responsive chitosan-based microgels
TRANSCRIPT
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.
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For My Family
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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
4.5 References and Notes ..................................................................................... 163
5. Conclusion and Outlook ...................................................................................... 167
5.1 References and Notes ..................................................................................... 170
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
Summary/Zussamenfassung
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.
Summary/Zussamenfassung
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).
Summary/Zussamenfassung
4
Summary/Zussamenfassung
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
Summary/Zussamenfassung
6
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.
Summary/Zussamenfassung
7
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.
Summary/Zussamenfassung
8
List of Abbreviations
9
List of Abbreviations
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
List of Abbreviations
10
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
List of Abbreviations
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)
List of Abbreviations
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
List of Abbreviations
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
List of Abbreviations
14
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.
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-
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.
1. Introduction
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.
1.6 References and Notes
1. Kyrey, T.; Witte, J.; Pipich, V.; Feoktystov, A.; Koutsioubas, A.; Vezhlev, E.;
Frielinghaus, H.; von Klitzing, R.; Wellert, S.; Holderer, O., Influence of the cross-
linker content on adsorbed functionalised microgel coatings. Polymer 2019, 169, 29-
35.
2. Nöth, M.; Gau, E.; Jung, F.; Davari, M. D.; El-Awaad, I.; Pich, A.;
Schwaneberg, U., Biocatalytic microgels (μ-Gelzymes): synthesis, concepts, and
emerging applications. Green Chem. 2020.
3. Newsom, J. P.; Payne, K. A.; Krebs, M. D., Microgels: modular, tunable
constructs for tissue regeneration. Acta Biomater. 2019, 88, 32-41.
4. Li, F.; Lyu, D.; Liu, S.; Guo, W., DNA hydrogels and microgels for biosensing
and biomedical applications. Adv. Mater. 2020, 32 (3), 1806538.
5. Caputo, T. M.; Aliberti, A.; Cusano, A. M.; Ruvo, M.; Cutolo, A.; Cusano, A.,
Stimuli-responsive hybrid microgels for controlled drug delivery: Sorafenib as a model
drug. J. Appl. Polym. Sci. n/a (n/a), 50147.
6. Farjami, T.; Madadlou, A., Fabrication methods of biopolymeric microgels and
microgel-based hydrogels. Food Hydrocolloids 2017, 62, 262-272.
7. Nooshkam, M.; Varidi, M., Maillard conjugate-based delivery systems for the
encapsulation, protection, and controlled release of nutraceuticals and food bioactive
ingredients: a review. Food Hydrocolloids 2020, 100, 105389.
8. Shang, S.; Liu, J.; He, Y.; Zhu, P., Smart conducting PNIPAM-co-AAc
microgels with controllable phase transition and stimuli responsibility. Mater. Lett.
2020, 127862.
9. Pergushov, D. V.; Sigolaeva, L. V.; Balabushevich, N. G.; Sharifullin, T. Z.;
Noyong, M.; Richtering, W., Loading of doxorubicin into surface-attached stimuli-
1. Introduction
46
responsive microgels and its subsequent release under different conditions. Polymer
2020, 123227.
10. Bergman, M. J.; Pedersen, J. S.; Schurtenberger, P.; Boon, N., Controlling the
morphology of microgels by ionic stimuli. Soft Matter 2020, 16 (11), 2786-2794.
11. Ghanbarinia Firozjah, R.; Sadeghi, A.; Khoee, S., Ultrasonic de-cross-linking
of the pH- and magneto-responsive PHEMA/PMMA microgel to janus nanoparticles:
a new synthesis based on “grafting from”/“grafting to” polymerization. ACS Omega
2020, 5 (42), 27119-27132.
12. Sanzari, I.; Buratti, E.; Huang, R.; Tusan, C. G.; Dinelli, F.; Evans, N. D.;
Prodromakis, T.; Bertoldo, M., Poly(N-isopropylacrylamide) based thin microgel films
for use in cell culture applications. Sci. Rep. 2020, 10 (1), 6126.
13. Varga, I.; Kardos, A.; Borsos, A.; Gilányi, T., Effect of internal charge
distribution on the electrophoretic mobility of poly(N-isopropylacrylamide) based
core-shell microgel particles. J. Mol. Liq. 2020, 302, 111979.
14. Xue, Y.; Chen, Y.; Yu, Y.; Yong, Y., Bacterial nanoencapsulation with
cytocompatible atom transfer radical polymerization for improved Cr(VI) removal.
Chem. Eng. J. 2020, 387, 124068.
15. Zeng, M.; Li, X.; Zhang, Y.; Chen, X.; Sui, X.; Yuan, J., Tailoring the droplet
size of Pickering emulsions by PISA synthesized polymeric nanoparticles. Polymer
2020, 206, 122853.
16. Dieuzy, E.; Aguirre, G.; Auguste, S.; Chougrani, K.; Alard, V.; Billon, L.;
Derail, C., Microstructure-driven self-assembly and rheological properties of multi-
responsive soft microgel suspensions. J. Colloid Interface Sci. 2021, 581, 806-815.
17. Minami, S.; Yamamoto, A.; Oura, S.; Watanabe, T.; Suzuki, D.; Urayama, K.,
Criteria for colloidal gelation of thermo-sensitive poly(N-isopropylacrylamide) based
microgels. J. Colloid Interface Sci. 2020, 568, 165-175.
18. Gavrilov, A. A.; Rudyak, V. Y.; Chertovich, A. V., Computer simulation of the
core-shell microgels synthesis via precipitation polymerization. J. Colloid Interface Sci.
2020, 574, 393-398.
19. Galdioli Pellá, M.; Simão, A.; Lima-Tenório, M.; Tenório-Neto, E. S.; Scariot,
D. B.; Nakamura, C. V.; Rubira, A. F., Chitosan hybrid microgels for oral drug delivery.
Carbohydr. Polym. 2020, 239, 116236.
20. Kim, S. I.; Yim, S.; Chandrasekharan, A.; Seong, K.-Y.; Lee, T. W.; Kim, B.;
Kim, K.; Choi, S.; Yang, S. Y., On-site fabrication of injectable 131I-labeled microgels
for local radiotherapy. J. Controlled Release 2020, 322, 337-345.
21. Tai, Z.; Huang, Y.; Zhu, Q.; Wu, W.; Yi, T.; Chen, Z.; Lu, Y., Utility of
Pickering emulsions in improved oral drug delivery. Drug Discovery Today 2020, 25
(11), 2038-2045.
