstructures3, biocompatibility4, biodegradability5 and adjustable chemical

and mechanical properties6. Moreover, their large surface area offers

opportunities for multivalent bioactive conjugates and an interior network for

drug encapsulation7. Microgels can swell or shrink in response to external

stimuli, such as alterations in temperature8, pH value9, ionic strength10, and

light11. These unique properties offer great potential for fabricating functional

microgels for use in a diverse range of applications as vehicles for drug

encapsulation and delivery, incorporated in vitro for cell expansion and

proliferation and in vivo for tissue regeneration and reconstruction3.

1.1.1 Biopolymer-Based Microgels

Natural and synthetic polymers are commonly utilized to form microgels.

Examples of synthetic polymers include poly(acrylic acid) (PAA)12,

poly(methacrylic acid) (PMAA)13, poly(4-vinylpyridine) (PVP)14, poly(N,N-

dimethylaminoethyl methacrylate) (PDMAEMA)15, poly(N-vinylcaprolactam)

(PVCL)16 and poly(N-isopropylacrylamide) (PNIPAm)17. These synthetic

polymers are commonly used to prepare synthetic micro- or nanogels in the

presence of multifunctional cross-linkers18.

Recently, biopolymer-based microgels/nanogels (biomicrogels/bionanogels)

gained increasing interest because they can overcome some of the problems

associated with synthetic materials, such as poor biodegradability and their

toxicity to the environment. These biomicrogels/bionanogels not only exhibited

1. Introduction

17

the properties of synthetic counterparts but also possessed unique properties

such as biocompatibility19, biodegradability20, bioaccessibility21, nontoxicity22

and low price23. Moreover, biopolymer-based micro- or nanogels exhibit a

variety of functional groups containing hydroxyl, amino, and carboxylic acid

groups, which are developed for cross-linking with various cross-linkers and

conjugating with cell-targeting agents24.

The typical examples of naturally occurring biopolymers are polysaccharides,

e.g. chitosan (CS)25, dextran (Dex)26, cellulose (CL)27, pectin (PE)28, hyaluronan

(HA)29, pullulan (PuL)30, chondroitin sulfate (ChS)31, agar32, agarose (AG)33,

and alginate (Alg)34, which are commonly used for preparing biocompatible

microgels. Some of the most common naturally occurring biopolymers are

shown in Fig. 135.

Fig. 1. Natural biopolymers that can be used in drug delivery systems35.

1.1.1.1 Chitosan

Of the many biopolymers in existence, chitosan, a natural polysaccharide, is

gaining attention, and as such, is being widely considered for micro- and

nanoparticle preparation36.

1. Introduction

18

Chitosan is a linear polysaccharide composed of β-(1-4)-linked D-

glucosamine (deacetylated unit) and N-acetyl-D-glucosamine (acetylated unit).

It is obtained from the partial deacetylation of chitin, the second most abundant

natural polymer, and is composed of a series of polymers varying in their degree

of molecular weight, viscosity, pKa etc37. It has an average molecular weight

between 50 kDa and 2,000 kDa38. Chitin and chitosan can be commercially

obtained from shellfish sources such as crabs, shrimps, and krill, as well as

insects and fungi. They have gained considerable attention because not only can

they be obtained from a renewable resource but also they are non-toxic,

compatible biomaterials that show a broad range of potential applications,

especially in the biomedical field39.

Chitosan contains three types of reactive groups (an amino group and two free

hydroxyl groups) with the primary and secondary hydroxyl groups being located

in the repeating glucosidic residue. Due to these functional groups, the chemical

modification of these groups of chitosan has offered a number of useful materials

used in a wide range of fields for a wide of applications, such as food, cosmetics,

as well as biomedical and pharmaceutical applications40. Chemically modified

chitin and chitosan structures are obtained by generating free radicals upon the

chitosan chain, which react with polymerizable monomers, resulting in a grafted

chain41. Moreover, upon ionization of amino groups, the increased charge

density renders chitosan suitable for chemical reactions such as alkylation,

acylation and carboxyl-methylation42.

The quaternization of amino groups makes chitosan a cationic polyelectrolyte

with a pKa of about 6.5. At low pH values, the amino groups become protonated

and positively charged. Its polycationic surface makes chitosan suitable for

adhering to negatively charged substrates, aggregating polyanionic compounds,

and chelating with different metal ions, such as Ca2+, Ba2+ and Al3+ 43.

1. Introduction

19

For characteristics such as biocompatibility, biodegradability, bioadhesivity,

bioactivity and low toxicity, chitosan has been used as a pharmaceutical

excipient in a wide range of biomedical fields including cell proliferation44,

tissue engineering45 and targeted drug delivery systems46.

1.1.1.2 Fabrication of Microgels Based on Chitosan

Due to their excellent biocompatibility and biodegradability, polysaccharides

(e.g., chitosan and dextran) are often taken to be the ideal candidates in the fields

of medicine and biotechnology for designing and fabricating micro- and

nanoparticles47. Additionally, owing to the large number of amino groups in its

chain, chitosan was employed as an ideal material for preparing microgels that

exhibit significant pH-responsive behaviors48.

Different methods for the fabrication of polymer-based micro- or nanogels

can be carried out by cross-linking polymer chains via chemical or physical

interactions.

Chemical cross-linking is a suitable strategy to generate chitosan-based

microgel networks by implying covalent cross-linking reactions among

functional groups of chitosan (e.g., amino groups and hydroxyl groups) and

different kinds of cross-linking agents, e.g., glutaraldehyde (GA)49, glyoxal50,

genipin51 or succinimide-end-functionalized polyethylene glycol52. The reaction

between chitosan and glutaraldehyde can be achieved through the amine groups

of chitosan and the aldehyde groups of glutaraldehyde, forming covalent imine

bonds. Another form of cross-linker is genipin, an aglycon derived from

geniposide. It represents an excellent natural cross-linker for cross-linking with

chitosan, proteins, collagen and gelatin whist also being less toxic than many

other synthetic cross-linkers. As such, it has been widely used in biomedical

applications53.

https://www.sciencedirect.com/science/article/pii/S0142961211008404

1. Introduction

20

Wang et al. introduced a novel biocompatible microgel via naturally derived

cross-linked polymers, such as chitosan and gelatin, with succinimide-end

polyethylene glycol (NHS-PEG-NHS)54. The obtained microgels are

biocompatible due to their naturally derived components, gelatin and chitosan,

as well as the cross-linker PEG. Moreover, they meet the requirements of

biocompatibility, drug encapsulation, and size control. The biocompatibility of

the prepared microgels were evaluated using MTT (3-(4,5-dimethylthiazol-2-

yl)-2,5-diphenyltetrazolium bromide) assay in vitro. The microgels had the

ability to encapsulate hydrophobic drugs for sustained delivery.

Another crosslinking method for fabricating chitosan-based microgels is

physical interactions, such as hydrogen bonding55, van der Waals forces56,

electrostatic interactions57, or hydrophobic associations58.

These physical cross-linking strategies that are achieved through various

synthesis routes were also investigated by many researchers to prepare chitosan-

based microgels. For example, as described in Chapter 2 59, Li et al. introduced

an electroactive supramolecular microgel cross-linked via hydrogen bonds. We

prepared a redox-active microgel with dual responsiveness, and pH and redox-

responsiveness. The microgels were cross-linked by hydrogen bonds between

chitosan and poly(hydroquinone) in an inverse miniemulsion system. The

microgels can encapsulate doxorubicin (DOX), which will be released from the

microgel in the presence of lysozyme. Therefore, the obtained electroactive

microgels could be developed for use in biomedical fields.

Gu et al. introduced a pH-responsive microgel cross-linked via electrostatic

interaction60. A physically cross-linked pH-responsive injectable microgel,

consisting of a chitosan matrix, glucose-specific enzyme and recombinant

human insulin, was fabricated to achieve glucose-responsive closed-loop insulin

delivery. In these systems, the chitosan matrix was cross-linked by

tripolyphosphate (TPP) through electrostatic interactions which also entrapped

1. Introduction

21

the enzyme-loaded nanocapsules, glucose oxidase (GOx)-/catalase (CAT)-

containing enzyme nanocapsules, and insulin in the matrix. GOx is a glucose-

specific enzyme that can catalyze glucose to gluconic acid. Due to the enzymatic

conversion of glucose immobilized within microgels into glucomic acid, this

pH-sensitive matrix, an enzymatic nanocapsule-containing microgel, swelled

due to the decreased microenvironmental pH value, as shown in Fig. 2. The in

vivo studies indicated that these systems can release insulin and control blood

glucose levels in a mouse model of type 1 diabetes.

Fig. 2. Schematic representation of insulin and enzyme nanocapsules-loaded microgels.

Reprinted with permission from Ref. [60]. Copyright 2013 American Chemical Society.

Vahedifar et al. reported the calcium and chitosan-mediated clustering of

whey protein particles58. Based on the nature of biopolymers, proteins

complexed with chitosan could form particles in which biopolymeric clusters

are formed. In these particles, chitosan can interact with whey protein via

electrostatic interactions between deprotonated carboxyl groups of whey protein

and protonated amine groups of chitosan. Such reactions are hydrophobic

interactions between hydrophobic patches of whey proteins and the acetyl

groups of chitosan, along with hydrogen bonding through hydroxyl groups

between whey proteins and chitosan.

1. Introduction

22

1.1.2 Conductive Polymer-Based Microgels

These novel classes of polymers are known as intrinsically conducting

polymers. They are notable as they present interesting electrical and optical

properties61. Based on their electrical properties or modifications of these

properties, conductive polymers have been applied to prepare nanomaterials as

drug-delivering systems62, organic electrode materials63, electrochemical

biosensing devices64, and for use in electrocatalysis65, chromatography66,

membrane separation67, lithium-ion battery68, environmental monitoring69 and

electrochromic devices70.

1.1.2.1 Conductive Polymers

Conductive polymers have recently attracted great attention due to their

particular electronic properties71. For instance, they contain intrinsic electronic,

magnetic and optical properties like metals or semiconductors. On this basis,

they have been termed “synthetic metals”72. There have conjugated π-electron

systems contained in the polymeric backbone that render them conductive73.

These distinctive structures provide them with electronic properties such as

electrical conductivity, low energy optical transitions, low ionization potential

and high electron affinity74. Along the polymer chain, they have single and

double bonds. The mechanism for conductivity in these polymers is diverse and

complicated. Heeger proposed that the conducting polymers showed electrical

conductivity by several orders of magnitude of doping, such as solitons, polarons

and bipolarons which are the charge-storage devices in conducting polymers75.

Kroschwitz suggested that some factors such as conjugation length, chain length

and the charge transfer to adjacent molecules can affect the conductivity of

conductive polymers76. Many conducting polymers, such as poly(hydroquinone)

(PHQ)77, poly(aniline) (PANI)78, poly(pyrrole) (PPy)79, poly(furan)80,

poly(indole)81, poly(thiophene)82, poly(acetylene)83, poly(terthiophene)84,

1. Introduction

23

poly(fluorine)85, poly(3-alkylthiophene)86, poly(tetrathiafulvalene)87,

poly(naphthalene)88, poly(p-phenylene sulfide)89, poly(3,4-ethylene

dioxythiophene)90 and poly(p-phenylene vinylene)s91, have been investigated

and widely used in various fields92.

Fig. 3. Redox transition process from hydroquinone to benzoquinone93.

Among these conductive polymers, one of the important members is quinones.

Quinones can be used in many fields such as batteries94, sensors or biosensors95,

supercapacitors96 and electrical conductors97. These polymers can undergo

reversible two-electron oxidation and reduction. Due to their interesting specific

capacity, high redox potential, and advanced electrochemical reversibility, they

1. Introduction

24

can be applied as a class of high energy density electroactive materials96. The

electric behavior of hydroquinone is shown in Fig. 3 93. Due to its strong

reducibility, hydroquinone can lose two electrons to form benzoquinone. One

electron is lost from hydroquinone and forms quinhydrone radicals; one more

electron is then lost to form benzoquinone. The hydroquinone-benzoquinone

charge-transfer redox couple is named quinhydrone98.

The conductive polymer, poly(aniline) (PANI), has also attracted much

attention due to its reversible doping or dedoping process which is achieved

through the protonation of the polymer chain’s backbone99. In general, chemical

dopings of the conducting polymers are classified as either p-doping (oxidation)

or n-doping (reduction). PANI is a p-type semiconductor that can easily

transport charges through the process of doping or dedoping, as shown in Fig. 4

100. Furthermore, PANI exhibits electrochromic behavior101. Tetsuhiko

Kobayashi et al. found that polyaniline films present color variations depending

on the different potentials, ranging from -0.2 to 1.0 V vs. SCE. The colors are

transparent yellow at 0.2 V, green at 0.5 V, dark blue at 0.8 V, and black at 1.0

V102.

Fig. 4. Chemical structure of PANI (A) before and (B) after doping100b.

1. Introduction

25

1.1.2.2 Doping

Typical conjugated polymers are insulators or semiconductors. They are

electrically conductive at several doping levels, which is an electrochemical

technique that brings about significant changes in electrical conductivity103.

Following the doping or dedoping process, there will be the generation or

removal of the charge carriers in the neutral polymer chain. In the doping process,

the polymer can be partially oxidized or reduced due to the increased or

decreased amount of π-electrons over the polymer backbone104. There are two

types of doping: p-doping (partial oxidation of π system of the polymer chain)105

and n-doping (partial reduction of π system of the polymer chain)106.

1.1.2.3 Applications of Conductive Polymer-Based Microgels

Due to its highly conjugated polymer chain, conductive polymer-based

microgels can be used in various promising applications such as sensors, drug

delivery systems and catalysis. Due to their remarkable conductive performance,

metal-containing microgels in which the mental centers serve as redox-

responsive or paramagnetic generating active species for charge transfer, can be

applied as satisfactory smart redox sensors. Through electrochemical reduction

or by reducing linkers in microgels, these redox-cleavable crosslinkers in

microgels are degraded in a reducing environment, such as glutathione (GSH),

thus suggesting a drug delivery system. Moreover, the conductive polymers

showing a high activity for catalyzing oxidation reactions can be applied as

catalysts.

Conductive polymer-based microgels can be used in chemical oxidation

sensing. Xiong et al. reported an oxidation-triggered degradable nanogel used

for Fe3+-chelating107. The nanogel (oxNG-DFO) was prepared through the

copolymerization of an oxidation-sensitive host-guest crosslinker between β-

cyclodextrin (β-CD) and ferrocene (Fc), metal chelating deferoxamine (DFO)

1. Introduction

26

and AAm monomers. The obtained nanogels exhibited excellent chelating

activity to Fe3+ ions and oxidation-triggered degradable behavior. The results

indicated that cellular ferritin expression can be effectively reduced, thereby

regulating intracellular iron levels. The conductive nanogels can be applied as

various metal chelation therapies in humans by serving as chemical oxidation

sensors.

Microgels with redox sensitivity can be applied in drug delivery. The multi-

responsive (pH/redox/ultrasound) core-shell microgels were prepared for use in

a double-locked drug delivery system by Liu et al108. The pH-sensitive poly(2-

(diisopropylamino ethylmethacrylate)-block-poly(ethyleneimine) (PDPA-b-PEI)

copolymers were synthesized as micelles and the redox-responsive shells were

formed by Michael addition of a primary amine group of branched PEI using

disulfide as a cross-linker, which was specifically cleaved by glutathione (GSH).

The anticancer drugs, DOX and perfluorohexane (PFH) were encapsulated and

the drug cumulative release amount was close to 90%. The results indicated that

this multi-responsive microgel could be used as an effective drug carrier for

cancer treatments.

Conductive microgels can also be designed to be catalysts. Pich et al. prepared

a selenium (Se) modified Poly(N-vinylcaprolactam) (PVCL) microgel as

colloidal catalyst109. Se-containing PVCL microgels exhibit high catalytic

activity and selectivity during oxidation reactions (e.g., the oxidation of acrolein

to acrylic acid and methyl acrylate). Moreover, the hydrodynamic radii of the

microgels before and after oxidation by H2O2 at 50 °C are smaller than that at

20 °C, indicating that the temperature-responsiveness of the microgels is not

influenced by the addition of Se. For the sample B1.5 Se2.0, the hydrodynamic

radii of the microgels before and after oxidation by H2O2 were changed from

192 nm (20 °C) to 87 nm (50 °C) and from 179 nm (20 °C) to 91 nm (50 °C),

respectively. The obtained Se-modified microgels enable oxidation reactions

1. Introduction

27

with high activity to be selectively catalyzed, which suggests that they may be

attractive for use technical processes to reduce energy consumption.

1.2 Properties of Functional Microgels

The attractive phenomenon of the volume change of microgels has been

noticed for several decades. These volume changes can be triggered by

temperature110, pH111, ionic strength of the media112, electric fields113,

magnetism114, redox-potential108, ultrasound115, and photo-irradiation in solution

conditions116. Different environmental triggers can cause the microgels to shrink

or swell approaching the extremes. These microgels are intelligent and termed

as stimuli-responsive microgels. For instance, pH-sensitive microgels are

capable of changing volume by means of changing the pH of the solution. For

temperature-sensitive microgels, heating and cooling can induce microgels to

change their volume in response to variations in temperature. For electric field-

sensitive microgels, water electrolysis was used as a driving force from outside

and the microgels can be responsible for a high voltage117.

1.2.1 pH-Sensitive Properties of Microgels

Chitosan is a potential material for use in the preparation of pH-responsive

drug carriers due to its primary amine groups that can form a micro- or nanogel

network with pH-responsive behaviors, such as exhibiting swelling properties in

acidic mediums (pH < pKa) and shrinking behaviors in a basic environment (pH >

pKa)118.

The acidic environment in tumor tissues has been identified as an ideal trigger

for the selective delivery of anticancer drugs. In an acidic environment, the

moieties in the drug carriers were protonated, resulting in the destabilization of

1. Introduction

28

nanocarriers, which then accelerates the release of the drugs, as shown in Fig. 5

119. Therefore, the nanocarriers derived from chitosan showed pH-responsive

properties and have been widely applied in biomedicine for loading multiple

kinds of cargoes, e.g., drugs, cells, proteins and genes, and controlling the release

of such cargoes in anti-tumor drug delivery systems due to the tumor’s acid

microenvironment120.

Fig. 5. Schematic representation of pH-responsive drug release behavior of the chitosan

conjugated nanocarrier due to the repulsive forces among protonated amino groups in

chitosan121.

1.2.2 Redox-Active Properties of Microgels

As mentioned above, the typical pH-sensitive microgels are introduced as

drug carriers. A new class of stimuli-responsive materials, the redox-active

polymers, are incorporated to prepare various kinds of redox-active microgels to

achieve redox responsiveness by undergoing reversible oxidation/reduction

reactions. Under redox stimuli, which can be applied chemically or

electrochemically, these designed gels are capable of responding reversibly to

the applied redox stimuli, in a controllable and predictable manner. It is indicated

1. Introduction

29

that the redox-active microgels are the ideal candidate to be applied as electrical

sensors122, actuators123 or energy conversion124 and storage devices125.

1.3 Applications of Chitosan-Based Microgels

Chitosan, the only natural cationic polysaccharide, has attracted considerable

attention and consideration for use in fabricating microgels owing to its

outstanding biocompatibility, biodegradability, low toxicity and bio-adhesive

nature in diverse applications such as drug delivery, functional coating, tissue

regeneration and filtration. In the field of drug delivery, biodegradable and

biocompatible chitosan-based microgels were introduced to achieve the

encapsulation and release of the entrapped drug. For functionalizing textiles with

the ability to control moisture, thermoregulation and antimicrobial activity,

chitosan-based microgels incorporating antimicrobial agents have been applied

for functionalizing cotton fabric. Chitosan-based microgels have also been

applied to modify composite materials to improve tissue regeneration in the field

of tissue engineering. Moreover, chitosan-based microgels with good adsorptive

properties due to amino and hydroxyl functional groups can be developed to

remove environmental pollution.

1.3.1 Drug Delivery

Cancer has become a major worldwide health problem126. During the cancer

chemotherapy, the use of conventional anti-tumor agents is limited during by

their poor solubility, high toxicity, narrow therapeutic window, and serious side

effects to normal tissues due to their non-specific sites of action, which might

lead the cancer treatment to fail127. Therefore, a targeted drug carrier system has

been widely applied to encapsulate a large number of drugs, enhance the

1. Introduction

30

therapeutic effects of anticancer drugs, diminish the undesirable effects, and

specifically deliver them to tumor cells for cancer treatment128.

For cancer therapy, the tumor microenvironment, which is different from that

in normal tissues, is considered to be one of the important factors for designing

new therapies. Therefore, chitosan-based microgels can be designed as stimuli-

responsive systems that respond to stimuli from the tumor microenvironment to

achieve drug delivery129. Due to their stimuli-responsive behaviors of swelling-

deswelling transitions, drugs can be encapsulated into the nanocarriers and then

released from the interior of the carriers when their volume changes. Therefore,

stimuli-responsive microgels were investigated for use as drug delivery systems.

Microgels loaded with drugs can be synthesized and functionalized, and their

volume transitions can be tailored to trigger the release of drugs from the

particles in the presence of external triggers including external stimuli such as

pH or temperature changes130, ionic strength131, ultrasound132, magnetic fields133,

electrical effects134 and irradiation, or biological stimuli such as interactions with

enzymes and proteins135.

Hu et al. synthesized the novel prodrug conjugates, carboxymethyl chitosan-

carboxybenzaldehyde-doxorubicin nanoparticles (CMCs-CBA-DOX NPs),

which can release DOX in the acidic environment of the tumor cells through

passive targeting136. These acid-sensitive passive targeting drug release systems

were formed by self-assembling the amphipathic polymeric drug conjugates, in

which a carboxymethyl chitosan polymer was applied as a carrier, and p-

carboxybenzaldehyde (p-CBA) was used as a micro molecule linker connecting

to DOX through the formation of an amide linkage. Cellular uptake and the

release of DOX were investigated. As shown in Fig. 6, CMCs-CBA-DOX NPs

enter the body via an intravenous infusion and then disperse into the tissues via

the blood circulation. The drug-loaded NPs enter into the tumor cells through

endocytosis. In the presence of the acidic local environment, the imine bond

1. Introduction

31

between the drug (DOX) and the carrier (CMCs) is cleaved, thus triggering the

release of the drug.

Fig. 6. Schematic illustrations of pH-dependent drug release of CMCs-CBA-DOX in

vivo. Reprinted with permission from Ref. [136]. Copyright 2005 American Scientific

Publishers.

It is indicated that drug-containing nanocarriers which exhibit long-

circulating times or stimuli-responsive behaviors can passively accumulate in

the tumor site due to their enhanced permeability and retention (EPR) effect137.

EPR concept was introduced by Maeda et al. in the late 1970s138. They

discovered that macromolecular drugs selectively accumulated in tumor tissues.

The passive accumulation of nanocarriers in tumor sites was ascribed to the

leaky architecture of the tumor vasculature with its disorganized endothelium of

tumor vessels and poor lymphatic drainage system. From then on, a large

number of studies have operationalized this concept for drug delivery systems.

1. Introduction

32

Based on the EPR effect, long-circulating nanocarriers have been investigated

as a means to enhance drug accumulation, representing a great opportunity to

reach the targeted tumor tissues. PEGylated chitosan nanoparticles have been

designed and investigated as long-circulating carriers to realize diverse drug

delivery. The surface of the chitosan-based nanoparticles that have been

modified with poly(ethylene glycol) (PEG) can not only increase physical

stability but also decrease the surface charge of the particle. They have been

applied to achieve a prolonged circulation time in blood and enhanced

accumulation of the drugs139. The long-circulating polyelectrolyte nanoparticles

(PENPs) based on two different polysaccharides (hydrochloride chitosan (HCS)

and hyaluronic acid (HA)), were prepared by Wang et al.140. The PNPs were

synthesized through the electrostatic interactions between positively charged

amino moieties of CS and negatively charged carboxyl groups of HA, coated

with methoxy poly(ethylene glycol) (mPEG) through hydrogen bonding and

Van der Waals forces. The mPEG coating on the surface of PENPs could provide

steric hindrance against the non-specific mononuclear phagocyte system (MPS)

clearance to facilitate the PENPs reaching the target site and triggering HA-

mediated cellular uptake. HA can interact with cell-surface receptors, such as

CD44 receptors. Therefore, HA-based PENPs could enhance the specificity of

drug treatments for tumor cells and accumulate in tumor tissues with high levels

by pathways of receptor-oriented endocytosis. In addition, mitoxantrone

hydrochloride (MTO) was chosen as a model drug, as it has been shown to be

successfully encapsulated into the PENPs. These MTO-loaded drug delivery

systems could be applied for the treatment of advanced breast and prostate

cancers, lymphoma, and leukemia (Fig. 7).

In order to deliver drugs into tumors with more specificity, it has been

suggested that receptor-specific ligands are conjugated onto the drug-loaded

carriers to overcome the obstacles and enable the therapeutic agents to reach the

1. Introduction

33

targeted sites. This will depend on the binding affinity between nanocarriers and

the specific antigens or receptors which were overexpressed at the targeted sites,

e.g., cancer cells, resulting in the active targeting ability141.

Fig. 7. Schematic illustrations of the fabrication of PEGylated PENPs as drug delivery

carriers in MCF-7 cells. Reprinted with permission from Ref. [140]. Copyright 2018

Elsevier Science Ltd.

There were a variety of ligands that can be utilized in the active targeted

delivery system. Examples of various ligands have been reported, such as

biotin142, folic acid (FA)143, galactose (Gal)144, hyaluronic acid (HA)145,

glycyrrhetinic acid (GA)146 and lactobionic acid (LA)147. The researchers

1. Introduction

34

conjugated these receptor-specific ligands onto the microgel surface for

selective targeting to treat a specific disease or specific tumor cells.

Fig. 8. Schematic illustration of the synthesis of VP-16-encapsulated FA-CS-g-

PSBMA nanoparticles for tumor targeting delivery. Reprinted with permission from

Ref. [148]. Copyright 2016 the Royal Society of Chemistry.

A considerable amount of research has been devoted to the study of efficient

chitosan-based nanoparticles for drug delivery. Hua et al. first introduce self-

assembled chitosan (CS)-based nanoparticles coated with folic acid (FA) and

poly(sulfobetaine methacrylate) (PSBMA) for use in tumor-specific drug release

systems148. FA was applied as the active targeting moieties because the folate

receptor (FR) is overexpressed on the many epithelial tumor cell membranes,

such as in ovary, kidney, colon, prostate and lung cells. Therefore, after binding

with FR, FA-conjugated nanoparticles can be successfully internalized into

tumor cells by FR-mediated endocytosis (Fig. 8). The prepared nanoparticles

can encapsulate etoposide (VP-16), a widely-used chemotherapy drug, into the

inner hydrophobic core and release higher amount of the drug in an acidic

1. Introduction

35

phosphate-buffered saline than in neutral environments. Moreover, the drug-

loaded nanoparticles can be effectively internalized into HeLa cells. These

results suggest that the prepared FA-CS-g-PSBMA nanoparticles could be

applied as an active targeting nanocarrier in an anti-tumor drug delivery system.

Fig. 9. Schematic illustration of dual-ligands core/shell nanogels for active targeting of

hepatocellular carcinoma cells. Reprinted with permission from Ref. [149]. Copyright

2020 Dove Medical Press.

1. Introduction

36

Hefnawy et al. introduced a novel dual-ligand functionalized core-shell

chitosan-based nanocarrier for the treatment of hepatocellular carcinoma (HCC)

with an active targeting system. In this research, positively charged DOX was

complexed with negatively charged carboxymethyl chitosan-g-poly(acrylate)

through electrostatic interactions. A positively charged dual-ligand (lactobionic

acid and glycyrrhetinic acid)-conjugated chitosan was then coated on the

complex. These dual-ligand systems can provide two targeting moieties. One of

them is lactobionic acid, which can be used for selective targeting and is based

on the binding to asialoglycoprotein (ASGP) receptors, which are over-

expressed on the surface of HCC cells. Another ligand is glycyrrhetinic acid,

which can bind to the over-expressed surface proteins on an HCC. The

developed active targeting system can be used to achieve HCC-targeted delivery

of DOX (Fig. 9)149. Therefore, as a natural biodegradable polymeric material,

chitosan was chosen as an ideal candidate for preparing enzymatically

degradable chitosan-based micro- or nanogels that could be designed for

biomedical applications, such as controlled drug release.

1.3.2 Functional Coatings

A variety of physical and chemical methods have been investigated to

functionalize textile materials and endow them with enhanced protective

properties, thus combing the comfort of apparel with thermoregulation and

moisture management behaviors150. One of the innovative strategies used is to

incorporate a thin layer of a surface-modifying system, such as stimuli-

responsive microgels151. These reversible swelling/de-swelling properties of

particles can be responsive to changes in the environment. When applied to a

textile, they could dictate whether restrain or release vapor from the body by

decreasing or increasing the porosity of the textile material, thus allowing body

1. Introduction

37

vapor to be blocked or released152.

Chitosan-based microgels can be exploited for functionalizing textiles that are

able to achieve moisture management and thermoregulation activity. Moreover,

they can also be applied to textiles in combination with various antimicrobial

agents153 for the absorption and release of active substances as delivery media in

the field of medical textiles154.

Brigita Tomšič et al. prepared a stimuli-responsive cotton fabric using

temperature and pH-sensitive poly-N-isopropylacrylamide and chitosan

microgel (PNCS) encapsulating antimicrobially active 3-(trimethoxysilyl)-

propyldimethyloctadecyl ammonium chloride (Si-QAC), forming a bio-barrier

on the fiber surface35, 155. Si-QAC was applied to determine the antimicrobial

activity of the cotton fabric antibacterial to resist two types of bacteria, Gram-

positive Staphylococcus aureus and Gram-negative Escherichia coli. The results

show that PNCS microgel-functionalized cotton fabric is a smart stimuli-

responsive fabric that exhibits increased wearing comfort with simultaneous

moisture management and thermoregulation ability, and excellent antimicrobial

activity. They also developed a smart textile with silver embedded into a

temperature- and pH-responsive microgel for the control of antimicrobial

activities151. These PNCS microgels enable the release of silver triggered by

temperature and pH changes in the environment, which endowed the cotton

fabric with excellent antimicrobial activity against Gram-negative E. coli (>

99%) and Gram-positive S. aureus (> 85%).

Moreover, chitosan-based microgels can be exploited for coating on fabric for

self-cleaning application. Simoncic Barbara et al. prepared a novel stimuli-

responsive polyamide 6 (PA6) fabric with ZnO incorporated poly-(N-

isopropylacrylamide)/chitosan (PNCS) microgel coating for photocatalytic self-

cleaning156. The results showed that PNCS microgel coating with ZnO exhibited

temperature- and pH-responsive moisture management. In addition, in the

1. Introduction

38

presence of ZnO on the coating, the fabric exhibited UV protection and

photocatalytic self-cleaning properties.

1.3.3 Tissue Regeneration

Over the past decades, composite materials have been applied in the field of

tissue engineering to improve tissue regeneration, e.g., strontium-graphene

oxide (Sr-GO) nanocomposites157. However, the resistance of composite

materials to the human body limited their application in clinical research.

Paramagnetic or superparamagnetic nanoparticles (MNPs), such as

gadopentetate dimeglumine (Ga-DTPA)158, copper sulphide (CuS)159 and

iron(II,III) oxide (Fe3O4)160, could be applied as magnetic resonance imaging

(MRI) contrast agents to contribute to electronic stability as well as

pharmacodynamics and relaxivity. Although Fe3O4 MNPs have been widely

developed as MRI agents, they are nevertheless sensitive to magnetization and

oxidation161. Therefore, a superficial coating is essential for protection and

stability.

As a natural, renewable, non-toxic, biocompatible and biodegradable

compound, chitosan has attracted extensive attention for use coating the core of

metal oxides. Cui et al. exploited a multifunctional nanoprobe based on chitosan-

modified Fe3O4 nanoparticles for osteochondral magnetic resonance (MR)

diagnosis and regeneration162. The superparamagnetic nanoparticles Fe3O4-

CS/KGN MNPs were obtained through the self-aggregation of chitosan-grafted-

Fe3O4 oleic acid (Fe3O4-CS) and kartogenin (KGN). T2-weighted imaging using

Fe3O4-CS/KGN MNPs in vivo was employed to conduct the investigation. As

shown in Fig. 10A, the MRI results indicated that regarding the recovery from

the control, KGN alone and Fe3O4-CS/KGN groups were almost complete after

12 weeks of restoration. Compared with the control and KGN-treated group, the

1. Introduction

39

MRI results from the Fe3O4-CS/KGN group showed that the new cartilage layer

was integrated and lubricated, and new bone trabecula was reconstituted. From

the micro-computed tomography (micro-CT) diagnosis (Fig. 10B), part of the

freshly formed bone trabecula was mineralized under the cartilage, indicating

that Fe3O4-CS/KGN MNPs did not inhibit the osteochondral

reconstruction. These novel magnetic particles provided a noninvasive approach

for in vivo therapeutics of complex joint cartilage damage.

Fig. 10. MRI and micro-CT diagnose in vivo. (A) The T2-weighted MR images (red

circle: defect site; blue arrow: edema signals; green arrow: newly formed

cartilage). (B) micro-CT images of rabbit knees with Fe3O4-CS/KGN MNPs treatment

(W: weeks). Reprinted with permission from Ref [162]. Copyright Ivyspring

International Publisher.

For facial defects resulting from trauma, it is essential to repair adipose tissue

during the patient’s rehabilitation process163. Adipose tissue engineering

exhibits a great potential for repairing damaged adipose tissue164. Yin et al.

developed an injectable stem cell laden open porous microgel PLGA-g-HEMA

for adipose tissue regeneration165. Based on double-bonded poly(L-glutamic

acid)-g-2-hydroxyethyl methacrylate (PLGA-g-HEMA) and maleic anhydride-

javascript:;

1. Introduction

40

modified chitosan (MCS), PLGA-g-HEMA was synthesized using a water-in-

oil (W/O) emulsion method. The results for the neo-generated adipose tissues

were evaluated in vivo. The H&E staining treated with the PLGA-g-HEMA

group showed that adipose tissues had been formed locally. The Oil red O

staining results also exhibited the red intracellular lipid accumulation. After 12

weeks, the PLGA-g-HEMA group showed a ring-like morphology and vacuole

structure, further indicating that adipose tissue had formed. These results

demonstrated that these chitosan-based microgel systems showed excellent

potential for adipose tissue regeneration.

1.3.4 Filtration and Purification

Environmental pollution has become a global concern due to the disposal of

large amounts of water-soluble dyes. Most of the dye-bearing wastewater is non-

biodegradable and can pollute groundwater, thus posing a serious threat to

human life166. It is a major challenge to remove toxic dyes from the wastewater

and industrial effluents due to the fact that the dyes can be easily discharged into

the effluents in the environment167. Whist membrane technologies have been

applied to achieve dye removal, they still have drawbacks, such as inherent

fouling and disagreement between water flux and rejection which limits their

application in industrial wastewater treatment168. To overcome these hindrances,

a variety of materials (e.g., inorganic nanoparticles, functionalized hydrophilic

polymers and antibacterial agents) have been used169.

As a semi-crystalline, non-toxic biopolymer with a good adsorptive nature

due to its amino and hydroxyl functional groups, chitosan has been extensively

applied in membrane technologies170. Moreover, it is indicated that the

bioactivity of chitosan nanoparticles can be further enhanced through the

incorporation of metal ions such as Ag+, Cu2+, Zn2+, Mn2+ or Fe2+ 171. Therefore,

1. Introduction

41

chitosan-based microgels can be studied to discern their potential for resolving

membrane fouling issues. In order to improve the membrane hydrophilicity and

other physicochemical properties, the synergistic interaction of nanomaterials

with the polymer chains regulated the fouling tendency.

Due to their excellent antimicrobial properties, Asiri et al. incorporated

chitosan-based nanoparticles into nanocomposite to fabricate hollow-fiber

membranes172. The chitosan and silver-loaded chitosan nanoparticles were

prepared by ionic gelation to fabricate hollow-fiber membranes using a dry-wet

spinning technique. The prepared nanocomposite hollow-fiber membranes

displayed a superior anti-biofouling performance. The anti-biofouling study

showed that by incorporating 0.30 wt% of the chitosan and silver-loaded

chitosan nanoparticles, the antifouling properties of the hollow-fiber membranes

with a flux recovery ratio were enhanced to 81.21 and 86.13%, respectively. The

dye rejection study demonstrated that the nanocomposite membranes showed a

maximum rejection of 89.27% and 86.04% for Reactive Black 5 and Reactive

Orange 16, respectively. The presence of Ag+ in silver-loaded chitosan

nanoparticles further improved the microbial inhibition which was tested for

biofilm inhibition property using model strains of bacteria such

as Mycobacterium smegmatis, Staphylococcus aureus, and Escherichia

coli. Therefore, the nanocomposite hollow-fiber membrane with silver-loaded

chitosan nanoparticles can be applied in the treatment of industrial dye effluents.

In addition, wastewater treatment takes a long operation time and thus

requires preparation of stable antifouling nanomaterials for preventing biofilm

formation. Sujoy K. Das developed an environmentally benign facile synthesis

process to synthesize a core-shell magnetic chitosan microsphere coating with

silver nanoparticles (MCSM) as a smart antifouling nanomaterial for efficient of

dyes and microbial contaminants removal (Fig. 11A)173.

https://pubs.acs.org/action/doSearch?field1=Contrib&text1=Sujoy+K.++Das

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42

Fig. 11. (A) Schematic representation of the silver nanoparticles synthesis and easy

separation using external magnetic field leading to recycling and reuse. (B) Chemical

structures of different anionic and cationic dyes (AB-113, BCG, BPB, CR, EY, SB,

SDB and Y-5GN); color images of dye solution before and after treatment and

1. Introduction

43

percentages of dye adsorption by silver nanoparticles at different pH values (2.0-10.0).

Reprinted with permission from Ref. [173]. Copyright 2015 American Chemical

Society.

As shown in Fig. 11B, eight different commercially used dyes, including both

cationic and anionic dyes, were treated with MCSM at low and high pH

values. The results showed that MCSM exhibited pH-dependent adsorption

properties. Almost 99% of anionic dyes (AB-113, BCG, CR, EY, SB, SDB, Y-

5GN) was removed at a low pH range (2.0-4.0), whereas the cationic dye (BPB)

was removed at higher pH values (pH > 8.0). Moreover, the bacterial growth

inhibition ability of MCSM was assessed against E. coli and P.

aeruginosa using turbidity measurement. The bacterial growth kinetics

indicated that MCSM completely inhibited the growth of E. coli and P.

aeruginosa, indicating the stellar antibacterial properties of MCSM. The core-

shell MCSM provided an environmentally sustainable technology for eco-

friendly and cost-effective water purification.

1.4 Aim and Motivation

The major challenges for controlled drug delivery systems are the

biocompatibility and stability of the delivery systems. To achieve efficient

delivery of therapeutics into tumor cells, it is suggested that delivery vehicles

composed of naturally occurring systems are applied to overcome these

challenges174.

Herein, the aim of the Thesis is to design and prepare biopolymer-based

microgel systems with good biocompatibility, pH-sensitivity and

biodegradability. Due to their unique properties such as non-toxicity,

biodegradability, and biocompatible behaviors, as well as reactive functional

1. Introduction

44

groups, chitosan has been introduced to fabricate microgels in the biomedical

and pharmaceutical fields. However, the limited solubility of chitosan at

physiological pH values creates challenges in drug delivery utilization. The

modifications on its amino and hydroxyl groups enable chitosan to be imparted

with new properties, thus achieving a specific biomedical purpose. With

functionalization on its chain, the designed biocompatible and biodegradable

chitosan-based microgels could be developed as the drug carriers for the

encapsulation and site-specific controlled release of therapeutics.

1.5 Scope of the Thesis

Chapter 1 gives an overview of the functional microgels which can be

fabricated by two systems: biopolymer-based systems and conductive polymer-

based systems. The properties of biopolymers and conductive polymers offer the

functional microgels various properties, such as pH-sensitivity, conductivity and

biodegradability. As one of the unique polysaccharides, chitosan can be utilized

as a scaffold material in manufacturing microgels, opening up many potential

applications such as drug delivery, functional coatings, tissue regeneration,

filtration and purification.

Chapter 2 describes the synthesis of a redox-active chitosan-based microgel

for controlled drug delivery. Using chitosan as a matrix and poly(hydroquinone)

as the redox-active polymer, a series of microgels were prepared with a tunable

ratio of chitosan:poly(hydroquinone), with the obtained microgels showing pH-

and redox-responsibility. Moreover, the prepared microgels can encapsulate

DOX to be released in the presence of lysozyme in an acidic environment, which

could be applied to carriers in a controlled drug delivery system.

Chapter 3 introduces the biodegradable microgels in which chitosan was

applied as a matrix, poly(aniline) was grafted on the matrix to introduce

1. Introduction

45

conductivity, and glutaraldehyde was used as a cross-linker. These microgels

possessed the pH-sensitivity and redox-activity, and can be degraded at a high

rate in the presence of lysozyme at pH 6, presenting good biodegradability.

Chapter 4 introduced a series of biodegradable pH-responsive microgel based

on modified biopolymers, alkyne-modified chitosan and azide-modified dextran,

cross-linked via “click chemistry” without any extra cross-linkers. In addition,

the microgels can be degraded in the presence of model dextranase, an enzyme

present in the colon. It can also encapsulate an antibiotic, VM, and release it in

a controlled manner, suggesting that such “smart” microgels have great potential

for biomedical applications as drug carriers for targeted therapies in the colon.

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1. Introduction

58

2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels

59

2. Redox-Active Supramolecular Poly(hydroquinone)-

Chitosan Microgels

This Chapter has been reproduced from Helin Li, Olga Mergel, Puja Jain, Xin

Li, Huan Peng, Khosrow Rahimi, Smriti Singh, Felix A. Plamper and Andrij

Pich, Soft Matter, 2019, 15, 8589-8602. Copyright Royal Society of Chemistry.

Reproduced with permission.