22. Michel, S. E. S.; Dutertre, F.; Denbow, M. L.; Galan, M. C.; Briscoe, W. H.,
Facile synthesis of chitosan-based hydrogels and microgels through thiol–ene
photoclick cross-linking. ACS Appl. Bio Mater. 2019, 2 (8), 3257-3268.
23. Zhao, J.; He, N., A mini-review of embedded 3D printing: supporting media
and strategies. J. Mater. Chem. B 2020.
24. Jo, Y. K.; Lee, D., Biopolymer microparticles prepared by microfluidics for
biomedical applications. Small 2020, 16 (9).
1. Introduction
47
25. Pellá, M. C. G.; Simão, A. R.; Lima-Tenório, M. K.; Scariot, D. B.; Nakamura,
C. V.; Muniz, E. C.; Rubira, A. F., Magnetic chitosan microgels: Synthesis,
characterization, and evaluation of magnetic field effect over the drug release behavior.
Carbohydr. Polym. 2020, 250, 116879.
26. Volpatti, L. R.; Facklam, A. L.; Cortinas, A. B.; Lu, Y.-C.; Matranga, M. A.;
MacIsaac, C.; Hill, M. C.; Langer, R.; Anderson, D. G., Microgel encapsulated
nanoparticles for glucose-responsive insulin delivery. Biomater. 2021, 267, 120458.
27. Chu, B.; Wu, C.; Tang, S.; Tu, M., Sprayable agarose-derived dopamine-grafted
microgels for promoting tissue adhesion in skin regeneration. React. Funct. Polym.
2020, 154, 104665.
28. Saavedra Isusi, G. I.; Bindereif, B.; Karbstein, H. P.; van der Schaaf, U. S.,
Polymer or microgel particle: Differences in emulsifying properties of pectin as
microgel or as individual polymer chains. Colloids Surf., A 2020, 598, 124793.
29. Heida, T.; Köhler, T.; Kaufmann, A.; Männel, M. J.; Thiele, J., Cell-free protein
synthesis in bifunctional hyaluronan microgels: a strategy for In situ immobilization
and purification of his-tagged proteins. ChemSystemsChem 2020, 2 (3), e1900058.
30. P., F.; O., Ó. F.; M., P.; P., G.; R., F., In vivo drug delivery applications of
nanogels: a review. Nanomed. 2020, 15 (27), 2707-2727.
31. Zhao, S.; Zhou, Y.; Wei, L.; Chen, L., Low fouling strategy of electrochemical
biosensor based on chondroitin sulfate functionalized gold magnetic particle for
voltammetric determination of mycoplasma ovipneumonia in whole serum. Anal. Chim.
Acta 2020, 1126, 91-99.
32. Montheil, T.; Echalier, C.; Martinez, J.; Subra, G.; Mehdi, A., Inorganic
polymerization: an attractive route to biocompatible hybrid hydrogels. J. Mater. Chem.
B 2018, 6 (21), 3434-3448.
33. Fredrikson, J. P.; Brahmachary, P.; Erdoğan, E.; Archambault, Z.; June, R. K.;
Chang, C. B., Metabolomic profiling and mechanotransduction of single chondrocytes
encapsulated in alginate microgels. bioRxiv 2020, 2020.09.28.317008.
34. Feng, R.; Wang, L.; Zhou, P.; Luo, Z.; Li, X.; Gao, L., Development of the pH
responsive chitosan-alginate based microgel for encapsulation of Jughans regia L.
polyphenols under simulated gastrointestinal digestion in vitro. Carbohydr. Polym.
2020, 250, 116917.
35. Oh, J. K.; Lee, D. I.; Park, J. M., Biopolymer-based microgels/nanogels for drug
delivery applications. Prog. Polym. Sci. 2009, 34 (12), 1261-1282.
36. Malerba, M.; Cerana, R., Chitin- and chitosan-based derivatives in plant
protection against biotic and abiotic stresses and in recovery of contaminated soil and
water. Polysaccharides 2020, 1 (1), 21-30.
37. Seyam, S.; Nordin, N. A.; Alfatama, M., Recent progress of chitosan and
chitosan derivatives-based nanoparticles: pharmaceutical perspectives of oral insulin
delivery. Pharmaceuticals 2020, 13 (10), 307.
38. Song, X.; Chen, Y.; Zhao, G.; Sun, H.; Che, H.; Leng, X., Effect of molecular
weight of chitosan and its oligosaccharides on antitumor activities of chitosan-selenium
nanoparticles. Carbohydr. Polym. 2020, 231, 115689.
39. Rogelio, R.-R.; Hugo, E.-A.; Cristina, V.-M.; Zaira Yunuen, G.-C., Composite
hydrogels based on gelatin, chitosan and polyvinyl alcohol to biomedical applications:
a review. Int. J. Polym. Mater. Polym. Biomater. 2020, 69 (1), 1-20.
1. Introduction
48
40. S., U., Chitin, characteristic, sources, and biomedical application. Curr. Pharm.
Biotechnol. 2020, 21 (14), 1433-1443.
41. Loganathan, P.; Gradzielski, M.; Bustamante, H.; Vigneswaran, S., Progress,
challenges, and opportunities in enhancing NOM flocculation using chemically
modified chitosan: a review towards future development. Environ. Sci.: Water Res.
Technol. 2020, 6 (1), 45-61.
42. Mu, M.; Li, X.; Tong, A.; Guo, G., Multi-functional chitosan-based smart
hydrogels mediated biomedical application. Expert Opin. Drug Delivery 2019, 16 (3),
239-250.
43. Azarova, Y. A.; Pestov, A. V.; Bratskaya, S. Y., Application of chitosan and its
derivatives for solid-phase extraction of metal and metalloid ions: a mini-review. Cellul.
2016, 23 (4), 2273-2289.
44. Bazmandeh, A. Z.; Mirzaei, E.; Fadaie, M.; Shirian, S.; Ghasemi, Y., Dual
spinneret electrospun nanofibrous/gel structure of chitosan-gelatin/chitosan-hyaluronic
acid as a wound dressing: In-vitro and in-vivo studies. Int. J. Biol. Macromol. 2020,
162, 359-373.
45. Asadpour, S.; Kargozar, S.; Moradi, L.; Ai, A.; Nosrati, H.; Ai, J., Natural
biomacromolecule based composite scaffolds from silk fibroin, gelatin and chitosan
toward tissue engineering applications. Int. J. Biol. Macromol. 2020, 154, 1285-1294.
46. Rostami, E., Progresses in targeted drug delivery systems using chitosan
nanoparticles in cancer therapy: A mini-review. J. Drug Delivery Sci. Technol. 2020,
58, 101813.