60

2.1 Introduction

Since poly(N-isopropylacrylamide) (PNIPAM) microgels were firstly

prepared by Pelton and Chibante in 1986, stimuli-responsive microgels have

received a great deal of interest, particularly in the last decade1. Due to their

unique properties which endow them with tailor-made properties and

sensitivities in response to external stimuli, such as temperature and pH, these

so-called “smart microgels” have been extensively investigated for controlled

drug delivery, tissue engineering, and enzyme or protein modification. Notably,

a lot of research has been conducted on PNIPAM microgels to investigate their

potential use for drug delivery. These PNIPAM microgels are thermosensitive

and exhibit a reversible coil-to-globule transition at their lower critical solution

temperature (LCST) of approximately 34 °C in an aqueous solution, which is

close to human body temperature. However, the application of PNIPAM-based

microgels has been limited due to their deficient biodegradability and

biocompatibility, such that their use may result in cytotoxicity in the body.

Therefore, to develop biomedical applications, intensive efforts focusing on

microgels concentrated on a range of natural polymers including chitosan2,

dextran3, gelatin4, inulin5, starch6, and sodium alginate7, etc., to overcome this

limitation8.

Natural polymers, obtained from renewable resources, such as polysaccharide,

protein, collagen, carrageenan, cellulose, hyaluronic acid, are increasingly

reported as biopolymers that have a wide range of applications when used to

prepare microgels for clinical use9. These materials are biodegradable,

biocompatible and non-toxic. Among these interesting natural polymers,

chitosan is a typical example that has been studied and subjected to chemical

modification with other synthetic stimuli-responsive polymers. Chitosan,


61

consisting of b-(1,4)-2-acetamido-2-deoxy-D-glucose units, is a weak cationic

natural polysaccharide that is obtained through the deacetylation of chitin10.

Specifically, it is a pH-responsive polymer due to its large number of amino

groups. Upon protonation or deprotonation of amino groups in chitosan, a phase

transition is induced with the increased or decreased hydrodynamic volume of

the polymer in solution. When the polymer is presented within the microgel

network, the variation of the microgel volume is triggered due to the pH-induced

phase transition11. Due to its biodegradability and biocompatibility, chitosan is

an ideal biomedical material that can be applied in several fields, such as targeted

drug delivery and release and tissue engineering.

In the last decade, the increased demand for renewable and sustainable energy

resources has meant that a lot of efforts have been directed towards the

development of power storage and delivery systems. Conducting polymers,

termed “synthetic metals”, have been widely used in electrochemical fields

owing to their intrinsic electronic properties that are inherent to metals or

semiconductors. The conductivity of these materials comes from the conjugated

double bonds over the polymeric chains, thus leading them to exhibit electronic

properties12.

The mechanisms of polymerization of conducting polymers are the oxidation

during an electrochemical process, including coupling and proton elimination.

Firstly, a radical cation appears due to the oxidation of one monomer, and forms

a dimer with another radical cation after the loss of two protons. This dimer can

then be further oxidized and coupled with another radical cation, and then

oligomers appear. This process will continue until the polymer is obtained.

However, different conductive polymers will undergo different polymerization

processes13.

Among these conductive polymers, redox-active polymers play an important

role in the application of electrochemical devices. Conductive polymers can be


62

reversibly oxidized or reduced due to their redox sites. Through the electron

exchange reaction, which takes place between the loss of electrons (oxidation)

or the reception of electrons (reduction), the electrons can be delivered14. The

ion and electron transfer via oxidation or reduction process can result in the

creation of electrochemical capacitors, also called pseudo-capacitors, in which

the allocated charge can be recycled by reduction during the oxidation process15.

Due to their unique properties, such as high electrical storage capacity and fast

charge-discharge character, these capacitors can be applied in electrical fields16.

Depending on their distinctive biomedical and electrochemical properties, these

polymers have an interesting range of different applications, such as in the

design of electrochemical devices for drug release profiles, and new types of

actuators17.

Based on the above-mentioned concepts, a redox-active polymer with a

conjugated backbone has attracted our attention. Moreover, this polymer

combines both intrinsic conductivity and redox activity due to its specific

property, that is, every unit of the polymer can be reversibly oxidized or reduced.

This enhances the addressability of redox-active sites of the polymer within

electroactive colloids. Therefore, this redox-active polymer can be employed to

enhance the electroactivity of the colloids. Additionally, it can also be used in

flow-cell batteries, biosensors, biofuel cells, and electrochromic devices18. The

size and payload release of the colloids could also be altered by adjusting the

redox state of the matrix. For such applications, we will focus on a redox-active

polymer, poly(hydroquinone), which has a conjugated backbone with high

electrochemical activity. Due to these characteristics, it could be used as a novel

redox-active polymer in electrochemistry19. We conducted poly(hydroquinone)

into the biodegradable microgel system in a facile way through physical cross-

linking. These microgels also have a potential application in the drug release

profile.


63

The aim of this work was to synthesize a redox-active microgel by

polymerizing hydroquinone in the presence of chitosan, inspired by the work of

Jian He et al20. Next, we advanced this approach to synthesize a redox-active

microgel in which chitosan was used as a matrix and poly(hydroquinone) was

the redox-active polymer. The microgels were physically cross-linked in an

inverse miniemulsion system. Herein, we developed a new colloidally stable

microgel with dual responsiveness: pH and redox-responsiveness. A series of

microgels were synthesized with a changeable ratio of

chitosan:poly(hydroquinone). In addition, these microgels are sensitive to the

equilibrium potential which was tuned using a bulk electrolytic approach, that

is, they could be triggered to alter from a swollen state to a shrunken state in an

electrochemical cell. The obtained microgels were degradable and in the

presence of urea or enzymes, they can be degraded into small fragments. This is

because urea can disrupt hydrogen bonds, and thus, disrupt the physical cross-

links in microgels. Moreover, an enzyme, lysozyme, can also degrade microgels

by cleaving the glucosidic linkages in the polysaccharide backbones of chitosan.

The degradable microgels could also be used as drug release devices. They

encapsulated doxorubicin (DOX), which could be released from the microgel in

the presence of lysozyme. Due to its drug release properties, the goal of our work

is to develop such an electroactive microgel to provide the impetus for further

development in diverse fields, such as tissue engineering, cell therapy and

energy storage.


64

2.2 Experimental Section

2.2.1 Materials

All reagents were purchased from commercial suppliers and used without

further purification unless chitosan. Hydroquinone ( ≥ 99%), acetic acid (99%),

glutaraldehyde (25% in H2O), cyclohexane ( ≥ 99.8%), ammonium hydroxide

solution (NH3·H2O, ≥ 25% in H2O), Span 80, dibasic potassium phosphate

(K2HPO4, ≥ 98%), monobasic potassium phosphate (KH2PO4, ≥ 98%), acetone

( ≥ 99.5%), methanol ( ≥ 99.8%), sodium hydroxide (NaOH, ≥ 98%, pellets,

anhydrous), lysozyme from chicken egg white (protein ≥ 90%, ≥ 40,000

units/mg protein) and bovine serum albumin (BSA) were bought from Sigma-

Aldrich and used as received. Hydrochloric acid (HCl, 37%) was purchased from

VWR International GmbH. Doxorubicin hydrochloride (DOX, 98%) was

purchased from TCI. Chitosan, medium molecular weight (190-310 kDa,

Sigma-Aldrich), was used with further purification according to a literature21.

Dialysis membranes (MWCO = 1.2 kDa) were purchased from Carl Roth.

Deionized water was obtained as a reaction medium for all experiments, and also

used for the preparation of PBS buffers at pH range 5 to 8, and other pH values

were adjusted with 0.1 M HCl or 0.1 M NaOH.

2.2.2 Synthesis of Microgels

The microgels were synthesized by inverse miniemulsion polymerization

using cyclohexane as an organic phase and acetic acid as an aqueous phase. In

an aqueous phase, chitosan (0.012 g) and hydroquinone with changed amounts

from 0.008 g to 0.072 g were dissolved in 1 mL 0.1 M acetic acid. In an organic

phase, Span 80 (0.258 g) was dissolved in cyclohexane (10 mL). The aqueous


65

phase and the organic phase were mixed and ultrasonicated using a Branson

Sonifier 450 (duty cycle of 50%, and output control of 40%) under ice cooling

for 10 minutes. Exposed to the air, the reaction mixture was stirred with a

condenser for 9 hours at 60 °C. Afterward, a series of microgel samples were

prepared with different mass ratios of chitosan to hydroquinone. The ratio was

changed from 1:0.33 to 1:3 by increasing the amount of hydroquinone. After the

reaction, the microgel dispersion was centrifuged for 20 minutes at 6000 rpm.

The supernatant was discarded and 10 mL of cyclohexane was added. This

process was repeated 3 times. The final precipitate was dispersed in 5 mL of

deionized water and dialyzed against water for further purification. The microgel

samples need to be freshly prepared each time in order to take subsequent

measurements. Sedimentation occurs when there is longer standing time in both

Tris and PBS buffers.

2.2.3 Degradation of Microgels

2.2.3.1 Degradation of Microgels by Urea

To quantify the degradation behavior of microgels in the presence of urea, the

size and morphology of microgels during degradation were investigated by

taking dynamic light scattering (DLS) and transmission electron microscopy

(TEM) measurements. As a hydrogen bond breaker, urea can be applied to

degrade microgels. Degradation of the microgels was carried out by evaluating

the altered size and morphology of microgels in the presence of urea through

DLS and TEM measurements.

To investigate the variation in the particle sizes during degradation, three

microgel samples were observed by measuring the variation of hydrodynamic

radius monitored by DLS. The obtained mixture used for DLS measurements


66

was produced by mixing the microgel dispersion and urea solution. The final

concentration of urea was kept at 8 M. At regular time intervals, the DLS

measurements proceeded at shorter time intervals within the first 400 minutes

and longer intervals from 1 day to 1 week. The microgel dispersion was under

vigorous stirring throughout the experiment.

The changed morphologies of the microgels during degradation were

observed by TEM. The samples used for TEM measurements were prepared by

mixing the original microgel sample dispersion with urea solution. The

concentration of urea solution was kept at 8 M and the mixture was under

vigorous stirring for the duration of experiment. At predetermined time intervals,

the mixture was withdrawn and dropped on a TEM grid. It was dried at room

temperature before measuring.

2.2.3.2 Degradation of Microgels by an Enzyme

Enzymatic degradation was conducted by monitoring the changed size,

weight and morphology in the presence of an enzyme, lysozyme, via DLS,

weight loss and TEM measurements. Lysozyme can disrupt the glucosidic

linkage of chitosan, thus resulting in the collapse of the microgel. On this basis,

it can be used as a model enzyme for degrading microgels.

To investigate the extent of enzymatic degradation, DLS and TEM

measurements were conducted by mixing the microgel dispersion with lysozyme

solution. The concentration of lysozyme was kept at 10 mg/mL. During the

whole testing process, the mixtures for measuring were constantly stirred and

their temperature kept at 37°C. The following procedures for DLS and TEM

measurements were monitored according to the process of urea-induced

degradation, as mentioned above. Then, the degradation process was

investigated by tracking the weight loss of microgels in the presence of the

enzyme. The measurements were as follows. 10 mg of dried microgel particles


67

were dispersed in 5 mL of buffer (pH = 6) and mixed with lysozyme to obtain a

concentration of 10 mg/mL. The mixture was transferred and incubated at 37°C

under continuous stirring. The sample was withdrawn at predetermined time

intervals. Further degradation was terminated by adjusting the environmental pH

value from 6 to 3 with 0.1 M HCl. This was because lysozyme’s active pH range

is from 6 to 9. All samples were dialyzed overnight, lyophilized and weighed

(wt). The weight loss ratio was calculated by Equ. (1):

Weight loss (%) = W0−Wt

W0 × 100 (1)

Where W0 is the initial weight of the microgel samples, and Wt is the weight of

the dried microgel samples after degradation as a function of incubation time t,

respectively. All analyses were carried out in triplicate.

2.2.4 Drug Loading and Release Studies

DOX, an anticancer drug, was chosen as a model drug to study the drug

loading and release profiles of microgels. The drug loading efficiency was

investigated by mixing 5 mg of dried microgels and 1 mg of DOX in 5 mL of

deionized water and stirring it overnight at room temperature. It was then

transferred to a dialysis bag to remove the free DOX. The DOX loading content

was quantified by a UV-Vis spectrometer at a wavelength of 496 nm, based on

the calibration curve of a series of standard DOX solutions constructed in

deionized water. The drug loading efficiency was quantified by measuring the

ratio of UV-Vis intensity of DOX, in the media of microgel dispersion after

dialysis, to total UV-Vis intensity within microgels before dialysis according to

Equ. (2):

Drug loading efficiency (%) = Md −Mf

Md× 100 (2)


68

Where Md is the initial weight of DOX, and Mf is the weight of free DOX,

respectively.

To release the loaded DOX, the drug release behaviors of the microgels were

carried out in the presence of lysozyme. The procedures were as follows. Firstly,

the DOX-loaded microgel dispersion and lysozyme solution were mixed

together whilst the concentration of lysozyme was kept at 10 mg/mL. The

mixture was subjected to vigorous stirring overnight at 37 °C to achieve

complete degradation. To release the loaded DOX, 1 mL of this mixture was

dialyzed against 9 mL of PBS buffer (pH = 6) in an incubator. It was then placed

on a shaker kept at 37 °C. At predetermined time intervals, 1 mL of released

PBS buffer was taken out for UV-Vis analysis to determine the amount of the

released free DOX. The buffer removed for sampling was then replaced by 1 mL

of fresh buffer to maintain a 10 mL incubation volume. The amount of DOX

released from dialysis bags was measured using UV-Vis spectrometry. The

amount of released DOX was quantified from the calibration curve of a series of

DOX solutions constructed in a PBS buffer at pH 6. DOX-loaded microgel

solutions with no lysozyme incorporated were also tested under the same

conditions according to the same procedures as described above to be used as a

control. All analyses were carried out in triplicate.

2.2.5 Electrochemical Assay

To evaluate the electrochemical behaviors of microgels, cyclic voltammetry

(CV) measurements were taken by an electrochemical workstation potentiostat

CHI760D (CH Instruments, Austin, Texas, USA). The measurements were

performed by scanning the potential in the respective potential window (-2 V ~

2 V) at a scan rate of 1 V s-1 at room temperature. A glassy carbon (GC) electrode

or a platinum (Pt) electrode was used as the working electrode, and an Ag/AgCl


69

electrode was used as the reference. A platinum wire electrode was used as a

counter electrode. Before conducting the measurement, the working electrode

was prepared via mechanical polishing. It was then rinsed with water and dried

with a stream of argon. The microgels were dispersed in a 0.05 M phosphate

buffer (pH = 7.3) and purged with argon for 10 minutes to remove any dissolved

oxygen. The CV experiments were performed in a standard three-electrode setup.

The three electrodes were put together in a cuvette where the microgels

dispersed in the PBS buffer (pH = 7.3) were stored. A platinum gauze electrode

(35 mm x 20 mm) served as the working electrode and attached to the bottom of

the cuvette. A platinum wire served as a counter electrode and was placed in

0.05 M PBS buffer (pH = 7.3) in a compartment separated by a diaphragm that

was also immersed in the microgel solution. An Ag/AgCl electrode stored in 1

M KCl served as the reference electrode. Bulk electrolysis measurements were

also performed using the same equipment.

Electrolysis experiments were undertaken at fixed potentials of 2 V for the

oxidation process and -2 V for the reduction process of the microgels. The

potential value was fixed due to the cyclic voltammetry data, which showed two-

step oxidation and reduction processes in some cases, being only sufficiently

activated below 2 V and above -2 V. Prior to the test, the microgel solution was

purged with argon for 10 minutes to remove any dissolved oxygen. After the

electrolysis measurements were taken, the microgel solution was extracted and

then quickly transferred to a glass cuvette for DLS measurements to analyze the

hydrodynamic radius of the microgels.

2.2.6 XTT Assay

The XTT cell proliferation assay of the microgels was monitored using the

XTT cell proliferation assay kit (Cat. No. 30-1011K, ATCC) based on the


70

manufacturer’s instructions. The targeted cells used for the experiment were the

mouse fibroblast cell line, L-929 (ATCC CCL-1). 1 × 104 cells per well were

seeded in a 96-well microtiter plate and precultured overnight at 37 °C under 5%

CO2 and 95% air incubator before assay. After removing the medium, the cells

were cultured in the medium containing microgels with different concentrations

of 0.100, 0.010 and 0.001 mg/mL per well. The mixtures were incubated in 5%

CO2 and 95% air for 24 hours at 37 °C. In order to determine the living cell

numbers, the XTT (sodium 2,3-bis-(2-methoxy-4-nitro-5-sulfophenyl)-5-

[(phenylamino)-carbonyl]-2H-tetrazolium) inner salt and PMS (N-methyl

dibenzopyrazine methyl sulfate) as electron carrier were employed according to

the supplier’s instructions. At the viable cell surface, XTT can only be reduced

by living cells to an orange water-soluble formazan dye with the assistance of

PMS. Formazan formation was quantified spectrophotometrically at 490 nm

(reference wavelength 630 nm) using a microtiter plate reader (Detection

Microplate Reader from BioTek). The analyses were performed in triplicate.

2.2.7 Characterization Methods

Fourier transmission infrared (FTIR) spectra were recorded by Nexus 470

(Thermo Nicolet) spectrometer. The freeze-dried microgel samples were pressed

into a KBr pellet at room temperature and analyzed using FTIR spectroscopy.

Attenuated total reflectance fourier transform infrared (ATR-FTIR)

spectrometry was collected using a Nexus 470 (Thermo Nicolet) which is

equipped with a smart split ATR single reflection Si crystal over the spectral

range of 4000-400 cm-1 with a resolution of 4 cm-1.

The hydrodynamic radius of the microgel in aqueous solution was studied

using a commercial laser dynamic light scattering (DLS) spectrometer

(ALV/DLS/SLS-5000) equipped with an ALV/LSE-5004 multiple tau digital


71

correlator and ALV/CGS-3 compact goniometer system with a helium-neon

laser (Uniphase 1145P, output power of 22 mW and wavelength of 632.8 nm)

as a light source set at a 90° scattering angle. Electrophoretic mobility of

microgels was carried out using Malvern Zetasizer Nano ZS. After equilibration

of 120 s, the measurements were carried out at 25 °C. Before measurements, all

samples were obtained and diluted with PBS buffers with a pH range of 3 to 11.

Transmission electron microscopy (TEM) analyses were carried out by Zeiss

LIBRA 120. The electron beam accelerating voltage was set at 120 kV. The

samples were produced by dropping the microgel solution onto a formvar-

carbon-coated copper grid with a mesh size of 400. Before preparing the samples,

the surface of the grid was pretreated with plasma for 120 s. A drop of the

microgels solution was placed on the grid after surface treatment. The grid was

then placed on filter paper and dried overnight at room temperature.

The colloidal stability of the microgels dispersions was determined by a

separation analyzer, LUMiFuge 114 (LUM GmbH, Germany). All the samples

were sedimented in polycarbonate cells at acceleration velocities of 4000 rpm at

25 °C. In order to measure the sedimentation velocity of microgels in the

corresponding solvent mixtures, the microgels in PBS buffers were investigated.

UV-Vis spectra were performed on a Perkin Elmer Lambda 35 UV-vis

spectrometer.

2.3 Results and Discussion

2.3.1 Synthesis of Microgels via Oxidative Polymerization

The synthesis of redox-active hydrogels through the oxidative polymerization

of hydroquinone using chitosan as a template has been reported by Jian He et

al20. In their study, the authors discovered that the color of the mixture of an


72

acidic solution of chitosan and hydroquinone would change from light yellow to

wine-colored. When exposed to the air, the mixture became increasingly viscous

until the hydrogels were prepared. This suggested that the hydroquinone, when

polymerizing to poly(hydroquinone), can form hydrogen bonds with chitosan.

Hence, a novel redox-active hydrogel was prepared.

Scheme 1. The preparation route of physically cross-linked microgels via the oxidative

polymerization20.

According to their work of redox-active hydrogel, we prepared biocompatible

and biodegradable microgels through inverse miniemulsion polymerization.

Firstly, the chitosan and hydroquinone were dissolved in the diluted acetic acid

(0.1 M) and were applied as a dispersed aqueous phase. Secondly, the organic

phase was prepared by adding the Span 80 in 10 mL of cyclohexane. Thirdly,

the water-in-oil (W/O) miniemulsion was prepared by sonicating the mixture of

the aqueous phase and organic phase prepared above. Following the same


73

procedure, a series of microgels, exhibiting the particular redox-active properties

shown in Scheme 1 and Table 1, were successfully prepared.

Table 1. Different microgel samples synthesized in this study.

Name of

sample

Mass ratio of

chitosan to

hydroquinone

Molar ratio of -NH2 of

chitosan to -OH of

hydroquinone

Chitosan

(g)

Hydroquinone

(g)

CHHQ-1 1:0.3 1:0.61 0.012 0.004

CHHQ-2 1:0.5 1:0.91 0.012 0.006

CHHQ-3 1:1 1:1.83 0.012 0.012

CHHQ-4 1:2 1:3.66 0.012 0.024

CHHQ-5 1:3 1:5.49 0.012 0.036

Under certain conditions, the redox couple presents considerable

electrochemical activity that includes the oxidation of hydroquinone and the

reduction of benzoquinone22, and undergoes a reversible two-electron, two-

proton oxidation/reduction process, wherein the redox couple provides the

charging and discharging approaches in the electrochemical reaction23.

2.3.2 Chemical Composition of Microgels

The FTIR spectrum of CHHQ microgels, chitosan and poly(hydroquinone)

were present in Fig. 1A. For the pure chitosan, the characteristic peaks located

in the region of 3200-3500 cm-1, ascribed to N-H stretching and O-H stretching

vibrations24. The asymmetric stretching of CH3 and CH2 of chitosan’s saccharide

structure were located at 2925 cm-1 and 2874 cm-1 25. The characteristic peaks at

1653 cm-1, 1552 cm-1 were ascribed to amides I and II, respectively26. The C-O

skeletal stretching from the saccharide structure of chitosan was observed at

1077 cm-1 and 1034 cm-1 27.


74

Fig. 1. (A) FTIR spectra of chitosan, poly(hydroquinone) and the different microgel

samples. (B) Linear fit of molar ratio of chitosan to hydroquinone and intensity ratio of

the C-O (1077 cm-1) to OH (1204 cm-1) IR bands present in the samples. (C) ATR-

FTIR spectra of the microgel (CHHQ-4) and poly(hydroquinone).

For the poly(hydroquinone), the absorption peaks at 1620 cm-1, 1501 cm-1

and 1450 cm-1 were ascribed to the vibrations of aromatic rings and C=C bonds

in the backbone of poly(hydroquinone)28. The peak located at 1204 cm-1 was

ascribed to the phenolic OH groups. The absorption peak at 822 cm-1 was

assigned to the out-of-plane vibrations of the C-H bond of the aromatic rings29.


75

Comparing the spectrum of chitosan, poly(hydroquinone) and the CHHQ

microgels, it was seen that the adsorption peaks of microgels included the

characteristic peaks of both chitosan and poly(hydroquinone). As shown in Fig.

1B and Table 2, the intensity ratio of C-O (1077 cm-1) to OH (1204 cm-1)

reflected a linear fit of the molar ratio of chitosan to hydroquinone. To verify

that the hydrogen bond was formed within microgels, ATR-FTIR spectra were

investigated. From the results shown in Fig. 1C, the absorbance of the peak at

3215 cm-1 was assigned to the stretching vibration of hydrogen-bonded O-H.

The absorption peak at 3361 cm-1 is the free O-H of poly(hydroquinone). The

absorption peak of O-H moved from 3361 cm-1 to 3215 cm-1, indicating that the

hydrogen bonds formed30.

Table 2. The synthesis of microgel particles with the different molar ratios of chitosan

to poly(hydroquinone).

Name of sample Molar ratio

(chitosan/hydroquinone)

Intensity ratio

(1077/1204)

CHHQ-1 0.180 0.340

CHHQ-2 0.270 0.500

CHHQ-3 0.550 1.220

CHHQ-4 1.092 2.340

CHHQ-5 1.640 3.360

2.3.3 Influence of pH on Microgel Size and Electrophoretic Mobility

The hydrodynamic radius (Fig. 2A) and electrophoretic mobility (Fig. 2B) of

microgels in PBS buffers at different pH values were investigated. As expected,

the pH of the aqueous solution affected the particles’ dimension. As shown in

Fig. 2A, the hydrodynamic radius of the microgels was measured as a function

of pH. The microgels swelled when exposed to an acidic medium due to the

protonation of amino groups. In the acidic environment, the amino groups (-NH2)


76

of chitosan were protonated (-NH3+) and carried net positive charges, thus

inducing the microgels to be positively charged. Subsequently, the microgels

swelled due to the intermolecular electrostatic repulsions and the osmotic

pressure of the uptaken counterions. When the environment changed from

slightly alkaline conditions to alkaline conditions, the microgels were negatively

ionized. At pH 11, the amino groups of chitosan were deprotonated and the

hydroquinone (HQ) units started to deprotonate to a singly (Q-) or totally

deprotonated (Q2-) state31. The intramolecular electrostatic repulsions and the

osmotic contributions were weak in the alkaline environment, resulting in the

shrinkage of microgels which caused the particle size to decrease.

In an aqueous solution, the volume of the microgels was affected by the

degree of cross-linking density, which was changed with the various mass ratios

of chitosan to hydroquinone, varying from 1:0.3, 1:0.5, 1:1, 1:2 to 1:3. The

number of physical cross-links increased with the increased content of

hydroquinone within the microgels. As shown in Fig. 2D, the swelling ratio of

microgels was influenced by the degree of cross-linking density, as well as pH

value. It is indicated that the microgels with the highest cross-linking density

will form the tightest network due to a large number of physical cross-links

within the microgels. The looser network a microgel contains, the higher

swelling or shrinking ratio the microgel exhibits. Moreover, the hydrophobic

forces will induce limited swelling where there is high hydroquinone amount

present.

As shown in Fig. 2C, the microgels swelled at pH 3 and collapsed at pH

10 within two circles, indicating that the pH responsiveness of microgels

is reversible.


77

Fig. 2. (A) Variation of hydrodynamic radius, (B) electrophoretic mobility, (C)

hydrodynamic radius by varying the pH between 3 and 10, (D) swelling ratio, and (E)

the photos of microgels as a function of pH.

In acidic media, microgel dispersions exhibited a slightly brownish

color as shown in Fig. 2E. In alkaline media, a dark brown color appeared

because of the complexation of hydroquinone and benzoquinone. Scheme

2 indicated that the hydroquinone was colorless, and benzoquinone and


78

quinhydrone were yellow and purple. Hydroquinone predominates in

microgels in acidic media, whist benzoquinone and quinhydrone

predominate in microgels when there is an increase in pH, inducing the

change in color.

Scheme 2. Complexation between hydroquinone and benzoquinone.

2.3.4 Colloidal Stability of Microgels

The colloidal stability behaviors of microgels were evaluated using

LUMiFuge at pH 3, 7, and 10. Fig. 3 indicates that the sedimentation

velocities of the microgels were affected by the chemical structure and pH

of the medium. These sedimentation velocities can be obtained by

analyzing the slope of the sedimentation curves.

It is probable that CHHQ-5 has the highest tendency to aggregate and

therefore the lowest colloidal stability, which is reflected in the

exceptionally high sedimentation velocity. The microgels sedimented

slower in pH 3 than in pH 7 and 10. This can be explained by the swelling

of the microgels and also that there are no strong indications for

aggregation, except for CHHQ-5.


79

Fig. 3. Sedimentation velocities of microgels at pH 3, 7 and 10.

2.3.5 Electrochemical Properties of Microgels

2.3.5.1 Cyclic Voltammetry

In order to investigate the electrochemical behavior of microgels, the CV

measurements (in 0.05 M PBS buffer pH 7.3, against an Ag/AgCl electrode)

were performed. In these redox-active microgels, chitosan acted as a matrix and

poly(hydroquinone) constituted the redox-responsive component. The redox

cycling of the hydroquinone-benzoquinone redox couple was measured using

CV with a GC electrode. In the redox process, hydroquinone (HQ) units along

the polymeric backbone can be oxidized to 1,4-benzoquinone (BQ) units. The

CVs of all microgel samples with different HQ content are shown in Fig. 4.


80

Fig. 4. Cyclic voltammograms of the microgels in 0.05 mol/L phosphate buffer (pH =

7.3) at a scan rate of 1 V s-1 at 23°C. The reference electrode is Ag/AgCl and the

working electrode is glassy carbon electrode.

Fig. 5. Cyclic voltammogram of the CHHQ-4 microgels in 0.05 mol/L phosphate

buffer (pH = 7.3) at different scan rates. The reference electrode is Ag/AgCl and the

working electrode is platinum.


81

The electrochemical responses can be seen from the cyclic

voltammograms. A significant increase in the two peaks indicates a linear

relationship for various microgel samples with an increased amount of

hydroquinone. These redox-active microgels undergo two-electron

oxidation and reduction. The two oxidation peaks and reduction peaks can

be detected from the cyclic voltammograms. The redox process was shown

as follows (Scheme 3). Firstly, hydroquinone was deprotonated during the

oxidation process. Secondly, hydroquinone lost one electron, thus forming

quinhydrone radicals. The quinhydrone radicals are active. Then,

benzoquinone is formed by losing another electron from quinhydrone32.

As described above, hydroquinone, a reductant, can lose two electrons to

form benzoquinone. The redox couple is a charge-transfer complex and

the redox process is a reversible conversion including electrochemical and

chemical reversible process. However, the oxidation of hydroquinone is

irreversible because benzoquinone decomposes between pH 9-1133.

As seen from the cyclic voltammograms as shown in Fig. 5, at high

potentials, the oxygen evolution can give rise to the acidification of the

electrolyte owing to water decomposition, which is not entirely

compensated by the capacity of the buffer. In addition, compared to the

CV obtained for glassy carbon electrodes, the CV is enriched in features,

which may be due to the joint effects of adsorption phenomena, water

decomposition and electrochemical microgel switching.

2.3.5.2 Bulk Electrolysis

It was observed from the CV measurements that the microgels are

redox-active and electrochemically active. The fact that a small number of

redox-active materials are capable of oxidizing or reducing at the electrode

surface, showing that only a small amount of conversion was observed


82

during the oxidation or reduction process34. In order to achieve a high

conversion, controlled potential bulk electrolysis measurement was

considered to obtain a high electrochemical response whilst the microgel

dispersion could be fully oxidized or reduced. Meanwhile, the

hydrodynamic radius of the microgels was measured following

electrolysis.

Fig. 6. (A) Hydrodynamic radius of CHHQ-4 microgel with altered electrochemical

potentials in three redox circles. Photography of the CHHQ-4 microgel solution after

electrolysis at (B) oxidized and (C) reduced states. (D) Schematic illustration of redox

process of hydroquinone and benzoquinone20.

Typically, bulk electrolysis was conducted in a 0.05 M PBS buffer (pH

=7.3) for three redox cycles. During the oxidation process, an

electrochemical potential was applied and kept constant at 2 V vs.

Ag/AgCl until the faradaic current was close to zero after 1.5 hours.

Contrastingly, it will be kept at -2 V vs. Ag/AgCl during the reduction

process. The charge indicated that the current was approximately 5% of


83

the original current after 5.5 hours. The color of the microgel solution

changed from white to light brown after transitioning to the oxidation state,

as shown in Fig. 6B and Fig. 6C. Moreover, it would change from light

brown to dark brown after changing from the oxidation state to the

reduction state. The sizes of the microgels were measured after these two

redox states for three circles as shown in Fig. 6A. The results showed that

the microgels swelled after the oxidation process and shrank after the

reduction state. The redox sensitivity was reversible.

Scheme 3. The oxidation/reduction process of hydroquinone/benzoquinone20, 56,

58-59.

The mechanism of the swelling-shrinking transition of microgels during

redox circles was as follows. Firstly, benzoquinone units prevail in the

microgels after oxidation and hydroquinone units, which can form stronger

hydrogen bonds than benzoquinone in microgels, prevail after reduction,

as shown in Fig. 6D. The microgels will swell owing to the looser microgel

structure during the oxidation and shrink during the reduction. Secondly,

the protons involved in the electrochemical reaction were different in the

oxidation and reduction states, respectively. During the oxidation process,

the electrolysis-induced pH changes occurred due to the proton enrichment

as shown in Scheme 3. The pH value of the microgel solution decreased

to 2.8 after oxidation and increased to 8.0 after reduction compared to the


84

original pH value (pH = 7.3), inducing the swelling-shrinking transition of

the microgels. The hydrodynamic radius results indicate that a swelling

ratio Rh(pH 3)/Rh(pH 8) = 1.6 is high when compared with pH variation

from pH 3.0 to pH 8.0 (Fig. 2A).

In conclusion, the size of microgels can be adjusted via redox circles by

means of monitoring environmental potentials. During the oxidation and

reduction process, the microgels swelled or shrank, suggesting that these

processes are reversible.


2.3.6.1 Degradation of Microgels by Urea

Gurney, Frank and Wen reported that urea molecules may rearrange the

neighboring water molecules when they spread homogeneously in microgel

solutions, thus preventing the molecules from participating in hydrogen-bonding

water clusters35. In order to investigate the urea-mediated degradation, urea was

applied as a hydrogen bond breaker in an aqueous solution to disrupt the physical

cross-linker in microgels, and thus, degrade the microgels. The degradability of

microgels was monitored by taking DLS and TEM measurements to assess the

changing size and morphology of microgels in the presence of urea.

A series of microgels were synthesized and applied to evaluate the effect of

urea-mediated degradation on microgels. As shown in Scheme 4 and Table 3,

three types of microgels with physical cross-linking-only (CHHQ), physical and

chemical cross-linking (CHHQGA) and chemical cross-linking-only (CHGA)

were prepared and applied for degradation by urea to verify that the physical

cross-linking within microgels can be disrupted. In the chemical-cross-linked

microgels, glutaraldehyde was used as a chemical cross-linker.


85

Table 3. The synthesis of particles with chitosan, hydroquinone and glutaraldehyde

amounts used for supramolecular assembly within the microgels.

Name of

sample

Mass ratio of

chitosan to

hydroquinone

Chitosan

(g)

Hydroquinone

(g)

Glutaraldehyde

(g)

Type 1 CHHQ-4 1:2 0.012 0.024 -

Type 2 CHHQGA-4 1:2 0.012 0.024 0.012

Type 3 CHGA - 0.012 - 0.012

As shown in Fig. 7, CHHQ-4 microgels, cross-linked by hydrogen bonds,

started to degrade when exposed to urea. During the first 10 minutes, the

microgels rapidly changed in size and subsequently swelled, as shown in Fig.

7A. The physical cross-linkers in microgels were disrupted which induced the

swelling. After 5 minutes, the size of microgels decreased due to the larger

number of disrupted hydrogen bonds. The microgels then decomposed slowly

over time and an equilibrium of a radius at approximately 100 nm was obtained,

indicating that the degradable products were small fragments. The physical

cross-linkers were broken and the remaining materials could be assigned to

poly(hydroquinone), which is not soluble in water36.

During degradation, the color of microgel dispersion changed from the

original milky color to transparent (Fig. 7B,C). Moreover, the degradation

behaviour was monitored using TEM as shown in Fig. 8. As shown in Fig. 8A,

the microgels started to swell following the addition of urea. Afterward, they

degraded into small fragments due to the disruption of their hydrogen bonds.


86

Scheme 4. (A) Schematic representation of preparation of chitosan-hydroquinone

(CHHQ), (B) chitosan-hydroquinone-glutaraldehyde (CHHQGA), and (C) chitosan-

glutaraldehyde (CHGA) microgels.


87

Fig. 7. (A) Determination of microgel degradation in the presence of urea via DLS.

Photograph of the CHHQ-4 microgel solution (B) before and (C) after degradation in

the presence of urea.

Microgels fabricated by physical cross-linkers were investigated as previously

discussed. In order to obtain more information about urea-mediated degradation,

two types of microgels, CHGA and CHHQGA-4, were employed as references.

The CHGA microgels, fabricated by chemical cross-links, remained unchanged

in the presence of urea over a period of 6 days as shown in Fig. 7A. This

indicated that urea has no effect on the CHGA microgels, which were cross-

linked only by glutaraldehyde. In addition, CHHQGA-4 microgels with physical

and chemical cross-links were studied by means of DLS (Fig. 7A) and TEM (Fig.

9). During the polymerization of CHHQGA-4 microgels, physical cross-links

were formed between chitosan and hydroquinone. Meanwhile, the chemical

cross-links were formed within the microgels with the help of glutaraldehyde

which reacted with the amino groups of chitosan. As shown in Fig. 7A, the DLS

results showed that the size of microgels remained unchanged because urea


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cannot disrupt the chemical cross-links. TEM images indicated that the

morphology of microgels maintains a spherical shape throughout the whole

degradation time, as shown in Fig. 9A. During microgel synthesis, both the

physical cross-linker and chemical cross-linker formed. After adding the urea,

the hydrogen bonds decomposed and the chemical cross-linker formed in

microgels. Therefore, the microgel network became loose and the particles

swelled slightly.

Fig. 8. (A) TEM images and (B) particle size distribution from DLS analysis of CHHQ-

4 microgels in the presence of urea at 0 h (r = 200.14 nm, PDI = 0.299), 1h (r = 487.83

nm, PDI = 0.223), 2h (r = 289.52 nm, PDI = 0.340), 4h (r = 163.28 nm, PDI = 0.368),

1 day (r = 96.91 nm, PDI = 0.362) and 7 days (r = 94.31 nm, PDI = 0.342).


89

Fig. 9. (A) TEM images and (B) particle size distribution from DLS analysis of

chemically cross-linked CHHQGA-4 microgels in the presence of urea taken at 0 h (r

= 268.51 nm, PDI = 0.159), 1h (r = 277.42 nm, PDI = 0.336), 1 day (r = 295.61 nm,

PDI = 0.168) and 7 days (r = 291.30 nm, PDI = 0.367).

2.3.6.2 Degradation of Microgels by an Enzyme

We next investigated how an enzyme degraded the CHHQ microgels.

Lysozyme, a model enzyme used to degrade microgels, can cleave the glucosidic

linkage of chitosan within microgels, as shown in Fig. 10A. Lysozyme can

hydrolyze microgels from the combination of N-acetylglucosamine residues in

chitosan and the active sites in lysozyme, referred to as the hexameric binding

sites37. The microgels were degraded via chitosan breakage. The DLS results

indicated that the size of microgels rapidly decreased during the first 20 minutes

and then decreased slowly, as shown in Fig. 10B. This means that the

degradation rate was high during the first stage, and becomes lower. Due to the

high enzyme concentration (10 mg/mL), the microgels’ network collapsed in the

presence of lysozyme. At this stage, the enzyme rapidly eroded the microgels

from the surface. The degradation rate was high due to the high content of

degradable units within the microgels. The degradation rate then slowed owning


90

to the decreased number of cleavage units. The microgel dispersion without

adding urea was applied as a control. The variation in the morphology of the

microgels was monitored using TEM as shown in Fig. 11A. The images showed

that the microgels collapsed within 1 hour and further degraded into nano-sized

fragments after 1 day.

Fig. 10. (A) Enzymatic degradation of CHHQ-4 microgels over time determined by

DLS in the presence of lysozyme and without lysozyme. (B) Weight loss of CHHQ-4

microgels in pH 6 buffer at 37°C over time. (C) Schematic illustration of enzymatic

degradation of chitosan by lysozyme38.

To further investigate the extent of the degradation behaviors, the weight

change of microgels during degradation was measured, as shown in Fig. 10C.

When exposed to lysozyme, up to approximately 70% of the weight loss of the

microgels occurred during the first 20 minutes. This then slowed during the time

that followed until reaching equilibrium. The weight loss was up to

approximately 76% after 1 day. Therefore, the microgels partially degraded and


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the weight loss tendency showed the same trend as the DLS and TEM results

reported above.

Fig. 11. (A) TEM images and (B) particle size distribution from DLS analysis of

CHHQ-4 microgels taken from the lysozyme dispersion at 0 h (r = 281.23 nm, PDI =

0.126), 1 h (r = 110.31 nm, PDI = 0.256) and 1 day (r = 76.85 nm, PDI = 0.285). (C)

Degradation of CHHQ-4 microgels over time determined by DLS in the presence of

lysozyme and bovine serum albumin (BSA).

In order to explore the degradation profiles induced by an enzyme at different

concentrations or a protein, the degradations of microgels triggered by an

enzyme or protein were carried out as shown in Fig. 11C. At a lower enzyme

concentration, the degradation rate is noticeably lower; with an increase in

enzyme concentration, the degradation rate is much faster. Therefore, the

degradation rate depended on the enzyme concentration. To compare the

degradation behaviors of enzymes and proteins, BSA-triggered degradation of


92

microgels was investigated. In the presence of a high concentration of BSA, the

size of the microgels remained constant, indicating that the microgels cannot be

degraded by a protein.

Scheme 5. Schematic illustrations of two different degradation approaches of

microgels via (A) urea and (B) enzyme.

As shown in Scheme 5, the proposed two degradation mechanisms were

illustrated and discussed as follows. In the presence of urea, the microgels firstly

swelled due to the partially disrupted hydrogen bonds and then degraded during

the following time owing to the further disruption. The microgels degraded into

small fragments after 1 day, indicating that microgels were physically cross-

linked and the hydrogen bond within microgels could be degraded by urea. On

the contrary, enzyme-induced degradation demonstrated a different mechanism

of action. Lysozyme can degrade the microgels by cleaving chitosan’s

glycosidic linkages. Moreover, the enzyme-induced degradation was faster than

urea-induced degradation. The enzyme degraded the microgels by eroding their

surface from outside to inside. Therefore, the size of particles decreased over

time.


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The drug loading and controlled releasing behaviors were investigated by

simply mixing a modal drug, DOX, into the microgel dispersion. These were

determined using a UV-Vis spectrometer at an absorbance of 496 nm. The drug

loading was obtained by incubating DOX in microgels for 16 hours and stirring

them overnight, in the approach to equilibrium. The loading efficiency was

quantified by Equ. (2) and the encapsulation efficiency was 80.9%. The

mechanism for DOX encapsulation was based on physical entrapment by means

of π - π stacking with quinone part between poly(hydroquinone) and DOX, and

hydrogen bonding with hydroxyl groups and amino groups between chitosan

and DOX.