47. Peuler, K.; Dimmitt, N.; Lin, C. C., Clickable modular polysaccharide
nanoparticles for selective cell-targeting. Carbohydr. Polym. 2020, 234, 115901.
48. Wang, Y.; Zhu, L.; Zhang, H.; Huang, H.; Jiang, L., Formulation of pH and
temperature dual-responsive Pickering emulsion stabilized by chitosan-based microgel
for recyclable biocatalysis. Carbohydr. Polym. 2020, 241, 116373.
49. Patil, P. S.; Mansouri, M.; Leipzig, N. D., Fluorinated chitosan microgels to
overcome internal oxygen transport deficiencies in microtissue culture systems. Adv.
Biosyst. 2020, 4 (8), 1900250.
50. (a) Guaresti, O.; Maiz–Fernández, S.; Palomares, T.; Alonso–Varona, A.;
Eceiza, A.; Pérez–Álvarez, L.; Gabilondo, N., Dual charged folate labelled chitosan
nanogels with enhanced mucoadhesion capacity for targeted drug delivery. Eur. Polym.
J. 2020, 134, 109847; (b) Klein, M. P.; Hackenhaar, C. R.; Lorenzoni, A. S. G.;
Rodrigues, R. C.; Costa, T. M. H.; Ninow, J. L.; Hertz, P. F., Chitosan crosslinked with
genipin as support matrix for application in food process: Support characterization and
β-d-galactosidase immobilization. Carbohydr. Polym. 2016, 137, 184-190.
51. Novikov, I. V.; Pigaleva, M. A.; Levin, E. E.; Abramchuk, S. S.; Naumkin, A.
V.; Li, H.; Pich, A.; Gallyamov, M. O., The mechanism of stabilization of silver
nanoparticles by chitosan in carbonic acid solutions. Colloid Polym. Sci. 2020, 298 (9),
1135-1148.
52. Thakur, G.; Rodrigues, F. C.; Dathathri, E.; Nayak, S. K.; Pal, K., 2 - Protein-
based gels: preparation, characterizations, applications in drug delivery, and tissue
engineering. In Polymeric Gels, Pal, K.; Banerjee, I., Eds. Woodhead Publishing: 2018;
pp 31-54.
1. Introduction
49
53. Sun, Y.-S.; Lin, Y.-A.; Huang, H.-H., Using genipin to immobilize bone
morphogenetic protein-2 on zirconia surface for enhancing cell adhesion and
mineralization in dental implant applications. Polymers 2020, 12 (11), 2639.
54. Wang, K.; Lin, S.; Nune, K. C.; Misra, R. D. K., Chitosan-gelatin-based
microgel for sustained drug delivery. J. Biomater. Sci. Polym. Ed. 2016, 27 (5), 441-
453.
55. Mora-Boza, A.; Mancipe Castro, L. M.; Schneider, R. S.; Han, W. M.; García,
A. J.; Vázquez-Lasa, B.; San Román, J., Microfluidics generation of chitosan microgels
containing glycerylphytate crosslinker for in situ human mesenchymal stem cells
encapsulation. Mater. Sci. Eng. C 2020, 111716.
56. Månsson, L. K.; de Wild, T.; Peng, F.; Holm, S. H.; Tegenfeldt, J. O.;
Schurtenberger, P., Preparation of colloidal molecules with temperature-tunable
interactions from oppositely charged microgel spheres. Soft Matter 2019, 15 (42),
8512-8524.
57. Ji, Y.; Lin, X.; Yu, J., Preparation and characterization of oxidized starch-
chitosan complexes for adsorption of procyanidins. LWT 2020, 117, 108610.
58. Vahedifar, A.; Madadlou, A.; Salami, M., Calcium and chitosan-mediated
clustering of whey protein particles for tuning their colloidal stability and flow
behaviour. International Dairy Journal 2017, 73, 136-143.
59. Li, H.; Mergel, O.; Jain, P.; Li, X.; Peng, H.; Rahimi, K.; Singh, S.; Plamper, F.
A.; Pich, A., Electroactive and degradable supramolecular microgels. Soft Matter 2019,
15 (42), 8589-8602.
60. Gu, Z.; Dang, T. T.; Ma, M.; Tang, B. C.; Cheng, H.; Jiang, S.; Dong, Y.; Zhang,
Y.; Anderson, D. G., Glucose-Responsive Microgels Integrated with Enzyme
Nanocapsules for Closed-Loop Insulin Delivery. ACS Nano 2013, 7 (8), 6758-6766.
61. Sethumadhavan, V.; Zuber, K.; Bassell, C.; Teasdale, P. R.; Evans, D.,
Hydrolysis of doped conducting polymers. Commun. Chem. 2020, 3 (1), 153.
62. Hamdy Makhlouf, A. S.; Perez, A.; Guerrero, E., Chapter 13 - Recent trends in
smart polymeric coatings in biomedicine and drug delivery applications. In Advances
in Smart Coatings and Thin Films for Future Industrial and Biomedical Engineering
Applications, Makhlouf, A. S. H.; Abu-Thabit, N. Y., Eds. Elsevier: 2020; pp 359-381.
63. Wu, J.-G.; Chen, J.-H.; Liu, K.-T.; Luo, S.-C., Engineering antifouling
conducting polymers for modern biomedical applications. ACS Appl. Mater. Interfaces
2019, 11 (24), 21294-21307.
64. Su, H.; Liu, H.-Y.; Pappa, A.-M.; Hidalgo, T. C.; Cavassin, P.; Inal, S.; Owens,
R. M.; Daniel, S., Facile generation of biomimetic-supported lipid bilayers on
conducting polymer surfaces for membrane biosensing. ACS Appl. Mater. Interfaces
2019, 11 (47), 43799-43810.
65. Lahiri, A.; Chutia, A.; Carstens, T.; Endres, F., Surface-oxygen induced
electrochemical self-assembly of mesoporous conducting polymers for electrocatalysis.
J. Electrochem. Soc. 2020, 167 (11), 112501.
66. Meng, J.; Wang, X., Microextraction by packed molecularly imprinted polymer
combined ultra-high-performance liquid chromatography for the determination of
levofloxacin in human plasma. J. Chem. 2019, 2019, 4783432.
1. Introduction
50
67. Zhang, L.; Biesheuvel, P. M.; Ryzhkov, I. I., Theory of ion and water transport
in electron-conducting membrane pores with pH-dependent chemical charge. Phys.
Rev. Appl. 2019, 12 (1), 014039.
68. Zhang, H.; Li, Z.; Yu, S.; Xiao, Q.; Lei, G.; Ding, Y., Carbon-encapsulated
LiMn2O4 spheres prepared using a polymer microgel reactor for high-power lithium-
ion batteries. J. Power Sources 2016, 301, 376-385.
69. Yang, Y.; Deng, Z. D., Stretchable sensors for environmental monitoring. Appl.
Phys. Rev. 2019, 6 (1), 011309.
70. Zhang, L.; Wang, B.; Li, X.; Xu, G.; Dou, S.; Zhang, X.; Chen, X.; Zhao, J.;
Zhang, K.; Li, Y., Further understanding of the mechanisms of electrochromic devices
with variable infrared emissivity based on polyaniline conducting polymers. J. Mater.
Chem. C 2019, 7 (32), 9878-9891.
71. Bafekry, A.; Farjami Shayesteh, S.; Peeters, F. M., Introducing novel electronic
and magnetic properties in C3N nanosheets by defect engineering and atom
substitution. Phys. Chem. Chem. Phys. 2019, 21 (37), 21070-21083.
72. Magu, T. O.; Agobi, A. U.; HITLER, L.; Dass, P. M., A review on conducting
polymers-based composites for energy storage application. J. Chem. Rev. 2019, 1 (1),
19-34.
73. Zhang, X.-Y.; Yang, S.; Yang, L.; Zhang, D.; Sun, Y.; Pang, Z.; Yang, J.; Chen,
L., Carrier dynamic monitoring of a π-conjugated polymer: a surface-enhanced Raman
scattering method. Chem. Commun. 2020, 56 (18), 2779-2782.
74. Xu, K.; Sun, H.; Ruoko, T.-P.; Wang, G.; Kroon, R.; Kolhe, N. B.; Puttisong,
Y.; Liu, X.; Fazzi, D.; Shibata, K.; Yang, C.-Y.; Sun, N.; Persson, G.; Yankovich, A.
B.; Olsson, E.; Yoshida, H.; Chen, W. M.; Fahlman, M.; Kemerink, M.; Jenekhe, S. A.;
Müller, C.; Berggren, M.; Fabiano, S., Ground-state electron transfer in all-polymer
donor–acceptor heterojunctions. Nat. Mater. 2020, 19 (7), 738-744.
75. Nowak, M.; Rughooputh, S. D. D. V.; Hotta, S.; Heeger, A. J., Polarons and
bipolarons on a conducting polymer in solution. Macromolecules 1987, 20 (5), 965-
968.
76. Kroschwitz, J. I., Electrical and electronic properties of polymers: a state-of-
the-art compendium. Wiley: New York, 1988.
77. Ivanko, I.; Lindfors, T.; Emanuelsson, R.; Sjödin, M., Conjugated redox
polymer with poly(3,4-ethylenedioxythiophene) backbone and hydroquinone pendant
groups as the solid contact in potassium-selective electrodes. Sens. Actuators, B 2020,
129231.
78. Jin, W.; Wang, R.; Huang, X., Horseradish peroxidase-catalyzed oxidative
polymerization of aniline in bicontinuous microemulsion stabilized by AOT/SDS. J.
Mol. Liq. 2020, 302, 112529.
79. Czichy, M.; Zhylitskaya, H.; Zassowski, P.; Navakouski, M.; Chulkin, P.;
Janasik, P.; Lapkowski, M.; Stępień, M., Electrochemical polymerization of pyrrole–
perimidine hybrids: low-band-gap materials with high n-doping activity. J. Phys. Chem.
C 2020, 124 (26), 14350-14362.
80. Yao, W.; Shen, L.; Liu, P.; Liu, C.; Xu, J.; Jiang, Q.; Liu, G.; Nie, G.; Jiang, F.,
Electrochemical doping engineering tuning of the thermoelectric performance of a π-
conjugated free-standing poly(thiophene-furan) thin-film. Mater. Chem. Front. 2020,
4 (2), 597-604.
1. Introduction
51
81. Inamuddin; Shakeel, N.; Imran Ahamed, M.; Kanchi, S.; Abbas Kashmery, H.,
Green synthesis of ZnO nanoparticles decorated on polyindole functionalized-MCNTs
and used as anode material for enzymatic biofuel cell applications. Sci. Rep. 2020, 10
(1), 5052.
82. Jiang, L.; Rogers, D. M.; Hirst, J. D.; Do, H., Force fields for macromolecular
assemblies containing diketopyrrolopyrrole and thiophene. J. Chem. Theory Comput.
2020, 16 (8), 5150-5162.
83. Das, T.; Verma, B., Polyaniline-acetylene black-copper cobaltite based ternary
hybrid material with enhanced electrochemical properties and its use in supercapacitor
electrodes. Int. J. Energy Res. 2020, 44 (2), 934-949.
84. De Geest, B. G.; Van Camp, W.; Du Prez, F. E.; De Smedt, S. C.; Demeester,
J.; Hennink, W. E., Degradable multilayer films and hollow capsules via a 'Click'
strategy. Macromol. Rapid Commun. 2008, 29 (12-13), 1111-1118.
85. Ansari, S. A.; Ahmed, A.; Ferdousi, F. K.; Salam, M. A.; Shaikh, A. A.; Barai,
H. R.; Lopa, N. S.; Rahman, M. M., Conducting poly(aniline blue)-gold nanoparticles
composite modified fluorine-doped tin oxide electrode for sensitive and non-enzymatic
electrochemical detection of glucose. J. Electroanal. Chem. 2019, 850, 113394.
86. Harris, J. K.; Ratcliff, E. L., Ion diffusion coefficients in poly(3-alkylthiophenes)
for energy conversion and biosensing: role of side-chain length and microstructure. J.
Mater. Chem. C 2020, 8 (38), 13319-13327.
87. Gordillo, M. A.; Benavides, P. A.; Panda, D. K.; Saha, S., The advent of
electrically conducting double-helical metal–organic frameworks featuring butterfly-
shaped electron-rich π-extended tetrathiafulvalene ligands. ACS Appl. Mater.
Interfaces 2020, 12 (11), 12955-12961.
88. Soyleyici, H. C., Electrochromic properties of multifunctional conductive
polymer based on naphthalene. Opt. Mater. 2019, 90, 208-214.
89. Khalili, R.; Zarrintaj, P.; Jafari, S. H.; Vahabi, H.; Saeb, M. R., Electroactive
poly (p-phenylene sulfide)/r-graphene oxide/chitosan as a novel potential candidate for
tissue engineering. Int. J. Biol. Macromol. 2020, 154, 18-24.