To investigate the DOX release behavior in the presence of lysozyme, the

drug release profiles were conducted by measuring the content of released DOX

from DOX-loaded microgels whilst exposed to lysozyme as shown in Fig. 12.

During the first stage, approximately 43% of DOX at pH 6.0 and 10% of DOX

at pH 7.4 were released with a high release rate during the first 1 hour. The

release rate then decreased after the initial burst release. After 10 hours, the

release amount of DOX reached equilibrium and about 70% of DOX was

released at pH 6.0 and 36% at pH 7.4. In the presence of lysozyme, the release

rate was faster at pH 6.0 than that at pH 7.4. The enzymatic activity of lysozyme

reached its maximum at pH 6.0 and decreased at pH 7.4, thus, degrading DOX-

loaded microgels faster at pH 6.0 than that at pH 7.4. Moreover, the swelling

properties induced different release rates. Due to the fact that the microgels

would swell at pH 6.0, the particles were larger at pH 6.0 than that at pH 7.4.

Therefore, the diffusion of the DOX from the microgels was quite easier at pH

6.0. These factors induced different drug release properties. It was observed that

the microgels could be used as the drug carriers for the controlled release of the

drug.


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Fig. 12. Cumulative drug release profiles of CHHQ-4 microgels loaded with

doxorubicin hydrochloride in PBS buffers (pH 6.0 and pH 7.4) at 37 °C with or without

the addition of lysozyme.

2.3.8 Cytotoxicity Evaluation

To obtain more information on the biocompatibility of microgels, an

evaluation of cytotoxicity study was conducted by means of XTT cell

proliferation assay, that is, the L-929 murine cells, used as target cells, were

exposed to a series of CHHQ microgels with increased concentrations as shown

in Fig. 13. The results showed a dramatic decrease in the cell viability while the

cells were exposed to the microgels dispersion with a high concentration.

Therefore, the cytotoxicity of the microgels is dose-dependent. These results

suggested that the non-toxic doses of microgels ranged from 0.001 mg/mL to

0.010 mg/mL. For these concentration levels, the microgels showed no toxic

effects, indicating that the microgels had high biocompatibility. Meanwhile,


95

when the microgels concentration increased above 0.100 mg/mL, they exhibited

toxicity. Moreover, the results showed that at a high concentration of 1 mg/mL,

the cell survival was approximately low to zero. The three types of microgels at

the same concentration levels had no significant effect on the cell viability. It

was concluded that the cytotoxicity of the microgels was dose-dependent and

the non-toxic dose was from 0.001 mg/mL to 0.010 mg/mL.

Fig. 13. Cell viability of L-929 cells after exposure to CHHQ-1, CHHQ-3 and CHHQ-

5 microgel dispersions assessed by XTT cell proliferation assay.

2.4 Conclusions

In summary, we successfully synthesized a series of novel

redox/pH/electrochemical potential stimuli-responsive microgels via the

oxidative polymerization of hydroquinone in the presence of chitosan in an

inverse miniemulsion system. The obtained microgels can be prepared by

physical cross-links, that is, hydrogen bonding, and can also be degraded when

exposed to either urea or an enzyme environment. These obtained pH-responsive

microgels showed swelling behaviors in acidic mediums and shrunk in alkaline


96

surroundings due to the amino groups within the microgels that can carry the

positive charge in an acidic environment. These properties render the microgels

colloidally stable. Moreover, the microgels are redox-active and exhibit swelling

or shrinking behavior in response to the equilibrium potential changes that were

adjusted by electrochemical means investigated by DLS measurement. The

electrochemical behavior indicated that the microgels showed a two-electron or

proton redox behavior, including the oxidation of hydroquinone and reduction

of benzoquinone which occurred at the two potentials, respectively. The

microgels are biodegradable. In the presence of urea or lysozyme, the microgels

can be degraded via the cleavage of the hydrogen bonds or the glucosidic

linkages. They can encapsulate an anticancer drug, DOX, which is also released

in the presence of lysozyme. The DOX release can be triggered by exposing the

microgel to the lysozyme environment. In conclusion, due to their excellent

biocompatibility, biodegradability and redox-activity, the microgels can act as

the smart drug delivery devices for drug delivery systems, energy converters or

biosensors for energy conversion and storage technology.


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3. Polyaniline-Chitosan Microgels

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102

3.1 Introduction

In recently years, conducting polymers have attracted growing attention and

become promising materials because they display distinctive electronic

properties due to their unique conjugated π-electron system1. The conductivity

of conductive polymers can occur upon doping, which is the process in which

the polymers are oxidized or reduced. The mechanism of conductivity of

polymers is doping. During doping, conjugated polymers produce high

conductivity and the polymers are then oxidized or reduced. The oxidization is

generated by the acceptance or removal of electrons, which induces a radical

hole on the chain. Conductive polymers can be used in various fields, like

electronic devices, such as transistors2, biosensors3, chromatography4, energy-

storage cells5, alternative energy sources6, catalysts7 and indicators8.

Furthermore, as conducting polymers can transport small electronic signals in

human body, they can be applied to biotechnology applications such as artificial

nerves9. More interesting conducting polymers, such as polymer-coated

electrodes, have been prepared by scientists that can be used as a

neurotransmitter applied as a drug release system in the brain10. Conducting

polymers are also promising candidates for tissue engineering. Min Zhao et al.

discovered that at a genetic level, electrical stimulation or electric cues can play

an important role in wound healing, and can also identify genes, which is a

necessary aspect of electrical-signal-induced wound healing11. Aref Shahini et

al. prepared a conductive platform by including the incorporation of poly(3,4-

ethylenedioxythiophene):poly(4-styrene sulfonate) (PEDOT:PSS) in the

composition of BaG/Gel scaffolds. It can attain effective electrical or magnetic

stimuli during the bone healing process using tissue engineering techniques12.

Electric stimuli play a vital function in wound healing, nerve regeneration and


103

recovery from spinal cord damage13. There were also several kinds of

morphologies of polyaniline products, such as nanotube composites14,

nanofibers, electroactive films15, hydrogels16, nanoparticles17, capsules18, and

nanowires19.

Scheme 1. Different forms of polyaniline at varying degrees of oxidation20.

Among these diverse conducting polymers, polyaniline is an intrinsically

conductive polymer that exhibits reversible redox properties such as doping or

dedoping, and can be synthesized by oxidative polymerization of aniline, which

is seen as a type of polycondensation. The growth of polymer chains drives from

continuous addition of monomers to the end of the chain and the redox process

takes place in aniline that serves as a reducer, and the growing chain, which

serves as an oxidant21. The conductivity of polyaniline is generated due to the

strong acids that can stimulate the protonated states of polyanilines and at the

same time, stabilize the charge of the polyaniline. The polyconjugated structure

of polyaniline forms a zigzag chain lying in one plane, and an overlap of the π-

electron cloud is above and below the plane of the chain. The nitrogen lone


104

electron pair performs the same function as the π-electron. This assures

polyconjugation and the polyconjugated system, which increases the charge

carrier mobility. As the environmental pH changes, polyaniline can be rendered

conductive due to the protonation. Polyaniline switches between protonated and

unprotonated states and undergoes two particular redox processes as the

environmental pH changes between acid and base. There are two different

situations in which polyaniline can change its form from a conducting state to a

non-conducting state. The first way is to introduce electrons into polyaniline and

reduce nitrogen atoms. The second way is to remove the polaron-stabilizing acid,

which caused the polyconjugation of the polyaniline to disappear. The

conductivity properties of polyaniline depended on its degree of protonation and

the state of oxidation. Moreover, the protonation of polyaniline is reversible.

During the protonation process, a polyaniline chain is bound to acid molecules.

The reversible process is performed by adding a base. Scheme 1 showes four

different structures of polyaniline in various redox states, with the oxidation

centers forming from nitrogen atoms. Polyaniline has fully reduced and

oxidative forms named leucoemeraldine (LM) and pernigraniline (PNA),

respectively. As shown in Scheme 1, the polyaniline polymers present four

different forms at varying degrees of oxidation: fully oxidized pernigraniline

base (PNA) (all nitrogen atoms are imine), 75% oxidized nigraniline (NA), 50%

oxidized emeraldine (EM) (the ratio of imine to amine is 0.5) and fully reduced

leucoemeraldine (LM) (all nitrogen atoms are amine) that contain different

proportions of quinonoid imine and benzenoid amine20. The protonation degree

of the polymer depends on its environmental pH. Polyaniline can be fully

protonated by an aqueous hydrochloric acid solution. The partially protonated

form of polyaniline can be prepared by the oxidative chemical or

electrochemical polymerization of aniline. On the contrary, the deprotonated

form is the same as a semiconductor and can be induced by an aqueous


105

ammonium hydroxide solution. The most stable state among them is emeraldine,

in which the oxidized and reduced units are equal due to every second nitrogen

atom being oxidized.

Furthermore, the electrochemical property of polyaniline is electrochromic

behavior22. The color change occurs during the oxidation-reduction cycle. It

turns transparent yellow at -0.2 V, green at 0.5 V, dark blue at 0.8 V, and black

at 1.0 V20. Both the isolating form (emeraldine base) and conducting form

(emeraldine salt) are stable in air23. The oxidation (coloration) corresponds to

the proton-elimination and the reduction (decoloration) is accompanied by

proton-addition, and both of these processes present quick and reversible

responses20.

However, polyaniline has some drawbacks due to its poor processability, low

solubility and infusible character with other systems24. In order to solve these

problems, the chemical modification of aniline has been studied to improve its

properties, such as by doping with acids or forming polyaniline

nanocomposites25. In the latter, incorporating polyaniline into natural polymeric

materials can combine the conductivity of polyaniline and the processability of

the natural matrix. A variety of natural and synthetic biodegradable polymers

have been applied to tissue engineering applications and drug delivery systems,

such as cellulose26, poly(lactic acid)27, chitosan28, gelatin29 and other

biomaterials30. Among these biodegradable polymers, chitosan is a natural

biopolymer that is a partially N-deacetylated derivative of chitin, which is

produced from waste crab and krill shells obtained by the fishing industry.

Because of its specific biodegradability and biocompatibility, it has been widely

used in lots of fields such as in controlled drug delivery systems31, treatment of

waste water32, tissue engineering33, gene therapy34, wound healing accelerators35

and biosensors36. More importantly, chitosan has excellent biocompatibility and

biodegradability for medical modification due to the amino and hydroxyl groups


106

along the main chains which offer the possibility of diverse chemical

modifications by introducing cross-linking agents, as well as easy

functionalization37. These characteristics make it a suitable material for grafting

with polyaniline. Therefore, chitosan has been selected as a matrix and the

chitosan-graft-polyaniline copolymer is a prime candidate for our study.

This chitosan/polyaniline composite is prepared through a grafting reaction,

in which aniline is oxidized by using chitosan as a steric stabilizer. Oxidative

polymerization proceeds in two different ways. One can be considered as

polycondensation, wherein the polymer grows and the cation radical oxidation

sites recombine. The other one is a form of electrophilic substitution, in which

the oxidized nitrogen-containing structure substitutes one proton of the ring of

another aniline molecule by attacking the phenyl ring. After that, the ring and

the nitrogen-containing structure lose one proton, the monomer units bound

together, and the chain becomes longer. During polymerization, aniline can be

polymerized through chemical or electrochemical polymerization to form

polyaniline which grafts onto the chitosan and spreads out into the chitosan

network. The obtained composite then exhibits the processability of the chitosan,

which was used as a matrix, and the electrical conductivity of the conductive

polymer. During the oxidation, nitrogen atoms of polyaniline act as oxidation

centers and the charge carriers are produced in the polymer. In addition, the

number of oxidized nitrogen atoms in polyaniline may alter from 0

(leucoemeraldine, reduced states) to nearly 1 (pernigraniline, oxidized states).

The reduced and oxidized forms of polyaniline will change into the oxidized

state without the external potential. The best charge stabilizer for polyaniline is

a strong acid, which renders the polymer conductive. The oxidative

polymerization of aniline and chitosan depends on the pH of the reaction media,

which can occur in the low pH range. In an acidic medium, the polymerization

of aniline yields dark green powder with high conductivity. Contrastingly, where


107

polymerization proceeds in basic media the synthesis products will yield low

conductivity.

Previous studies have reported that chitosan-graft-polyaniline composites

could be prepared and applied in several fields, such as the hydrogels38,

nanofibers39, films40 or as the candidate with metal composites41. In this study,

a novel pH-sensitive and redox-active conducting microgel is prepared using

inverse miniemulsion polymerization. The aim of the work is to synthesize the

conductive and biodegradable microgels which incorporated the different

amounts of conducting polymers into the microgel matrix to be used in a drug

delivery system. The swelling ratio of the microgels can be controlled by

adjusting the amount of grafted aniline. During the preparation of the microgels,

polyaniline is grafted onto chitosan and the copolymers are cross-linked by

glutaraldehyde. The microgels are the combination of conductive polymers and

the swelling/de-swelling biopolymer products which present the dual

characteristics of biocompatibility and conductivity. In addition, the chitosan-

based microgels contain amino and imino groups that can be protonated in acidic

mediums, causing the microgels to swell. Due to its conductivity,

biocompatibility and biodegradability, it can be considered as an attractive

biomaterial for diverse applications such as drug delivery systems42, diverse

biomedical applications43, biosensors44, and also a scaffold well-suited to tissue

engineering45.


3.2.1 Materials

All reagents were purchased commercially and used without further

purification except chitosan. Chitosan of medium molecular weight (75-85%


108

deacetylated, Sigma-Aldrich) was used with further purification based on the

steps from the literature46. Aniline (≥99.5%), ammonium persulfate (APS,

≥98%), methylpyrrolidone (NMP, 99%), glutaraldehyde (25% in H2O), acetic

acid (99%), ammonium hydroxide solution (NH3·H2O, ≥25% in H2O),

cyclohexane (≥99.8%), Span 80, dibasic potassium phosphate (K2HPO4, ≥98%),

monobasic potassium phosphate (KH2PO4, ≥98%), methanol (≥99.8%), acetone

(≥99.5%), ethanol absolute (≥99.8%), lysozyme from chicken egg white (protein

≥ 90%, ≥40,000 units/mg protein), were bought from Sigma-Aldrich and used

as received. Deionized (DI) water was used in all experiments. Dialysis

membranes (MWCO = 1.2 kDa) were purchased from Carl Roth. DI water was

obtained as a reaction medium.

3.2.2 Synthesis of Chitosan-Grafted-Polyaniline (CH-g-Ani)

Copolymers

CH-g-Ani copolymers were prepared with different grafting ratios. Firstly,

different CH-g-Ani samples were prepared as follows. Purified chitosan was

dissolved in 0.1 M acetic acid and then stirred overnight at room temperature

until it completely dissolved. 10 mL of chitosan solution (0.01 g/mL in acetic

acid) was added to a 50 mL round bottom flask and then the different amount of

aniline that dissolved in 10 mL 1 M HCl was added dropwise. Next, different

amounts of ammonium persulfate were dissolved in 2.5 mL 1 M HCl that was

added dropwise to the previous mixture. The reaction mixture was cooled in an

ice-bath for 1 hour whilst avoiding light. Then the ice bath was removed and the

stirring continued at room temperature for 5 hours. The grafting reaction lasts

for 6 hours altogether. The same amount of 1 M NaOH was added to neutralize

the reaction mixture and the copolymer was precipitated by adding 200 mL of

ethanol absolute. The product was filtered and then washed with NMP several


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times to remove unreacted aniline. The product was finally washed one time

with deionized water to remove NMP. It was dried in the oven at 60 °C for 2

days and kept at room temperature for further use47. As shown in Table 1,

different grafting degrees of anilines to chitosans in copolymers were prepared

by using the different amounts of amino molar ratios of chitosan to aniline in the

reaction process.

Table 1. Ratios of chitosan and aniline synthesized in chitosan-grafted-polyaniline

copolymers.

Name of the

copolymers

Chitosan

content (g)

Aniline

content (g)

Molar ratio of -NH2

in chitosan and in

aniline

(NH4)2S2O8

content (g)

CH-g-Ani-1 0.1 0.023 1:0.5 0.028

CH-g-Ani-2 0.1 0.046 1:1 0.057

CH-g-Ani-3 0.1 0.092 1:2 0.113

CH-g-Ani-4 0.1 0.139 1:3 0.170

CH-g-Ani-5 0.1 0.231 1:5 0.283

3.2.3 Synthesis of Microgels (W/O miniemulsion)

The dry copolymer chitosan-grafted-polyaniline was dissolved in 1 M HCl,

stirred for 3 days to totally dissolve, resulting in the doped polyaniline in the

chitosan backbones. The microgels were prepared by inverse miniemulsion

polymerization using 1 M HCl as an aqueous phase and cyclohexane as an

organic phase. In the aqueous phase, these doped copolymer solutions with the

different suitable amounts of CH-g-Ani were capable of functioning as the

matrix whilst glutaraldehyde was used as a cross-linker. In the organic phase,

Span 80 (0.258 g) was used as a surfactant that was dissolved in 10 mL of

cyclohexane. The mixture of the aqueous phase and organic phase were

ultrasonicated using a Branson Sonifier 450 at the duty cycle of 50% and output


110

control of 40% under the ice cooling for 10 minutes. After sonication, the

miniemulsion stirred at room temperature for 6 hours. Five different samples

(CH-g-Ani-1, CH-g-Ani-2, CH-g-Ani-3, CH-g-Ani-4, and CH-g-Ani-5) were

developed with the different grafting ratios, as shown in Table 1. These

microgels contained different amounts of polyaniline, but had the same chitosan

content. After synthesis, the microgel dispersion was purified by centrifuging 10

minutes at 4000 rpm several times. The supernatant was discarded, and the

precipitated was washed several times with 10 mL of cyclohexane. The final

precipitate was re-dispersed in 5 mL of DI water and underwent dialysis for

further purification.

3.2.4 Characterization

Fourier transmission infrared (FTIR) measurements were performed by

Nexus 470 (Thermo Nicolet) at room temperature. Microgel dispersions were

dried by lyophilization and pressed in a KBr pellet.

Transmission electron microscopy (TEM) measurements using a Zeiss

LIBRA 120. The electron beam accelerating voltage was set at 120 kV. The

samples were produced by drop coating the microgel solution on a formvar-

carbon-coated copper grid with 400 meshes. The surface of the grid was

pretreated with plasma for 120 s before sample preparation. After surface

treatment, one drop of the microgel solution was added to a grid that was placed

on a piece of filter paper and left to dry overnight at room temperature.

The hydrodynamic radius of the microgel particles in the aqueous medium

was measured using an ALV/LSE-5004 Light Scattering Electronics Multiple

Tau Digital Correlator with the scattering angle set at 90° and equipped with an

ALV-5000/EPP multiple digital time correlator and laser goniometry system

ALV/CGS-8F S/N 025 with a helium-neon laser (Uniphase 1145P, output power


111

of 22 mW and wavelength of 632.8 nm) as a light source. The electrophoretic

mobility of microgels was determined using a Malvern Zetasizer Nano ZS. The

measurements were carried out in the pH 3-11 range at 25 °C after equilibration

for 120 s. Before measuring, all samples were diluted with PBS buffers as a

function of pH in the range of 3 to 11.

Cyclic voltammetry (CV) measurements were performed by scanning the

potential in the respective potential window (-0.2 V~1 V) at a scan rate from

0.01 V s-1 ~ 1 V s-1 at room temperature. A conventional three-electrode cell was

used with a glassy carbon (GC) electrode as a working electrode and an Ag/AgCl

electrode stored in 1 M KCl served as a reference electrode. All potentials in the

text and figures refer to an Ag/AgCl couple. A platinum wire electrode served

as a counter electrode. Before performing each measurement run, the working

electrode was polished with 1 µm diamond and subsequently polished with 0.05

µm alumina, rinsed with water and then dried with a stream of argon. CV

measurements were performed in 0.1 M HCl and the microgel solution was

purged with argon for 10 minutes to remove any dissolved oxygen.

3.2.5 Enzymatic Degradation of Microgels

To investigate the microgel degradation process, the size and morphology of

microgels were estimated by taking DLS and TEM measurements. Due to the

glycosidic linkage in chitosan, which can be hydrolyzed by lysozyme, DLS and

TEM measurements of the microgels in the presence of lysozyme were carried

out to investigate the changing radii and altered morphology of microgels as a

function of time.

The degradation behaviors of the two different microgel samples with

different polyaniline amounts were investigated by testing the variation

tendency of the particle size by means of DLS. Firstly, the original microgel


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dispersion was diluted in deionized water and then the proper concentration for

DLS was determined. After that, the determined concentration of microgel

dispersion was injected into the lysozyme solution (10 mg/mL) under continuous

stirring at 37 °C and the DLS measurements were carried out. Meanwhile, the

DLS measurement proceeded at regular time intervals, which were shorter time

intervals during the first 100 minutes and longer intervals over the later period

with a maximum interval of 1 day until the microgels degraded completely.

The morphology of particle degradation was tested by TEM measurement.

The sample without adding lysozyme was prepared by drops of original sample

solution to a TEM grid which was then dried at room temperature for testing.

The other samples for degradation were prepared by mixing the microgel and

lysozyme solution to form the mixture, which was stirred throughout the whole

degradation process. The samples for degradation measurement were prepared

by adding several drops of mixed dispersion to the TEM grid after 1 hour and 1

day, which were then placed at room temperature for drying.


3.3.1 Synthesis of Microgels

The microgels were prepared in two steps. Firstly, CH-g-Ani copolymers

were prepared through the oxidative polymerization of aniline in the presence of

chitosan at different grafting ratios. Secondly, the microgels were prepared by

means of crosslinking CH-g-Ani copolymers, using glutaraldehyde as a cross-

linker.

The mechanism of oxidative polymerization of aniline was investigated and

the important polymerization step is the process through which the monomer can

form dimers that control the rate of polymer growth in the polymerization


113

process. Ammonium persulfate was used as a strong oxidant, which can generate

primary radicals that are sulfate ion radicals (SO4-.). In the presence of a strong

oxidant, the aniline monomers can be oxidized to radical cations to form the

dimers. By means of electrophilic aromatic substitution, the dimers can be

oxidized, deprotonated, and react with aniline. During further oxidation and

deprotonation steps, the tetramers form via the reaction between oxidized

trimers and aniline. During the whole polymerization process, this step occurred

repetitively to develop polyaniline, as shown in Scheme 2. In addition, the

mechanism of grafted copolymerization between chitosan and polyaniline was

discovered and discussed. In the presence of the strong acid and oxidant, the

oxidative polymerization of aniline was initiated via a cationic radical as an

intermediate to form polyaniline. Meanwhile, polyaniline radical cation

introduced the chitosan macro radicals by the abstraction of hydrogen from the

-OH and -NH2 groups of chitosan at the same time48. Next, the copolymerization

reaction took place between the polyaniline radical cations and the chitosan

radical cations and the reactant, CH-g-Ani, was generated. Scheme 3 showed the

graft copolymerization of CH-g-Ani. In the acidic environment, the oxidative

polymerization of aniline was triggered by ammonium persulfate, and

poly(aniline) radicals were formed (Scheme 3A). Chitosan macro radicals

(Scheme 3B) and poly(aniline) cation radicals (Scheme 3A) are combined to

form CH-g-Ani (Scheme 3C)47.


114

Scheme 2. Mechanism of polyaniline polymerization49.


115

Scheme 3. Electrochemical copolymerization synthesis of (A) polyaniline radical

cation, (B) chitosan macro radical, (C) chitosan-grafted-polyaniline copolymer50 and

(D) CH-PANI microgel48a.

Table 2. Compositions of chitosan (CH) and CH-PANI microgels.

Sample name Chitosan

(mg)

Aniline

(mg)

Glutaraldehyde

(mg)

(NH4)2S2O8

(mg)

CH-PANI-1 10 2.31 5 5.66

CH-PANI-2 10 4.62 5 1.13

CH-PANI-3 10 9.25 5 2.26

CH-PANI-4 10 13.87 5 3.39

CH-PANI-5 10 23.11 5 0.56

CH 10 - 5 -

Following the successful preparation of CH-g-Ani, the microgels were

synthesized by inverse miniemulsion. In aqueous droplet, CH-g-Ani was


116

dissolved in and glutaraldehyde was used as a cross-linker to cross-link

copolymers. The aldehyde groups in glutaraldehyde are attached to amino

groups in CH-g-Ani copolymers. Thus, they were cross-linked in the aqueous

phase and formed the microgels’ network (Scheme 3D). The amounts of each

component are shown in Table 2.

3.3.2 FTIR Spectra of Microgels

Fig. 1 shows the FTIR spectra of CH-g-Ani copolymers. As shown in Fig. 1A,

the characteristic peaks of chitosan are located at 3500-3300 cm-1, attributed to

the stretching peaks of the -NH2 group51. The peaks at 2940 cm-1 and 2873 cm-1

are due to aliphatic C-H stretching mode52. The characteristic peaks of

polyaniline appear at 1507, 1483 and 1110 cm-1. These peaks are related to C=C

stretching of quinoid rings, C=C stretching vibration of benzenoid rings and the

absorption band of the N=Q=N bending vibration, respectively (Q=quinonoid)53.

The new absorption band at 747 cm-1 is from the -NH group, indicating that the

polyaniline has been grafted onto chitosan54.

The FTIR spectra of CH-PANI microgels are shown in Fig. 2. The peak at

3380 cm-1 could be assigned to -NH2 vibration stretching mode of chitosan, and

the peaks at 2923 cm-1 and 2855 cm-1 are related to aliphatic C-H stretching

vibrations. The peaks at 2923 cm-1 and 2855 cm-1 are due to the aliphatic C-H

stretching mode.


117

Fig. 1. FTIR spectra of chitosan, CH-g-Ani-3 and CH-g-Ani-5.

The C=C and C-C stretching and bending modes for the quinoid ring and

benzenoid ring in polyaniline show vibrations at 1590 cm-1 and 1500 cm-1,

respectively. The peak at 1170 cm-1 is the absorption band of the N=Q=N

bending vibration of protonated polyaniline55. The C-N stretching absorptions

are seen at 1380, 1315 and 1245 cm-1 which belonged to the stretching of C-N

in QB(trans-form)Q units, QB(cis-form)Q, QBB, BBQ units, and BBB units,

respectively (Q = quinonoid; B = benzenoid)20. The peak at 731 cm-1 is assigned

to the -NH bands. In addition, a new peak appears at 1654 cm-1 in the curves of

the CH-PANI microgels, attributed to the Schiff base group (-N=CH-), thus

indicating that a cross-linking reaction took place between the amino groups in

chitosan and aldehyde groups in glutaraldehyde56.


118

Fig. 2. FTIR spectra of chitosan and the CH-PANI microgels with different grafting

ratio.


Both chitosan and polyaniline contain nitrogen-containing groups (amino and

imino groups), which can be protonated to different levels when changing the

pH value of the surrounding environment. These microgels charges are

dependent on amino and imino groups in polyaniline and unreacted amino

groups in chitosan within microgels. The protonation of amino and imino groups

induced electrostatic repulsion in microgels, such that they were positively

charged in the acidic medium. While at the base, the amount of hydronium

[H3O+] reduced and the number of OH− anions increased. This may be due to

the formation of a Stern layer by means of the negatively charged counter ions

overcompensating for the positive surface charge57.


119

Fig. 3. (A) Hydrodynamic radius and (B) electrophoretic mobility of microgels in

buffers.

In order to compare the swelling behaviors of chitosan and polyaniline, six

samples, including chitosan microgels (CH), CH-PANI-1, CH-PANI-2, CH-

PANI-3, CH-PANI-4, and CH-PANI-5 microgels were investigated. Among

them, the chitosan microgels were prepared through inverse miniemulsion

polymerization, in which the chitosan was cross-linked by glutaraldehyde.

Therefore, CH were prepared without the copolymerization of polyaniline. As

shown in Fig. 3A, sample CH swelled most rapidly compared to the CH-PANI

microgels. Moreover, it had the highest amino groups amongst all the samples.

Without copolymerizing to polyaniline, more -NH2 groups were protonated into

-NH3+ groups. The microgels network expanded due to the electrostatic

repulsion of -NH3+ groups, as well as the increase in the hydrophilic interactions

of chitosan and decreased number of hydrogen bonds amongst the amino group

and hydroxyl group58. The microgels, therefore, swelled in the acidic medium

and shrank as the pH value increased. The effect of amino groups on the swelling

of microgels was obvious among all of the microgel samples.


120

For the CH-PANI samples, it was found that the swelling behaviors of these

samples were less obvious compared to the CH sample due to the low content

of the amino groups (-NH2) and high content of the imino groups (-NH-) and

disubstituted amino groups (-N=). Moreover, the swelling behaviors of CH-

PANI samples showed a different trend as compared to the CH sample.

For the polyanilines, there are two different kinds of nitrogen-containing

structure groups: imino (nitrogen atoms were oxidized) and disubstituted amino

groups (nitrogen atoms were not oxidized). The imino groups were protonated

at higher pH values, whilst the amino groups were protonated at lower pH

values59. As shown in Fig. 3B, when the pH level is lower than 4, both chitosan

and the polyaniline were protonated and the microgels accordingly carried

higher positive charges. Above pH 4, polyaniline becomes non-protonated and

its charge weakens60. The hydrodynamic radius in Fig. 3A also exhibited the

same trend. As a result of the protonated amino and imino groups, electrostatic

repulsion is generated in the microgels leading the particles to expand when the

environment is acidic.

3.3.4 Electrochemical Properties

The electrochemical behaviors of the CH-PANI-1 and CH-PANI-5 microgels

were investigated using a cyclic voltammogram. In 0.1 M HCl solution, the

oxidation and reduction peaks were investigated using a cyclic voltammogram

in the potential range from -0.2 V to 1.0 V with a scan rate of 1 V/s. The

electrochemical properties of microgels were adjusted by changing the ratio of

the initial amount of aniline during the polymerization process.


121

Fig. 4. Cyclic voltammograms of the CH-PANI-1 and CH-PANI-5 microgels in 0.1 M

HCl.

As shown in Fig. 4, the electrochemical properties of the two microgel

samples were investigated and the cyclic voltammograms showed a little

difference due to their different respective amounts of initial aniline. The CH-

PANI-1 microgel, which has a low content of polyaniline, showed less clearly

defined peaks. The spectrum CH-PANI-5 microgel showed distinct peaks, with

two pairs of oxidation peaks and two pairs of reduction peaks, respectively.

These two redox processes are due to the two reversible redox reactions of

polyaniline, which were the oxidation and reduction of polymers. These two

peaks correspond to the transitions among the three states of polyaniline. The

first redox pair (peak A and C) correspond to the first redox process that

indicated the transitions from fully reduced forms of polyaniline

(leucoemeraldine) to semiquinone forms of polyaniline (emeraldine). Further,

peak A corresponds to the first step of the oxidation process and peak C

corresponds to the further oxidation of polyaniline. The second redox pair (peak

B and D) correspond to the further redox process that shows the conversions


122

between the semiquinone forms of polyaniline (emeraldine) and the fully

oxidized forms of polyaniline (pernigraniline) which are shown in Scheme 149a,

49b, 61. The first pair shows a narrower peak than the second; this is because the

charge transformation in the first redox step is easier than the second one23.

Additionally, the critical point is that the electrochemical behavior of polyaniline

was depended on the pH of the environmental medium. The microgels dispersed

in a strong acid will show conductive properties, but not when they are dispersed

in neutral or alkaline mediums21b.


The microgel based on chitosan was biodegradable as lysozyme can

biodegrade chitosan. Therefore, the whole chitosan network is biodegradable.

The microgel dispersion was incubated with lysozyme (10 mg/mL) at 37 °C and

pH 6 under continuous stirring to investigate the enzymatic degradation behavior.

The degradation process was characterized by DLS and TEM.

As shown in Fig. 5, the hydrodynamic radius of the microgels underwent two

stages. During the first 20 minutes, the size of the microgels rapidly decreased.

During the following experimental time, the particle size slowly decreased until

it was close to 10 nm within one day. As shown in Scheme 4, the enzymatic

degradation of microgels in the presence of lysozyme is due to the cleavage of

the β-(1-4) glycosidic linkages in chitosan which can be degraded into chitosan

oligomers. The breakage was due to the combination of N-acetylglucosamine

residues in chitosan and the active sites in lysozyme, named the hexameric

binding sites. Therefore, the network of chemical cross-linking microgels will

collapse after an enzyme is added to it.


123

As shown in Fig. 5, two different microgel samples with different cross-

linking densities had nearly the same enzymatic degradation rates because the

chitosan content in these samples was the same and lysozyme can degrade the

polysaccharides in the chitosan chain. As shown in Fig. 6, the particles changed

their spherical shape during the 1 hour period and degraded into small fragments

after 1 day. The DLS and TEM results proved that these chitosan-based

microgels can be degraded into biocompatible byproducts through enzyme-

catalyzed hydrolysis.

Fig. 5. Degradation of the CH-PANI-1 and CH-PANI-5 microgels over time as

determined by DLS.


124

Fig. 6. TEM images of CH-PANI-3 microgels over time before and after adding the

enzyme in microgel dispersion.

Scheme 4. CH-PNI microgel degraded by lysozyme.

3.4 Conclusion

Polyaniline, one of the most interesting conductive polymers, has been

introduced and investigated in these biopolymer-based microgels which

exhibited pH-sensitive, biodegradable, and conductive properties. These

microgels were based on chitosan and blended with different amounts of

polyaniline. We characterized the chemical and electrical properties of

microgels using FTIR, DLS, CV and TEM. The results demonstrated that the

microgels exhibited pH-dependent phase transition behaviors in aqueous


125

solutions, whilst in acid environments, the microgels became negatively charged

due to the protonation of amino and imino groups. They presented

electrochemical behavior with two oxidation peaks and two reduction peaks

measured with cyclic voltammogram that corresponded to transitions among

three states of polyaniline (leucoemeraldine, emeraldine and pernigraniline). In

the presence of an enzyme, the microgels can be degraded at 37 °C and pH 6,

which was due to the cleavage of glucosidic linkage in chitosan within microgels.

As a result of their biocompatibility, pH-responsiveness, conductivity and

biodegradable characteristics, these microgels can be applied to several fields

such as functioning as scaffolds for tissue engineering, drug delivery vehicles

and biosensors.


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4. Dual-Degradable Dextran-Chitosan Microgels

131


This Chapter has been reproduced from Helin Li, Xin Li, Puja Jain, Huan Peng,

Khosrow Rahimi, Smriti Singh and Andrij Pich, Biomacromolecules, 2020, 21,

12, 4933-4944. Copyright 2020 American Chemical Society. Reproduced with

permission.


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4.1 Introduction

Natural biopolymers have recently attracted growing interest from researchers

investigating their use as bioactive materials for applications in tissue

engineering and sustainable drug delivery systems due to their renewable and

sustainable nature1. For sustained delivery drug purposes, injectable micro/nano

device systems can be fabricated, depending on the versatility of the natural

macromolecules. This is possible because of their biocompatibility,

biodegradability and capability of encapsulating bioactive agents into the

biocompatible carriers2. Therefore, these natural sources of bio-polymeric soft

materials have been widely used as both the building blocks to explore

responsive micro/nanogels as well as promising candidates for a variety of

biomedical applications3. In recent decades, a considerable amount of work has

been devoted to the development of biopolymer-based microgel preparations,

such that they have become a promising area of research for biomedical and

therapeutic applications. The utilization of biopolymers in the preparation of

microgels offers more efficient approaches to the encapsulation, stabilization,

culture and release processes of biologically active agents and molecules, such

as enzymes, cells, genes, peptides, proteins and drugs4. With these features, the

microgels, as powerful models for drug delivery vehicles, can be widely applied

in cancer therapeutics5.

To explore such applications, an increasing amount of studies have been

carried out over the last decade to look into biopolymer-based microgels. The

microgels’ properties and functions play an essential role in the self-assembly of

biological macromolecules, such as the attachment, spreading, and proliferation

of the cells for biomedical use6. Lee et al. successfully synthesized cross-linked

complex particles obtained from chitosan and poly(N-isopropylacrylamide)


133

(PNIPAM) through the surfactant-free dispersion copolymerization method and

soapless dispersion polymerization, which can be applied to drug delivery

systems. The copolymer particles exhibited a core-shell morphology, with 2, 2’-

azoisobutyronitrile (AIBN) acting as an initiator7. Boissiere et al. investigated

the design of biopolymer-based nanohybrids. They reported two technical

approaches for the design of hybrid capsules in the submicrometer size range,

the poly-L-lysine/alginate microparticles, which are promising carriers for drug

delivery devices. Moreover, they can be degraded by fibroblast cells.

Biopolymer-based microgels exhibit not only good biocompatibility.

Regarding their potential use as drug carriers, biodegradable microgels have

been widely viewed as promising candidates for their ability to be applied as

drug delivery vehicles. The mechanism of the biodegradable system was

introduced including the controlled uptake of bioactive guests, protection of the

embedded compounds from hydrolysis, degradation of the drug delivery

vehicles and controlled release of the encapsulated compounds8. Different

methods have been proposed to degrade the microgels, such as chemical

hydrolysis and enzymatic degradation9. The purpose of these delivery systems

is to encapsulate the drug which is stored in the drug delivery vehicles, protect

it from degradation, and subsequently selectively target it to a specific region in

a well-defined manner prompted by an external trigger.

Currently, polysaccharides, as the most abundant biopolymers in nature, are

being widely used in tissue engineering fields10. Their properties, such as non-

toxicity, biocompatibility, biodegradability and ease of chemical modification

in relation to synthetic polymers, make them attractive candidates as

biomaterials for drug delivery and tissue engineering11. Polysaccharides can be

utilized for the targeted and controlled delivery of drugs because they are non-

toxic, biofunctional, bioadhesive and biodegradable. Moreover, polysaccharides

can also be applied in drug engineering for tissue adhesives, surgical repair,


134

tissue regeneration and prolonging the targeting cells or the drug residence time

because they are bioadhesive12. These features have attracted a great deal of

interest because of their applicability in developing polysaccharide-based

biomaterials for biomedical applications as diverse as drug delivery, cell

encapsulation, regenerative medicine and tissue engineering13.

Natural polysaccharides are of great interest in developing colon-specific drug

delivery systems, in which the drugs are both active and protected against

hydrolysis in the stomach and small intestine, whilst also being capable of being

triggered and delivered when they enter into the colon region. Polysaccharide-

based microgels can be designed as a promising colon-specific drug targeting

matrix. The major strategies include the utilization of the protective coating on

the drug core, entrapment of the drug in biodegradable drug delivery vehicles

and the formulation of prodrugs depending on drug-saccharide conjugation.

Moreover, for local therapies targeting the colon, such as Crohn’s disease and

colon cancer, drug targeting reduces not only the necessary drug dose but also

the harmful adverse effects14.

Several polysaccharides, such as alginate15, dextran16, chitosan17, cellulose18,

pullulan19, hyaluronan20 and chondroitin sulfate21 are good candidates to be

applied as controlled or sustained drug release carriers at a targeted site for

possible biomedical purposes, e.g., regenerative medicine and tissue engineering

scaffolds22. Among these available polysaccharides, chitosan and dextran are

currently of great interest for biomedical use due to their uniquecharacteristics,

such as low toxicity23, low immunogenicity24, renewable resources25, the

abundant functional groups26, biocompatibility27 and biodegradability. For the

purpose of meeting different demands, distinct methods have been employed to

fabricate chitosan-based nanoparticles. A series of methods for chemical

modifications are capable of fabricating various chitosan-based microgels28.


135

Chitosan, a natural polycationic linear polysaccharide composed of

glucosamine units29 derived from the alkaline deacetylation of chitin, is

considered to be the most versatile biopolymer due to its unique physical and

chemical properties30. It has enriched functionalities, such as amine and

hydroxyl groups, which can allow for modifications with a variety of ligands31.

In addition, chitosan is a typical pH-responsive polymer that respond to changes

in physicochemical character with varying pH values through the protonation or

deprotonation of amino groups. Furthermore, injectable chitosan could be

degraded by enzymes in vivo, such as lysozyme and chitosanase, which render

it biodegradable for the controlled delivery of therapeutic medicines32.

However, the limited solubility of chitosan in neutral and alkaline solvents

could hinder its application in terms of the drug dissolution rate, influencing the

bioavailability of the oral drug, and thus posing challenges to the exploitation of

natural polysaccharides in drug delivery and tissue reconstruction33. The

solubility of chitosan mainly depends on its physicochemical properties. At low

pH, the amino groups in chitosan become protonated and positively charged,

resulting in it being cationic when in an acid solution. On the contrary, chitosan

is insoluble at high pH due to its deprotonated amino group34.

In order to address this problem, modified chitosan could be exploited to

enhance its reduced solubility in a neutral medium, like water, to allow for more

efficient drug absorption. After modifying with alkyne groups, chitosan can

dissolve in water. In this works, we will have developed a class of

polysaccharide-based microgels, comprising from self-crosslinking of water-

soluble alkyne-modified chitosan and azide-modified dextran. Through this

approach, a novel series of microgels were reported here in a one-step emulsion

polymerization procedure. This procedure did not need to utilize any additional

cross-linking agents because the pre-functionalized precursors can be cross-

linked directly in the presence of Copper(II) and catalysts through a Copper(II)-


136

catalyzed azide-alkyne (CuAAC) click reaction. Under sonication, the microgels

were prepared by inverse miniemulsion polymerization at room temperature.

Cross-linking density can be facilely tuned as a function of the ratio of moles of

the azide group to moles of the alkyne group by changing the degree of

substitution of the two pre-functionalized precursors. The pH-responsive

microgel particles were investigated using dynamic light scattering (DLS),

electrophoretic mobility and also transmission electron microscopy (TEM),

which characterized the degradation of microgel particles.

Moreover, the microgels are pH- and enzymatically-degradable and showed

good degradability in the presence of an alkali or dextranase. The enzymatic

degradation of the microgels can be triggered at pH 6.0 by a model enzyme,

dextranase, which is a special kind of bacterial enzyme present in the colon35.