90. Wu, B.; Cao, B.; Taylor, I. M.; Woeppel, K.; Cui, X. T., Facile synthesis of a
3,4-ethylene-dioxythiophene (EDOT) derivative for ease of bio-functionalization of
the conducting polymer PEDOT. Front. Chem. 2019, 7 (178).
91. Paula, F. L. d. O.; Castro, L. L. e.; Junior, L. A. R.; Júnior, R. T. d. S.; Silva, G.
M. e.; Neto, P. H. d. O., Dynamical mechanism of polarons and bipolarons in poly(p-
phenylene vinylene). Sci. Rep. 2019, 9 (1), 18131.
92. Tomczykowa, M.; Plonska-Brzezinska, M. E., Conducting polymers, hydrogels
and their composites: preparation, properties and bioapplications. Polymers 2019, 11
(2), 350.
93. Yamamoto, T.; Kimura, T., Preparation of pi-conjugated poly(hydroquinone-
2,5-diyl) and poly(p-benzoquinone-2,5-diyl) and their electrochemical behavior.
Macromolecules 1998, 31 (8), 2683-2685.
94. Han, C.; Li, H.; Shi, R.; Zhang, T.; Tong, J.; Li, J.; Li, B. Q., Organic quinones
towards advanced electrochemical energy storage: recent advances and challenges. J.
Mater. Chem. A 2019, 7 (41), 23378-23415.
95. Bollella, P.; Katz, E., Enzyme-based biosensors: tackling electron transfer
issues. Sensors 2020, 20 (12), 3517.
1. Introduction
52
96. Tisawat, N.; Samart, C.; Jaiyong, P.; Bryce, R. A.; Nueangnoraj, K.; Chanlek,
N.; Kongparakul, S., Enhancement performance of carbon electrode for
supercapacitors by quinone derivatives loading via solvent-free method. Appl. Surf. Sci.
2019, 491, 784-791.
97. Itoi, H.; Tazawa, S.; Hasegawa, H.; Tanabe, Y.; Iwata, H.; Ohzawa, Y., Study
of the pore structure and size effects on the electrochemical capacitor behaviors of
porous carbon/quinone derivative hybrids. RSC Adv. 2019, 9 (47), 27602-27614.
98. Cheng, M.; Yang, X. C.; Zhang, F. G.; Zhao, J. H.; Sun, L. C., Efficient Dye-
Sensitized Solar Cells Based on Hydroquinone/Benzoquinone as a Bioinspired Redox
Couple. Angew. Chem., Int. Ed. 2012, 51 (39), 9896-9899.
99. Siva, T.; Bharathidasan, T.; Sathiyanarayanan, S., Anionic surfactant doped
synthesis of Poly Aniline Dendritic (PANID) fibers and its anti-corrosion performance.
Mater. Today Commun. 2020, 23, 100812.
100. (a) Kanazawa, K. K.; Diaz, A. F.; Geiss, R. H.; Gill, W. D.; Kwak, J. F.; Logan,
J. A.; Rabolt, J. F.; Street, G. B., Organic metals-polypyrrole, a stable synthetic metallic
polymer. J. Chem. Soc. 1979, (19), 854-855; (b) Ghasemi-Mobarakeh, L.; Prabhakaran,
M. P.; Morshed, M.; Nasr-Esfahani, M. H.; Baharvand, H.; Kiani, S.; Al-Deyab, S.;
Ramakrishna, S., Application of conductive polymers, scaffolds and electrical
stimulation for nerve tissue engineering. J. Tissue Eng. Regener. Med. 2011, 5 (4), E17-
E35; (c) Bendrea, A. D.; Cianga, L.; Cianga, I., Review paper: progress in the field of
conducting polymers for tissue engineering applications. J. Biomater. Appl. 2011, 26
(1), 3-84.
101. Liu, Y.; Liu, T. Y.; Pang, L.; Guo, J.; Wang, J.; Qi, D.; Li, W.; Shen, K., Novel
triphenylamine polyamides bearing carbazole and aniline substituents for multi-
colored electrochromic applications. Dyes Pigm. 2020, 173, 107995.
102. Kobayashi, T.; Yoneyama, H.; Tamura, H., Polyaniline film-coated electrodes
as electrochromic display devices. J. Electroanal. Chem. 1984, 161 (2), 419-423.
103. Lin, C. C.; Li, R. L.; Robbennolt, S.; Yeung, M. T.; Akopov, G.; Kaner, R. B.,
Furthering our understanding of the doping mechanism in conjugated polymers using
tetraaniline. Macromolecules 2017, 50 (15), 5892-5897.
104. Ma, Z.; Shi, W.; Yan, K.; Pan, L.; Yu, G., Doping engineering of conductive
polymer hydrogels and their application in advanced sensor technologies. Chem. Sci.
2019, 10 (25), 6232-6244.
105. Goel, M.; Heinrich, C. D.; Krauss, G.; Thelakkat, M., Principles of structural
design of conjugated polymers showing excellent charge transport toward
thermoelectrics and bioelectronics applications. Macromol. Rapid Commun. 2019, 40
(10), 1800915.
106. Jia, H.; Lei, T., Emerging research directions for n-type conjugated polymers.
J. Mater. Chem. C 2019, 7 (41), 12809-12821.
107. Liu, Z.; Wang, Y.; Purro, M.; Xiong, M. P., Oxidation-induced degradable
nanogels for iron chelation. Sci. Rep. 2016, 6 (1), 20923.
108. Lei, B.; Chen, M.; Wang, Y.; Zhang, J.; Xu, S.; Liu, H., Double security drug
delivery system DDS constructed by multi-responsive (pH/redox/US) microgel.
Colloids Surf., B 2020, 193, 111022.
1. Introduction
53
109. Tan, K. H.; Xu, W.; Stefka, S.; Demco, D. E.; Kharandiuk, T.; Ivasiv, V.;
Nebesnyi, R.; Petrovskii, V. S.; Potemkin, I. I.; Pich, A., Selenium-modified microgels
as bio-inspired oxidation catalysts. Angew. Chem., Int. Ed. 2019, 58 (29), 9791-9796.
110. Freeman, K. G.; Adamczyk, J.; Streletzky, K. A., Effect of synthesis
temperature on size, structure, and volume phase transition of polysaccharide microgels.
Macromolecules 2020, 53 (21), 9244-9253.
111. Mutharani, B.; Ranganathan, P.; Chen, S.-M., Temperature-reversible switched
antineoplastic drug 5-fluorouracil electrochemical sensor based on adaptable thermo-
sensitive microgel encapsulated PEDOT. Sens. Actuators, B 2020, 304, 127361.
112. Çalılı, F.; Kaner, P.; Aro, G.; Asatekin, A.; Çulfaz-Emecen, P. Z., Ionic
strength-responsive poly(sulfobetaine methacrylate) microgels for fouling removal
during ultrafiltration. React. Funct. Polym. 2020, 156, 104738.
113. Schmitt, J.; Hartwig, C.; Crassous, J. J.; Mihut, A. M.; Schurtenberger, P.;
Alfredsson, V., Anisotropic mesoporous silica/microgel core–shell responsive particles.
RSC Adv. 2020, 10 (42), 25393-25401.
114. Chen, J.; Ma, X.; Gnanasekar, P.; Qin, D.; Luo, Q.; Sun, Z.; Zhu, J.; Yan, N.,
Synthesis of recoverable thermosensitive Fe3O4 hybrid microgels with controllable
catalytic activity. New J. Chem. 2020, 44 (45), 19440-19444.
115. Das, S. S.; Bharadwaj, P.; Bilal, M.; Barani, M.; Rahdar, A.; Taboada, P.;
Bungau, S.; Kyzas, G. Z., Stimuli-responsive polymeric nanocarriers for drug delivery,
imaging, and theragnosis. Polymers 2020, 12 (6), 1397.
116. Afif, S.; Ghaleh, H.; Nasiri, M.; Maher, B. M.; Abbasi, F., Adhesion,
proliferation, and detachment of cells on poly(N-isopropyl acrylamide) brushes
tethered on polystyrene using surface-initiated atom transfer radical polymerization.
Mater. Today Commun. 2020, 25, 101566.
117. Nishiyama, H.; Odashima, S.; Asoh, S., Femtosecond laser writing of
plasmonic nanoparticles inside PNIPAM microgels for light-driven 3D soft actuators.
Opt. Express 2020, 28 (18), 26470-26480.
118. Benltoufa, S.; Miled, W.; Trad, M.; Slama, R. B.; Fayala, F., Chitosan hydrogel‐
coated cellulosic fabric for medical end-use: Antibacterial properties, basic mechanical
and comfort properties. Carbohydr. Polym. 2020, 227, 115352.
119. Lee, I.; Park, M.; Kim, Y.; Hwang, O.; Khang, G.; Lee, D., Ketal containing
amphiphilic block copolymer micelles as pH-sensitive drug carriers. Int. J. Pharm.
2013, 448 (1), 259-266.
120. Gui, W.; Wang, W.; Jiao, X.; Chen, L.; Wen, Y.; Zhang, X., Dual-cargo
selectively controlled release based on a pH-responsive mesoporous silica system.
ChemPhysChem 2015, 16 (3), 607-613.
121. Ghaz-Jahanian, M. A.; Abbaspour-Aghdam, F.; Anarjan, N.; Berenjian, A.;
Jafarizadeh-Malmiri, H., Application of Chitosan-Based Nanocarriers in Tumor-
Targeted Drug Delivery. Molecular Biotechnology 2015, 57 (3), 201-218.
122. Vishnu S. K, D.; Ranganathan, P.; Rwei, S.-P.; Pattamaprom, C.; Kavitha, T.;
Sarojini, P., New reductant-free synthesis of gold nanoparticles-doped chitosan-based
semi-IPN nanogel: A robust nanoreactor for exclusively sensitive 5-fluorouracil sensor.
Int. J. Biol. Macromol. 2020, 148, 79-88.
1. Introduction
54
123. Khodeir, M.; Antoun, S.; van Ruymbeke, E.; Gohy, J.-F., Temperature and
redox-responsive hydrogels based on nitroxide radicals and oligoethyleneglycol
methacrylate. Macromol. Chem. Phys. 2020, 221 (6), 1900550.
124. Yuan, L.; Wan, C.; Ye, X.; Wu, F., Facial synthesis of silver-incorporated
conductive polypyrrole submicron spheres for supercapacitors. Electrochim. Acta 2016,
213, 115-123.
125. Lai, Y. Y.; Li, X.; Zhu, Y., Polymeric active materials for redox flow battery
application. ACS Appl. Polym. Mater. 2020, 2 (2), 113-128.
126. Augustine, S.; Singh, J.; Srivastava, M.; Sharma, M.; Das, A.; Malhotra, B. D.,
Recent advances in carbon based nanosystems for cancer theranostics. Biomater. Sci.
2017, 5 (5), 901-952.
127. Muhamad, N.; Plengsuriyakarn, T.; Na-Bangchang, K., Application of active
targeting nanoparticle delivery system for chemotherapeutic drugs and
traditional/herbal medicines in cancer therapy: a systematic review. Int J Nanomedicine
2018, 13, 3921-3935.
128. Liyanage, P. Y.; Hettiarachchi, S. D.; Zhou, Y.; Ouhtit, A.; Seven, E. S.; Oztan,
C. Y.; Celik, E.; Leblanc, R. M., Nanoparticle-mediated targeted drug delivery for
breast cancer treatment. Biochim. Biophys. Acta, Rev. Cancer. 2019, 1871 (2), 419-433.
129. Chen, B.; Dai, W.; He, B.; Zhang, H.; Wang, X.; Wang, Y.; Zhang, Q., Current
multistage drug delivery systems based on the tumor microenvironment. Theranostics
2017, 7 (3), 538-558.
130. Shang, S.; Liu, J.; He, Y.; Zhu, P., Smart conducting PNIPAM-co-AAc
microgels with controllable phase transition and stimuli responsibility. Mater. Lett.
2020, 272, 127862.
131. García, M. C., 14 - Ionic-strength-responsive polymers for drug delivery
applications. In Stimuli Responsive Polymeric Nanocarriers for Drug Delivery
Applications, Makhlouf, A. S. H.; Abu-Thabit, N. Y., Eds. Woodhead Publishing: 2019;
pp 393-409.
132. Boissenot, T.; Bordat, A.; Fattal, E.; Tsapis, N., Ultrasound-triggered drug
delivery for cancer treatment using drug delivery systems: From theoretical
considerations to practical applications. J. Controlled Release 2016, 241, 144-163.
133. Bi, H.; Han, X., Magnetic field triggered drug release from lipid microcapsule
containing lipid-coated magnetic nanoparticles. Chem. Phys. Lett. 2018, 706, 455-460.
134. Szunerits, S.; Teodorescu, F.; Boukherroub, R., Electrochemically triggered
release of drugs. Eur. Polym. J. 2016, 83, 467-477.
135. Wu, S. Y.; Chou, H. Y.; Yuh, C. H.; Mekuria, S. L.; Kao, Y. C.; Tsai, H. C.,
Radiation-sensitive dendrimer-based drug delivery system. Adv. Sci. 2018, 5 (2),
1700339.