Meanwhile, hydrolytic degradation takes place above pH 9.0, rendering the

microgels suitable for the controlled release of drugs in the colon. Furthermore,

the cytotoxicity in vitro was evaluated and these microgels showed no

significant cytotoxicity up to a concentration of 0.1 mg/mL. These microgels are

fabricated from biomaterials, making them highly suitable as tiny bioactive

devices for drug delivery systems. Therefore, an antibiotic, vancomycin

hydrochloride (VM), was encapsulated into the microgels and delivered in the

presence of an enzyme, dextranase, in the colon, suggesting that the DE-CH

microgel can be applied as a local antibiotic delivery system for colonic diseases.


4.2.1 Materials

Purification of chitosan of medium molecular weight (190000-310000 g/mol,

Sigma-Aldrich) was conducted based on a literature36. All other reagents were


137

commercially available and used without further purification. Dextran (Mw ~

40 kDa), 3-bromo-1-propanol (97%), sodium azide ( ≥ 99.5%), N,N-

Dimethylformamide (DMF, ≥ 99%), 2-(N-morpholino)ethanesulfonic acid,

(MES, ≥ 99%), 4-pentynoic acid (97%), L-ascorbic acid (99%), 1-1’-

carbonyldiimidazole (CDI, ≥ 97%), N-(3-dimethylaminopropyl)-N’-

ethylcarbodiimide hydrochloride (EDC, ≥ 99%), N-hydroxysuccinimide (NHS,

98%), Span 80, cyclohexane ( ≥ 99.8%), potassium phosphate (K2HPO4, ≥ 98%),

monobasic potassium phosphate (KH2PO4, ≥ 98%), sodium chloride ( ≥ 99%),

chloroform-d (CDCl3, 99.8 atom % D), deuterium oxide (D2O, 99.9 atom % D),

deuterium chloride (DCl, 99 atom % D), ethyl acetate ( ≥ 99%), magnesium

sulfate (99.5%), dimethyl sulfoxide (DMSO, 99.9%) and endo-dextranase from

Penicillium sp. (lyophilized powder, 11 units/mg solid) were purchased from

Sigma-Aldrich and used as obtained. Vancomycin hydrochloride (VM) was

bought from DUCHEFA Biochemie (Haarlem, Netherlands). Dialysis

membranes (MWCO = 12 kDa, 6 kDa and 3.5 kDa) were provided by Carl Roth.

Deionized water was used in reactions and also applied in the preparation of PBS

buffers from pH 5 to pH 8, and other buffers at pH 3, pH 4, pH 9 and pH 10 were

pH-adjusted buffers adjusted by 0.1 M HCl or 0.1 M NaOH.

4.2.2 Synthesis of 3-Azidopropyl Carbonylimidazole

CDI (1.76 g, 10.85 mmol) was dissolved in 40 mL of ethyl acetate under

continuous stirring, and then 1-azido-3-propanol (1.392 mL, 15 mmol) was

added dropwise to the vigorously stirred suspension until the solution became

clear. The reaction mixture was stirred at room temperature for 4 h and then the

prepared solution was washed 3 times with deionized water. The extracts were

dried overnight with anhydrous magnesium sulfate. After evaporating the

reaction solution, 3-azidopropyl carbonylimidazole (AP-CI) was obtained as the


138

liquid37. 1H NMR (CDCl3, 400MHz, δ in ppm): 1.89 (m, 2H, CH2-CH2-CH2),

3.43 (t, 2H, N3-CH2), 4.45 (t, 2H, CH2-O), 6.83 (s, 1H, C=CH-N), 7.34 (s, 1H,

N-CH=C), 8.08 (s, 1H, N-CH=N) (Fig. 3B).

4.2.3 Synthesis of Azide Modified Dextran (Dextran-

Azidopropylcarbonate)

Dextran (1 g, 0.025 mmol) containing 6.168 mmol of glucopyranose repeating

units and AP-CI (1.204 g, 6.16 mmol) were dissolved in 20 mL of anhydrous

DMSO under continuous stirring for 16 h. 0.301 g (1.54 mmol) or 1.204 g (6.16

mmol) of AP-CI was then added to this mixture. After stirring overnight at 50 °C

under a nitrogen atmosphere, the solutions were dialyzed (Mw = 3.5 kDa)

against deionized water for 5 days. A white fluffy product was received through

lyophilization37. 1H NMR (CDCl3, 400MHz, δ in ppm): 2.01 (2H, C≡C-CH2),

4.26 (2H, CH2-CH2-O), 4.92 (1Hdextran, O-C(CH)-O) (Fig. 3C).

4.2.4 Synthesis of Alkyne Modified Chitosan (Alkyne-Pendant

Chitosan)

A certain amount of chitosan and 4-pentynoic acid (Table 1) were dissolved

in an MES buffer (0.1 M, pH adjusted to 5.0) and bubbled with nitrogen. Next,

EDC (0.32 g, 1.65 mmol) and NHS (0.57 g, 4.95 mmol) were progressively

injected into the flask. The reaction was carried out at room temperature under

constant stirring, under a nitrogen atmosphere, for 16 h. The reaction solution

was transferred into a dialysis tube (Mw = 3.5 kDa) against deionized water for

5 days and lyophilized38. As shown in Table 1, alkyne-pendant chitosan is one

of the precursors for the click reaction which were synthesized with different


139

degrees of substitution (DS) of alkyne groups, ranging from 20 mol% to 80

mol%. 1H NMR (D2O/DCl, 400MHz, δ in ppm): 1.93 (1H, HC≡C), 2.29 (2H,

C-CH2 -C=O), 2.52 (2H, C≡C-CH2-C), 3.02 (1H2 of chitosan, C-CH(NH)-C), 3.47-

3.82 (H3-6 of chitosan) (Fig. 4).

Table 1. Alkyne-modified chitosan synthesized in this work.

Name of

sample

Chito-

san

(g)

-NH2 in

Chitosan

(mmol)

4-

penty-

noic

acid

(g)

-C≡C in

4-penty-

noic acid

(mmol)

Molar

ratio of

-C≡C to

-NH2

(mol:mol)

DS of

-C≡C

(mol%)

-NH2

amount

(mol%)

Alkyne-CH-1 0.100 0.496 0.054 0.546 1.100 78 0

Alkyne-CH-2 0.100 0.496 0.037 0.372 0.750 57 20

Alkyne-CH-3 0.100 0.496 0.024 0.248 0.500 37 40

Alkyne-CH-4 0.100 0.496 0.012 0.124 0.250 20 60

4.2.5 Synthesis of Microgels via Click Cross-linking Reactions

The microgels were synthesized via a CuAAC click reaction in inverse

miniemulsion. 1 mL of 0.1 M MES buffer was used as an aqueous phase and 10

mL of cyclohexane was applied as an organic phase containing 0.258 g of Span

80 as a surfactant. In the aqueous phase, the two pre-polymers, modified chitosan

and modified dextran, were mixed at varying azide:alkyne molar ratios from

1:0.5 to 1:2 (Table 2) and dissolved in the MES buffer. After total dissolution,

2.82 mg of Cu(II)Br2/PMDETA complex and 15.84 mg of ascorbic acid were

rapidly added into the prepared aqueous phase in order to initiate the CuAAC

click reaction. The preparation of the Cu(II)Br2/PMDETA complex followed the

procedures set out in the literature39. The mixture of an aqueous phase and an

organic phase was sonicated at a duty cycle of 50% and output control of 40%

for 10 min in ice-bath cooling whilst being simultaneously bubbled with


140

nitrogen. After sonication, the obtained miniemulsion was incubated in a 50 mL

Schlenk flask bubbling with nitrogen continuously for 24 h. The microgels were

washed by centrifugation at 10000 rpm and re-suspended in 20 mL of

cyclohexane. The washing procedures were repeated 3 times. The obtained

microgels were re-dispersed in DMSO to remove residual surfactants and

dialyzed (Mw = 12 kDa) against water before use. DE-CH microgels were

obtained after freeze-drying.

Scheme 1. Synthesis of microgels by cross-linking of alkyne and azide modified pre-

polymers.

Table 2. The CuAAC click reaction of modified chitosan and dextran.

Name of

sample

DS of -N3 in

modified

dextran (%)

DS of -C≡C in

modified chitosan

(%)

Amount of -NH2

in modified

chitosan (%)

-N3: -C≡C in

microgels

(molar ratio)

DE-CH-1 3.33 78 0 1:2

DE-CH-2 3.33 57 20 1:1.5

DE-CH-3 3.33 37 40 1:1

DE-CH-4 3.33 20 60 1:0.5


141

4.2.6 Characterization Methods

The 1H NMR measurements were recorded at 400 MHz using a Bruker DPX-

400 FT NMR spectrometer. Chloroform-d (CDCl3), deuterium oxide (D2O) and

deuterium chloride (DCl) were used as solvents in these measurements.

Fourier transmission infrared (FTIR) measurements were carried out at room

temperature using a Nexus 470 (Thermo Nicolet). The lyophilized microgels

were then mixed with KBr tablets.

Transmission electron microscopy (TEM) observations were recorded using

a Zeiss LIBRA 120 at an accelerating voltage of 120 kV. The diluted microgel

solution was drop-cast on a plasma-treated formvar-carbon-coated 400 mesh

copper grid and dried at room temperature overnight.

Dynamic light scattering (DLS) measurements were conducted using a

commercial laser dynamic light scattering spectrometer (ALV/DLS/SLS-5000)

at a scattering angle of 90°. The spectrometer was equipped with an ALV/LSE-

5004 multi-𝜏 digital time correlator and an ALV/CGS-3 laser goniometer system

at a wavelength of 632.8 nm.

Electrophoretic mobility measurements were performed using a Malvern

Zetasizer Nano ZS particle analyzer. After an equilibration time of 120 s, the

measurements were performed at a pH range from 3 to 11 at 25 °C.

UV-Vis spectra were recorded on a Perkin Elmer Lambda 35 UV-Vis

spectrometer.

4.2.7 Alkaline-Induced Degradation

Two types of degradation pathways are explored for the synthesized

microgels: the pH-dependent hydrolytic degradation and enzymatic degradation.

To study the degradation behaviors, FTIR, 1H NMR, DLS and TEM


142

measurements were used to test the variation in chemical composition, swelling

degree and particle morphology. For pH-triggered hydrolytic degradation, the

obtained microgels, immersed in buffers at different pH values, were evaluated

by DLS measurements. At predetermined time intervals, samples were added to

the pH-adjusted buffers in the pH range of 3-10 and the size of the microgels

was obtained by DLS measurements. Moreover, in order to estimate the extent

of alkaline-induced degradation, the series of microgel dispersions were

dissolved in a pH-adjusted buffer at pH 10, monitored by DLS. In addition, at

predetermined time points, the degradation products were recorded using FTIR

and 1H NMR.

Furthermore, the morphology of the degraded microgels was observed by

TEM. TEM measurements were conducted after the addition of the sample in

pH 10 buffer at 1 h, 1 day and 7 days.

4.2.8 Enzymatic Degradation

DLS, FTIR, 1H NMR and TEM measurements were taken to investigate the

enzymatic degradation behavior of microgels in vitro. A microgel dispersion

mixed with dextranase (0.2 U/mL) was incubated in PBS buffers (pH 6) at 37°C

for the degradation experiments. The particle size was evaluated via DLS at

predetermined time intervals, with shorter time intervals during the first 20 min

and longer intervals over the period till 1 day. FTIR and 1H NMR were recorded

to value the chemical composition of the obtained degradation products at the

indicated time points. TEM observations were carried out to explore the changes

in particle morphology.


143


Drug loaded microgels were prepared by dissolving 5 mg of a DE-CH-3

microgel and 1 mg of VM in 5 mL of deionized water at room temperature for

24 h. The mixtures were centrifuged at 13000 rpm for 30 min and the process

was repeated 3 times. The amount of unloaded free VM of the final supernatant

was evaluated using a UV-Vis spectrophotometer at λ= 280 nm (Fig. 16A) to

determine the amount of unloaded free VM. The drug loading efficiency was

calculated according to Equ. (1):

Drug loading efficiency = Mtotal VM−Mfree VM

Mtotal VM (1)

where Mfree VM and Mtotal VM represent the mass of unloaded and initial VM,

respectively.

The in vitro drug release profiles of VM-loaded DE-CH-3 microgels

(VM@DE-CH-3) were investigated under three different environments, (a) with

dextranase (0.2 U/mL) at pH 6 under 37 °C, (b) without dextranase at pH 6 under

37 °C and (c) buffer at pH 10 under room temperature. In order to test the drug

release amount, microgels without VM loading were used as the control. A

certain amount of VM-loaded microgels and the control sample were added into

1 mL of buffers (a, b or c). After they were transferred into a dialysis bag

(MWCO = 6 kDa) and immersed in 19 mL of the same buffer, the release

experiments were conducted. 1 mL of the release medium was withdrawn to

determine the VM release amount via UV-Vis analysis at predetermined time

points and the same amount of fresh buffer was replenished to maintain the same

total volume (Fig. 16B).


144

4.2.10 Cytotoxicity Study in Vitro

The mouse fibroblast cell line, L-929 (ATCC CCL-1), was cultured to a

density of 1 × 104 cells per well in a 96-well microtiter plate overnight at 37 °C

under 5% CO2 and 95% air incubator for cell attachment. The cell viability

testing was carried out via the XTT cell proliferation assay kit (Cat. No. 30-

1011K, ATCC) according to the manufacturer’s instructions. After being

incubated with microgels at doses of 0.100, 0.010 and 0.001 mg/mL for 24 hours

in 5% CO2 and 95% air at 37 °C, the XTT (sodium 2,3-bis-(2-methoxy-4-nitro-

5-sulfophenyl)-5-[(phenylamino)-carbonyl]-2H-tetrazolium) inner salt and

PMS (N-methyl dibenzopyrazine methyl sulfate), as the electron carrier were

added to each well according to the supplier protocol. To determine the living

cell numbers, the optical absorbance was measured at 490 nm (reference

wavelength 630 nm) on a microtiter plate reader (Detection Microplate Reader

from BioTek).

4.2.11 Statistical Analysis

All analyses were performed in triplicate. The statistical significance

was analyzed using one-way ANOVA. A p value of 0.05 was used to

determine the statistical significance level and the data were marked with

(*) for p < 0.05, (**) for p < 0.01, and (***) for p < 0.001, respectively.


145


4.3.1 Chemical Structure of Microgels

Microgels were synthesized by cross-linking water-soluble functional

biopolymer in an inverse (water-in-oil; W/O) miniemulsion by CuAAC click

reaction. This was ascertained by the cross-linking of complementary reactive

side groups integrated into dextran and chitosan backbones without any other

cross-linking agent as shown in Scheme 1. The alkyne and azide modifications

of the chitosan and dextran were determined by 1H NMR spectra, respectively

(Fig. 3,4). The purification steps were conducted to remove the remaining

surfactant of Span 80, confirmed by 1H NMR (Fig. 1). In the purified microgel,

the two peaks c and d disappeared which were assigned from characteristic peaks

in Span 80.

Fig. 1. 1H NMR spectra of Span 80, two prepolymers (modified chitosan or dextran),

and DE-CH-3 microgel with/without purification in D2O.


146

The chemical structure of the obtained microgels was confirmed by FTIR

spectra (Fig. 2A). The peaks at 2106 cm-1 and 3311 cm-1 are due to the stretching

modes of the unreacted -N3 and C-H of alkyne groups, respectively. This

indicates that a small number of unreacted modification groups were still left in

the microgel which could further be utilized for post-modification. The peak in

the region (950-1300 cm-1) is assigned to the C-O stretching vibration. The peaks

at 1461 cm-1 and 1733 cm-1 are attributed to the stretching vibration of the -CH3

and C=O groups, respectively. The peaks at 2853 cm-1 and 2922 cm-1 correspond

to the stretching vibration of the C-H and O-H groups, respectively37. The peaks

at 1637 cm-1 and 3008 cm-1 attributed to C=N and C-H stretching of the 1,2,4-

triazole ring, confirming the CuAAC click reaction proceeded successfully40.

Scheme 2. Schematic illustration of the ideal network structures in synthesized

microgel samples.


147

Fig. 2. (A) FTIR spectra of pre-polymer building blocks (chitosan, dextran-

azidopropylcarbonate) and the series of synthesized microgels. (B) Enlarged FTIR

spectra at 1500-2500 cm-1 and (C) linear fit of molar ratio of C≡C to -N3 and intensity

of the C=N (1637 cm-1) IR bands present before and after the microgels synthesis in

this work.


148

Fig. 3. 1H NMR spectra of three types of azide modified dextran (dextran-

azidopropylcarbonate) synthesis, exhibiting (A) 1-azido-3-propanol, (B) 3-azidopropyl

carbonylimidazole, and (C) dextran-azidopropylcarbonate in CDCl3.

The enlarged part of the FTIR spectra at 1500-2500 cm-1 is shown in Fig. 2B.

The peak at 1637 cm-1 is assigned to the C=N stretching of the 1,2,4-triazole ring.

Comparing four different microgel samples, the peak intensity at 1637 cm-1

decreases from DE-CH-1 microgel to DE-CH-4 microgel, presenting the same

trend as the varying tendency of azide:alkyne molar ratios in the microgel

synthesis process changing from 1:0.5, 1:1, 1:1.5 to 1:2. Fig. 2C indicates that

the linear relationship between the intensity of the 1,2,4-triazole ring and the

molar ratio of alkyne to the azide group revealed a good linear relation. Scheme

2 illustrates the synthesis of all microgel samples with the different crosslinking

densities while changing the DS of two precursors, the DS of the alkyne group

in chitosan from highest (DS = 78) to lowest (DS = 20) and dextran with a fixed

degree of substitution of the azide group (DS = 3.33). These obtained microgels

with changed particle sizes were developed by tuning the number of clickable


149

functionalities in modified precursors, and thus changing the cross-linking

density of the microgels.

Fig. 4. 1H NMR spectra for four steps of alkyne modified chitosan (alkyne-pendant

chitosan) synthesis in D2O/DCl and the enlarged 1H NMR spectrum from 1.5 to 4.0

ppm (the integrated areas under the peaks were measured by MestRec NMR software).


As shown in Fig. 5, DLS and electrophoretic mobility were conducted to

observe the microgels’ pH-responsive behavior. As pH ranged from 3 to 11, the

DE-CH-2, DE-CH-3 and DE-CH-4 microgels exhibited a considerable change

in hydrodynamic radius due to the protonation or deprotonation of the amino


150

groups (-NH2) of chitosan in aqueous media in response to pH changes, which

are the pH-dependent donors (Fig. 5A). In an acidic environment, protonated

amino groups (-NH3+) carried net positive charges, which resulted in the

swelling of microgels due to their electrostatic repulsion and the presence of

counterions. The DE-CH-2, DE-CH-3 and DE-CH-4 samples showed a similar

trend but with different swelling ratios. Compared to the other samples, the DE-

CH-4 microgel exhibited the highest swelling ratio, Rh(pH 3)/Rh(pH 11) = 2.7,

because it had the lowest cross-linking density and the highest amount of amino

groups of all the microgels. The electrophoretic mobility tests of the microgels

indicate that the positive charges decreased as the pH varied from pH 3 to pH 8.

The microgels became negatively charged above pH 9 due to the effect of the

dextran’s OH- groups of (Fig. 5B). It is also indicated that hydrogen bonding

was formed between the O- groups of dextran and the hydroxyl group in

microgels41. On the contrary, the hydrodynamic radius of DE-CH-1 showed no

obvious difference over the whole pH range because of the fewer ionizable

groups and the weak electrostatic repulsion in the microgel network.

Fig. 5. (A) Hydrodynamic radius and (B) electrophoretic mobility of microgels in pH-

adjusted PBS buffers.


151


4.3.3.1 Effect of pH on Microgel Degradation

The degradation behaviors of the obtained microgels were carried out over the

whole pH range from 3 to 10. The size of the DE-CH-3 microgel exhibited no

obvious changes at pH 3-7 (Fig. 6). As pH increased above 9, degradation

occurred, indicating that the degradation extent is highly dependent on the

environmental pH value. In an alkaline solution, the microgels gradually

degraded due to the cleavage of carbonate ester bonds in dextran.

Fig. 6. Degradation processes of DE-CH-3 microgel in pH-adjusted PBS buffers with

various pH values, ranging from 3 to 10, measured by DLS as a function of time.


152

Fig. 7. Degradation processes of microgels at pH-adjusted PBS buffer (pH 10) over

time determined by DLS.

The degradation studies mentioned above showed that microgel degradation

can be tuned by pH values in the environment and occur in a basic medium.

Therefore, the alkaline-catalyzed degradation behavior of the DE-CH-2, DE-

CH-3 and DE-CH-4 microgels can be investigated by DLS, 1H NMR, FTIR and

TEM to observe the variations in the size, chemical composition and

morphology of microgels in an alkaline environment. At pH 10, the

hydrodynamic radius of the microgels decreased over time (Fig. 7). During the

first 10 min, the particle size sharply decreased, indicating the collapse in

microgels, which was probably due to the deprotonation of amino groups in the

microgel network. The microgel size gradually decreased from 10 min to 4 days.

It also can be observed that the degradation profiles are very similar among

different microgel samples due to the fact that all of the microgels studied

possessed the same amount of dextran in their structure which was subjected to

pH-dependent hydrolytic degradation, whereas the variation in the cross-linking

density in microgels has no influence on the degradation rate.


153

Fig. 8. 1H NMR spectra of DE-CH-3 and degradation products after 300 min and 4

days degradation at pH 10.

1H NMR and FTIR spectra were conducted to characterize the variation in the

chemical composition of degraded microgels over time. After degradation for

300 min and 4 days, 1H NMR spectroscopy showed that the signal a at 4.21 ppm

in the spectrum of DE-CH-3 disappeared when compared to the DE-CH-3

microgel (Fig. 8). A new signal n at 3.70 ppm appeared after 4 days of

degradation, indicating the cleavage of the carbonate ester bond. FTIR spectra

showed that the peak at 1733 cm-1 weakened after degradation which was due to

the hydrolysis of the carbonate ester bond (Fig. 9). The peak at 1439 cm-1 is

stronger because of the increased amount of -OH group in degraded microgels.


154

Fig. 9. FTIR spectra of DE-CH-3 and degradation products after 300 min and 4 days

degradation at pH 10.

Scheme 3. Schematic illustration of the pH-triggered degradation of microgels.


155

As shown in Fig. 10, the variation in microgel morphology was observed via

TEM. The microgels changed their morphology from a spherical shape to the

smaller polymer clusters after 1 h, 1 day and 7 days.

Fig. 10. TEM images of DE-CH-3 microgel over time before and after the degradation

in buffers (pH 10).

4.3.3.2 Effect of the Enzyme on Microgel Degradation

Moreover, 1,6-α-glucosidic linkages of dextran can be cleaved by dextranases

and degradation of microgels can be carried out in the presence of the enzyme

in a pH 6 buffer at 37 °C42. DLS, 1H NMR, FTIR spectra and TEM

measurements were introduced to investigate the degradation behaviors of

microgels. A fast degradation process occurred during the first 30 min indicating

the dextran chains in microgels can be rapidly cleaved during this stage (Fig. 11).

The degradation rates then decreased, standing for the second degradation stage.


156

This is probably due to the fact that the microgels possessed more cross-linking

points in their interiors than at their surfaces. Therefore, the dextranase accessed

the microgels for surface erosion at a rapid rate, and then changed into the

interior of the particles at a slower degradation rate.

Fig. 11. Degradation of microgels with dextranase over time, as determined by DLS.

1H NMR results showed that an anomeric signal d at 4.88 ppm, assigned to

typical 1,6-α-glucosidic linkages, became weaker at 50 min and 300 min of

degradation (Fig. 12). As shown in Fig. 13, FTIR spectra indicated that the

stretching vibration of -OH at 1456 cm-1 becomes stronger due to the fact that

the amount of -OH groups increased in degraded microgels. TEM images

observed the degradation process over time. It is indicated that the microgels

deformed after 1 h and broke into small fragments after 1 day (Fig. 14).


157

Fig. 12. 1H NMR spectra of DE-CH-3 and degradation products after degradation for

50 min and 300 min.

Scheme 4. Schematic illustration of pH-triggered microgels degradation.


158

Fig. 13. FTIR spectra of DE-CH-3 and degradation products after degradation for 50

min and 300 min.

Fig. 14. TEM images of DE-CH-3 microgel before and after the degradation in buffers

(pH 6).

4.3.4 Cytotoxicity Evaluation

As drug delivery vehicles, the most important point is that the microgels are

non-toxic and safe, thus allowing them to be developed for biomedical


159

applications. XTT cell proliferation assay using L-929 cells was performed to

test the cytotoxicity of microgels at various concentrations. Fig. 15 shows the

cytotoxicity of microgels exhibited in a dose-dependent manner. At doses

ranging from 0.001 mg/mL to 0.100 mg/mL, the microgels varied from non-

cytotoxic to a little toxic, when compared to the control sample. When the

concentration increase to 1 mg/mL, the microgels show decreased cell survival

because of the protonated amino groups in chitosan which exhibit toxicity43. In

addition, the cytotoxicity among the microgel samples showed no significant

difference at the same dose, indicating that the microgels can be used as a drug

delivery system for biomedical and biotechnological applications in doses below

0.100 mg/mL.

Fig. 15. Cytotoxicity in vitro of the microgels at various concentrations against L-929

cell lines.


An antibiotic glycopeptide, VM, was selected as a model drug to investigate

the drug loading and triggered release profiles of the obtained biohybrid


160

microgels. VM was encapsulated into the microgels in a deionized water

environment by electrostatic interactions between positively charged VM and

negatively charged microgels due to the effect of the existence of OH- groups of

dextran, thus attending to the formation of hydrogen bonding between the O-

groups of dextran and hydroxyl groups in microgels. The results showed that the

microgels carry different charges in a PBS buffer of pH 7 or in deionized water.

This may be due to the reason that the increased ionic strength of the solvent

allows for the compression of the counter-ion cloud (i.e., electric double layer),

which impacts the surface potential44. After 24 h of VM-encapsulating, the

loaded VM amount in microgels was estimated by a UV-Vis spectroscopy (Fig.

16).

Fig. 16. (A) UV-Vis spectra of a DE-CH-3 microgel, vancomycin hydrochloride (VM)

and a VM-loaded DE-CH-3 microgel (VM@DE-CH-3). (B) The standard calibration

curve of vancomycin hydrochloride (VM).

Furthermore, the drug release behaviors were investigated in different

conditions. The calculated drug loading efficiency by Equ. (1) was up to 93.67%

and the VM loading in microgels was 187.34 μg/mg. Dextranase, an enzyme


161

present in human colonic content, which can degrade dextran, was chosen as a

trigger for drug release.

Fig. 17. In vitro release profiles of vancomycin hydrochloride from DE-CH-3

microgels in buffers at pH 10 or pH 6 with or without dextranase under 37°C.

As shown in Fig. 17, both the dextranase- and pH-triggered drug-release

process of VM-loaded DE-CH microgels were studied to explore their potential

for precise drug delivery. At pH 6, a VM-loaded DE-CH-3 microgel in the

presence of dextranase (0.2 U/mL) showed an initial fast release of VM during

the first 1 h. This can be attributed to the rapid enzymatic degradation of the

microgel shell, where the highest amount of VM is localized. After 1 day, the

release amount was up to 89.47%. At pH 10, the released amount of VM from

the DE-CH-3 microgel showed a sustained release with a less rapid delivery rate

due to the slower microgel degradation behavior in an alkaline environment. On

the contrary, at pH 6, without adding dextranase, the release rate of VM from the

DE-CH-3 microgel was much slower. This effect may be attributable to the weak


162

bonds between VM molecules and microgels, thus inducing the VM leaching.

We believe that this leaching effect can be reduced via balancing the charge

density within microgel networks. The drug release results indicated that the

synthesized microgels showed broad applicability as a potential carrier for

colonic drug delivery.

4.4 Conclusion

Two modified biopolymers, chitosan and dextran, were applied to synthesize

a series of pH-sensitive and dual stimuli-responsive microgels with different

cross-linking densities via a facile “click chemistry” in an inverse miniemulsion.

The microgels can be obtained by cross-linking the two precursors, alkyne-

modified chitosan and azide-modified dextran, by means of CuAAC click

reaction without extra cross-linkers, which were characterized by 1H NMR and

FTIR. These microgels are pH-responsive and exhibit a sharp charge switch in

response to varying physiological pH values. Furthermore, these microgels show

pH- or enzyme-triggered degradation properties. They can be degraded above

pH 9 or in the presence of dextranase due to the hydrolysis of carbonate esters

in microgels or 1,6-α-glucosidic linkages in dextran structure, which were

characterized by DLS, 1H NMR, FTIR and TEM, showing the variation in size,

chemical composition and morphology during the degradation process. In

addition, drugs with positive charges, such as VM, an antibiotic, can be

encapsulated spontaneously into the microgels carrying negative charges in

water via electronic interaction. Meanwhile, the VM release can be controlled

by the pH conditions or an enzyme in the colon, e.g., dextranase. These

biodegradable microgels were demonstrated to have low cytotoxicity via XTT

cell proliferation assay using L-929 cells. Therefore, these microgel matrices are


163

expected to have potential applications as bioactive delivery vehicles for the

release of the drugs at specific sites, such as the colonic region.


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38. Bao, H. Q.; Li, L.; Leong, W. C.; Gan, L. H., Thermo-responsive association of

chitosan-graft-poly(N-isopropylacrylamide) in aqueous solutions. J. Phys. Chem. B

2010, 114 (32), 10666-10673.

39. Lonsdale, D. E.; Bell, C. A.; Monteiro, M. J., Strategy for rapid and high-purity

monocyclic polymers by CuAAC "Click" Reactions. Macromolecules 2010, 43 (7),

3331-3339.

40. Plech, T.; Wujec, M.; Majewska, M.; Kosikowska, U.; Malm, A.,

Microbiologically active Mannich bases derived from 1,2,4-triazoles. The effect of C-

5 substituent on antibacterial activity. Med. Chem. Res. 2013, 22 (5), 2531-2537.


166

41. (a) Khalkhali, M.; Sadighian, S.; Rostamizadeh, K.; Khoeini, F.; Naghibi, M.;

Bayat, N.; Habibizadeh, M.; Hamidi, M., Synthesis and characterization of dextran

coated magnetite nanoparticles for diagnostics and therapy. Bioimpacts 2015, 5 (3),

141-150; (b) Gardner, B., The effect of dextrans on zeta potential. Proc. Soc. Exp. Biol.

Med. 1969, 131 (4), 1115-1118.

42. (a) Yang, L.; Zhou, N.; Tian, Y., Purification, characterization, and biocatalytic

potential of a novel dextranase from Chaetomium globosum. Biotechnol. Lett. 2018,

40 (9), 1407-1418; (b) Shahid, F.; Aman, A.; Pervez, S.; Ul Qader, S. A., Degradation

of long chain polymer (dextran) using thermostable dextranase from hydrothermal

spring isolate (bacillus megaterium). Geomicrobiol. J. 2019, 36 (8), 683-693.

43. Fernandes, J. C.; Qiu, X. P.; Winnik, F. M.; Benderdour, M.; Zhang, X. L.; Dai,

K. R.; Shi, Q., Low molecular weight chitosan conjugated with folate for siRNA

delivery in vitro: optimization studies. Int J Nanomedicine 2012, 7, 5833-5845.

44. Sharma, A.; Cornejo, C.; Mihalic, J.; Geyh, A.; Bordelon, D. E.; Korangath, P.;

Westphal, F.; Gruettner, C.; Ivkov, R., Physical characterization and in vivo organ

distribution of coated iron oxide nanoparticles. Sci. Rep. 2018, 8 (1), 4916.

5. Conclusion and Outlook

167


In recent years, synthetic microgels have generally been prepared through the

polymerization of different synthetic monomers in the presence of

multifunctional cross-linkers for applications in tissue engineering, biomedical

devices, bionanotechnology and drug delivery. Biopolymer-based microgels

have recently attracted a great deal of attention in the fields of drug delivery,

tissue engineering, catalyst, an anode material and food applications due to their

unique properties including synthetic counterparts as well as biopolymers, such

as biodegradable, abundant in nature, renewable, nontoxic, and relatively cheap.

Moreover, biopolymer-based microgels possess diverse functional groups

including hydroxyl, amino, and carboxylic acid groups. The functional groups

can be applied in crosslinking with the functional cross-linkers for further

bioconjugation applications. A typical example of naturally occurring

biopolymers is chitosan which is a biopolymer made up of β-(1-4)-linked 2-

amino-deoxy-D-glucosamine units. Owing to its unique properties, such as

biodegradability, renewability, and abundance in nature, non-toxicity and low-

cost, it can be developed to prepare biodegradable microgels for tissue

engineering scaffolds and drug delivery carriers.

This Thesis introduced new approaches for the synthesis of chitosan-based

microgels for biomedical applications. Chapter 1 introduced the properties and

applications of functional microgels including biopolymer-conductive polymer-

based microgels. The biocompatible, biodegradable and pH-sensitive properties


168

of chitosan endow it with features well suited to fabricate microgels for drug

delivery, functional coatings, tissue regeneration and water filtration.

In Chapter 2, pH- and redox-sensitive microgels based on chitosan and

hydroquinone were developed. These microgels were formed via physical

crosslinking and hydrogen bonding, without the addition of other cross-linkers.

The obtained microgels were able to encapsulate an anticancer drug, DOX, and

could be biodegraded in the presence of an enzyme, lysozyme, thus releasing

DOX. These electroactive microgels can be utilized in diverse fields including

therapy of tumors, tissue engineering and energy storage.

Chapter 3 describes the chitosan-polyaniline conductive microgels

synthesized in an inverse W/O miniemulsion. The graft of polyaniline endows

the microgels with electrochromic behavior and conductivity. Moreover,

chitosan was chosen as the matrix, as it can be degraded by an enzyme, lysozyme.

Due to their enzymatic degradation behavior, these microgels exhibited pH-

sensitivity, conductivity and biodegradability that can be utilized as a good

candidate for biomedical application.

In Chapter 4, two modified biopolymers, chitosan and dextran, were cross-

linked without extra linkers via a CuAAC click reaction. These microgels were

pH-sensitive and dual-degradable in the presence of an alkaline environment or

an enzyme in the colon, dextranase. Additionally, these biodegradable and

biocompatible microgels can encapsulate an antibiotic, VM, and released it in a

controlled manner, which demonstrated their potential for use in developing

colon-specific drug delivery carriers.

For future research to improve the clinical application of the chitosan-based

nanoplatforms, the following directions can be considered: 1) Designing and

fabricating chitosan-based microgels integrated with various biologics to form a

bio-hybrid system. With recent advancements in synthetic biology, biomimetic

surfaces such as membranes from red blood cells (RBC), platelets, leukocyte,


169

cancer cells, stem cells, immune cells and platelets, have been extracted as the

membrane source to prepare bioinspired theranostic nanosystems that are

responsive to certain signals for various purposes1. Cancer cell membranes are

getting more attention for use as bio-stealth material nanoparticle coatings. For

anticancer therapeutic systems, cancer cell membrane cloaked nanoparticle

system can achieve a variety of properties, such as prolonged blood circulation,

immune escape, resistance to macrophage uptake and homologous cancer cell

targeting capabilities which allows them to be used as coating biomaterials to

functionalize synthetic nanoparticles2. Thus, the integration of chitosan-based

microgels functionalized with cell membranes becomes a potential candidate for

biotechnological clinical applications. 2) Incorporation with hierarchical

targeting drug release to obtain smart drug delivery. In recent years, stimuli‐

responsive nanocarriers have been exploited as drug delivery systems with

targeted drug release behaviors. Therefore, designing chitosan-based microgels

modified with one of diverse targeting ligand can be employed to cure a specific

cancer3. 3) The combination of ultrasound-targeted microbubble destruction

(UTMD) as a passive targeting technique has been extensively used for tumor

chemotherapy which improves the permeability of cancer cells, thus enhancing

the cellular uptake of drugs or genes4. In some cancers, such as pancreatic cancer

(PaCa) which has unique physical barriers, the dense extracellular matrix (ECM)

and hypovascular networks, prevent the penetration of chemotherapeutic drugs,

leading the treatment to be ineffective5. The ideal strategy is to enlarge the

permeability of vessels and the tumor cells to enhance drug delivery. Therefore,

in order to overcome the physical barriers of solid pancreatic tumors, UTMD

was applied to enhance cell membrane permeability and promote the

endocytosis of nanoparticles. Chitosan-based microgels can be employed to

prepare the nanocarriers with UTMD technology to ensure the effectiveness of

chemotherapy.


170

In conclusion, this Thesis introduced the properties and applications of

biopolymer based-microgels and also fabricated chitosan-based microgels

incorporate with conductive polymers or biopolymers for biomedical

applications.


1. (a) Parodi, A.; Quattrocchi, N.; van de Ven, A. L.; Chiappini, C.;

Evangelopoulos, M.; Martinez, J. O.; Brown, B. S.; Khaled, S. Z.; Yazdi, I. K.; Enzo,

M. V.; Isenhart, L.; Ferrari, M.; Tasciotti, E., Synthetic nanoparticles functionalized

with biomimetic leukocyte membranes possess cell-like functions. Nat. Nanotechnol.

2013, 8 (1), 61-68; (b) Stephan, M. T.; Irvine, D. J., Enhancing cell therapies from the

outside in: cell surface engineering using synthetic nanomaterials. Nano Today 2011,

6 (3), 309-325.

2. Rao, L.; Bu, L.; Cai, B.; Xu, J.; Li, A.; Zhang, W.; Sun, Z.; Guo, S.; Liu, W.;

Wang, T.; Zhao, X., Cancer cell membrane-coated upconversion nanoprobes for highly

specific tumor imaging. Adv. Mater. 2016, 28 (18), 3460-3466.

3. Mura, S.; Nicolas, J.; Couvreur, P., Stimuli-responsive nanocarriers for drug

delivery. Nat. Mater. 2013, 12 (11), 991-1003.

4. Gao, F.; Wu, J.; Niu, S.; Sun, T.; Li, F.; Bai, Y.; Jin, L.; Lin, L.; Shi, Q.; Zhu,

L.; Du, L., Biodegradable, pH-sensitive hollow mesoporous organosilica nanoparticle

(HMON) with controlled release of pirfenidone and ultrasound-target-microbubble-

destruction (UTMD) for pancreatic cancer treatment. Theranostics 2019, 9 (20), 6002-

6018.

5. Whatcott, C. J.; Diep, C. H.; Jiang, P.; Watanabe, A.; LoBello, J.; Sima, C.;

Hostetter, G.; Shepard, H. M.; Von Hoff, D. D.; Han, H., Desmoplasia in primary

tumors and metastatic lesions of pancreatic cancer. Clin. Cancer Res. 2015, 21 (15),

3561-3568.

6. Acknowledgement

171

6. Acknowledgement

Upon the completion of this Thesis, I would like to express my gratitude to

those who have offered me encouragement and support during my doctoral study.

Firstly, I would like to express my most heartfelt gratitude to my respectable

supervisor Prof. Dr. Andrij Pich who provided me the Ph.D. position as a CSC

scholarship holder. He patiently guides me, encourages me and his suggestions

are beneficial to me a lot. He provided me great help in selecting the research

topic, preparing the presentation, writing the paper and thesis, and correcting the

errors. In the preparation of the Thesis, he spent much time reading each draft

including the submitted papers, and provided me lots of valuable suggestions

and comments. Without his patient help and expert guidance, all the work and

the thesis would not be finished.

Secondly, I would like to present my deepest gratitude to my second

supervisor Prof. Dr. Felix A. Plamper who give me considerable help. During

the preparation of the paper whenever I sent him this article, he always gave me

lots of comments and suggestions that really improved my article a lot. For the

questions in the article, he has done a great favor to check and correct them to

be better.

Thirdly, I greatly thank all of the members of the Pich Group and DWI

members who offered me valuable help during the years of my study here. In

addition, I greatly appreciate Dr. Smriti Singh, Dr. Olga Mergel, Dr. Huan Peng

and Xin Li, who had cooperated the part of the work of the thesis and give me a

6. Acknowledgement

172

lot of support and help. The special thanks for Susanne Braun. She helped me to

do the nice translation and also corrections.

Finally, I would like to thank my family who gave me continuous support and

encouragement. They have always helped me out of difficulties without

complaint and also the thanks give to my friends who have put considerable time

and effort to help me insist on working on my studies.

Helin

05.12.2020

7. List of Publications

173


[1] H. Li, O. Mergel, P. Jain, X. Li, H. Peng, K. Rahimi, S. Singh, F. A.

Plamper, A. Pich, Electroactive and Degradable Supramolecular Microgels. Soft

Matter 2019, 15 (42), 8589-8602.

[2] I. V. Novikov, M. A. Pigaleva, E. E. Levin, S. S. Abramchuk, A. V.

Naumkin, H. Li, A. Pich, M. O. Gallyamov, The Mechanism of Stabilization of

Silver Nanoparticles by Chitosan in Carbonic Acid Solutions. Colloid Polym.

Sci. 2020, 298 (9), 1135-1148.

[3] H. Li, X. Li, P. Jain, H. Peng, K. Rahimi, S. Singh, A.Pich, Dual-

Degradable Biohybrid Microgels by Direct Cross-Linking of Chitosan and

Dextran Using Azide-Alkyne Cycloaddition. Biomacromolecules 2020, 21 (12),

4933-4944.

[4] X. Li, H. Li, C. Zhang, A. Pich, L. Xing, X. Shi, Intelligent Nanogels

with Self-Adaptive Responsiveness for Improved Tumor Drug Delivery and

Augmented Chemotherapy, Bioact. Mater. 2021, 6 (10), 3473-3484.

[5] X. Li, H. Sun, H. Li, C. Hu, Y. Luo, X. Shi, A. Pich, Multi-Responsive

Biodegradable Cationic Nanogels for Highly Efficient Treatment of Tumors,

Adv. Funct. Mater. 2021, 2100227.


174

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Responsive chitosan-based microgels