136. Hu, R.; Zheng, H.; Cao, J.; Davoudi, Z.; Wang, Q., Synthesis and In Vitro
Characterization of Carboxymethyl Chitosan-CBA-Doxorubicin Conjugate
Nanoparticles as pH-Sensitive Drug Delivery Systems. Journal of Biomedical
Nanotechnology 2017, 13 (9), 1097-1105.
137. Hao, Y.; Zheng, C.; Wang, L.; Hu, Y.; Guo, H.; Song, Q.; Zhang, H.; Zhang,
Z.; Zhang, Y., Covalent self-assembled nanoparticles with pH-dependent enhanced
tumor retention and drug release for improving tumor therapeutic efficiency. J. Mater.
Chem. B 2017, 5 (11), 2133-2144.
1. Introduction
55
138. Laporte, A.; Richard, H.; Bonnaud, E.; Henry, P.; Vital, A.; Georgescauld, D.,
A spin label study of myelin fluidity with normal and pathological peripheral nerves.
J. Neurol. Sci. 1979, 43 (3), 345-356.
139. Ait Bachir, Z.; Huang, Y.; He, M.; Huang, L.; Hou, X.; Chen, R.; Gao, F.,
Effects of PEG surface density and chain length on the pharmacokinetics and
biodistribution of methotrexate-loaded chitosan nanoparticles. J. Mater. Chem. B 2018,
13, 5657-5671.
140. Wang, J.; Asghar, S.; Yang, L.; Gao, S.; Chen, Z.; Huang, L.; Zong, L.; Ping,
Q.; Xiao, Y., Chitosan hydrochloride/hyaluronic acid nanoparticles coated by mPEG
as long-circulating nanocarriers for systemic delivery of mitoxantrone. Int. J. Biol.
Macromol. 2018, 113, 345-353.
141. Hayward, S. L.; Wilson, C. L.; Kidambi, S., Hyaluronic acid-conjugated
liposome nanoparticles for targeted delivery to CD44 overexpressing glioblastoma
cells. Oncotarget 2016, 7 (23), 34158-34171.
142. Paul, R.; Dutta, D.; Paul, R.; Dash, J., Target-directed azide-alkyne
cycloaddition for assembling HIV-1 TAR RNA binding ligands. Angew. Chem., Int.
Ed. 2020, 59 (30), 12407-12411.
143. Mi, P.; Cabral, H.; Kataoka, K., Ligand-installed nanocarriers toward precision
therapy. Adv. Mater. 2020, 32 (13), 1902604.
144. C., F.; H., G.; H., H., Sugar ligand-mediated drug delivery. Future Med. Chem.
2020, 12 (2), 161-171.
145. Yu, F.; Zhu, M.; Li, N.; Ao, M.; Li, Y.; Zhong, M.; Yuan, Q.; Chen, H.; Fan,
Z.; Wang, Y.; Hou, Z.; Qi, Z.; Shen, Y.; Chen, X., Imaging-guided synergistic
targeting-promoted photo-chemotherapy against cancers by methotrexate-conjugated
hyaluronic acid nanoparticles. Chem. Eng. J. 2020, 380, 122426.
146. Min, L.; Yan, W.; Shuai, J.; Yang, G.; Weijie, Z.; Shaobo, H.; Xiang, C.; Chen,
Z.; Ping, S.; Wenbo, K.; Guoliang, W.; Zifang, S.; Yong, Z.; Z., Q. C., Biodistribution
and biocompatibility of glycyrrhetinic acid and galactose-modified chitosan
nanoparticles as a novel targeting vehicle for hepatocellular carcinoma. Nanomedicine
2020, 15 (2), 145-161.
147. Hefnawy, A.; Khalil, I. H.; Arafa, K.; Emara, M.; El-Sherbiny, I. M., Dual-
ligand functionalized core-shell chitosan-based nanocarrier for hepatocellular
carcinoma-targeted drug delivery. Int J Nanomedicine 2020, 15, 821-837.
148. Hua, S.; Yu, J.; Shang, J.; Zhang, H.; Du, J.; Zhang, Y.; Chen, F.; Zhou, Y.; Liu,
F., Effective tumor-targeted delivery of etoposide using chitosan nanoparticles
conjugated with folic acid and sulfobetaine methacrylate. RSC Adv. 2016, 6 (94),
91192-91200.
149. Hefnawy, A.; Khalil, I. H.; Arafa, K.; Emara, M.; El-Sherbiny, I. M., Dual-
Ligand Functionalized Core-Shell Chitosan-Based Nanocarrier for Hepatocellular
Carcinoma-Targeted Drug Delivery. International journal of nanomedicine 2020, 15,
821-837.
150. Moiz, A.; Padhye, R.; Wang, X., A comparative study on the effect of surface
functionalization on the versatile protection of textiles. Fibers Polym. 2019, 20 (2),
348-357.
151. Štular, D.; Jerman, I.; Naglič, I.; Simončič, B.; Tomšič, B., Embedment of silver
into temperature- and pH-responsive microgel for the development of smart textiles
1. Introduction
56
with simultaneous moisture management and controlled antimicrobial activities.
Carbohydr. Polym. 2017, 159, 161-170.
152. Kulkarni, A.; Tourrette, A.; Warmoeskerken, M. M. C. G.; Jocic, D., Microgel-
based surface modifying system for stimuli-responsive functional finishing of cotton.
Carbohydr. Polym. 2010, 82 (4), 1306-1314.
153. Mattheis, C.; Zhang, Y.; Agarwal, S., Thermo-switchable antibacterial activity.
Macromol. Biosci. 2012, 12 (10), 1401-1412.
154. Hu, J. L.; Meng, H. P.; Li, G. Q.; Ibekwe, S. I., A review of stimuli-responsive
polymers for smart textile applications. Smart Mater. Struct. 2012, 21 (5).
155. Štular, D.; Jerman, I.; Simončič, B.; Tomšič, B., Tailoring of temperature- and
pH-responsive cotton fabric with antimicrobial activity: Effect of the concentration of
a bio-barrier-forming agent. Carbohydr. Polym. 2017, 174, 677-687.
156. Verbic, A.; Stojkoski, V.; Tomsic, B.; Spicka, N.; Stular, D.; Gorjanc, M.; Kert,
M.; Simoncic, B., Preparation of Functional Stimuli-responsive Polyamide 6 Fabric
with ZnO Incorporated Microgel. Tekstilec 2018, 61 (1), 15-26.
157. Chen, Y.; Zheng, Z.; Zhou, R.; Zhang, H.; Chen, C.; Xiong, Z.; Liu, K.; Wang,
X., Developing a strontium-releasing graphene oxide-/collagen-based organic–
inorganic nanobiocomposite for large bone defect regeneration via MAPK signaling
pathway. ACS Appl. Mater. Interfaces 2019, 11 (17), 15986-15997.
158. Kawano, T.; Murata, M.; Kang, J.-H.; Piao, J. S.; Narahara, S.; Hyodo, F. i.;
Hamano, N.; Guo, J.; Oguri, S.; Ohuchida, K.; Hashizume, M., Ultrasensitive MRI
detection of spontaneous pancreatic tumors with nanocage-based targeted contrast
agent. Biomaterials 2018, 152, 37-46.
159. Chu, Z.; Wang, Z.; Chen, L.; Wang, X.; Huang, C.; Cui, M.; Yang, D.; Jia, N.,
Combining magnetic resonance imaging with photothermal therapy of CuS@BSA
nanoparticles for cancer theranostics. ACS Appl. Nano Mater. 2018, 1 (5), 2332-2340.
160. Barandov, A.; Bartelle, B. B.; Williamson, C. G.; Loucks, E. S.; Lippard, S. J.;
Jasanoff, A., Sensing intracellular calcium ions using a manganese-based MRI contrast
agent. Nat. Commun. 2019, 10 (1), 897.
161. Waddington, D. E. J.; Boele, T.; Maschmeyer, R.; Kuncic, Z.; Rosen, M. S.,
High-sensitivity in vivo contrast for ultra-low field magnetic resonance imaging using
superparamagnetic iron oxide nanoparticles. Sci. Adv. 2020, 6 (29), eabb0998.
162. Hong, Y.; Han, Y.; Wu, J.; Zhao, X.; Cheng, J.; Gao, G.; Qian, Q.; Wang, X.;
Cai, W.; Zreiqat, H.; Feng, D.; Xu, J.; Cui, D., Chitosan modified Fe3O4/KGN self-
assembled nanoprobes for osteochondral MR diagnose and regeneration. Theranostics
2020, 10 (12), 5565-5577.
163. Wang, C. Y.; Johnson, J. A.; Zhang, Q.; Beahm, E. K., Combining
decellularized human adipose tissue extracellular matrix and adipose-derived stem
cells for adipose tissue engineering. Acta Biomater. 2013, 9 (11), 8921-8931.
164. Poon, C. J.; Pereira E. Cotta, C. V.; Sinha, S.; Palmer, J. A.; Woods, A. A.;
Morrison, W. A.; Abberton, K. M., Preparation of an adipogenic hydrogel from
subcutaneous adipose tissue. Acta Biomater. 2013, 9 (3), 5609-5620.
165. Xia, P.; Zhang, K.; Gong, Y.; Li, G.; Yan, S.; Yin, J., Injectable stem cell laden
open porous microgels that favor adipogenesis: in vitro and in vivo evaluation. ACS
Appl. Mater. Interfaces 2017, 9 (40), 34751-34761.
1. Introduction
57
166. Prasannan, A.; Udomsin, J.; Tsai, H.-C.; Sivakumar, M.; Hu, C.-C.; Wang, C.-
F.; Hung, W.-S.; Lai, J.-Y., Special wettable underwater superoleophobic material for
effective simultaneous removal of high viscous insoluble oils and soluble dyes from
wastewater. J. Membr. Sci. 2020, 603, 118026.
167. Oyewo, O. A.; Elemike, E. E.; Onwudiwe, D. C.; Onyango, M. S., Metal oxide-
cellulose nanocomposites for the removal of toxic metals and dyes from wastewater.
Int. J. Biol. Macromol. 2020, 164, 2477-2496.
168. Rezakazemi, M.; Khajeh, A.; Mesbah, M., Membrane filtration of wastewater
from gas and oil production. Environ. Chem. Lett. 2018, 16 (2), 367-388.
169. Villaseñor, M. J.; Ríos, Á., Nanomaterials for water cleaning and desalination,
energy production, disinfection, agriculture and green chemistry. Environ. Chem. Lett.
2018, 16 (1), 11-34.
170. Vatanpour, V.; Salehi, E.; Sahebjamee, N.; Ashrafi, M., Novel
chitosan/polyvinyl alcohol thin membrane adsorbents modified with detonation
nanodiamonds: Preparation, characterization, and adsorption performance. Arabian J.
Chem. 2020, 13 (1), 1731-1740.
171. Du, W.; Niu, S.; Xu, Y.; Xu, Z.; Fan, C., Antibacterial activity of chitosan
tripolyphosphate nanoparticles loaded with various metal ions. Carbohydr. Polym.
2009, 75 (3), 385-389.
172. Kolangare, I. M.; Isloor, A. M.; Karim, Z. A.; Kulal, A.; Ismail, A. F.;
Inamuddin; Asiri, A. M., Antibiofouling hollow-fiber membranes for dye rejection by
embedding chitosan and silver-loaded chitosan nanoparticles. Environ. Chem. Lett.
2019, 17 (1), 581-587.
173. Ramalingam, B.; Khan, M. M. R.; Mondal, B.; Mandal, A. B.; Das, S. K., Facile
synthesis of silver nanoparticles decorated magnetic-chitosan microsphere for efficient
removal of dyes and microbial contaminants. ACS Sustainable Chem. Eng. 2015, 3 (9),
2291-2302.
174. Yao, X.; Chen, L.; Chen, X.; Zhang, Z.; Zheng, H.; He, C.; Zhang, J.; Chen, X.,
Intracellular pH-sensitive metallo-supramolecular nanogels for anticancer drug
delivery. ACS Appl. Mater. Interfaces 2014, 6 (10), 7816-7822.
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.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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,
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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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)
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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)
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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 Degradation of Microgels
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.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
86
Scheme 4. (A) Schematic representation of preparation of chitosan-hydroquinone
(CHHQ), (B) chitosan-hydroquinone-glutaraldehyde (CHHQGA), and (C) chitosan-
glutaraldehyde (CHGA) microgels.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan 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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
88
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).
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
91
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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
93
2.3.7 Drug Loading and Release Studies
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.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
94
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,
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
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.
2.5 References and Notes
1. Pelton, R. H.; Chibante, P., Preparation of aqueous latices with N-
isopropylacrylamide. Colloids Surf., A 1986, 20 (3), 247-256.
2. Huang, X.; Luo, Z.; Guo, J.; Ruan, Q.; Wang, J.; Yang, X., Enzyme-adsorbed
chitosan nanogel particles as edible pickering interfacial biocatalysts and lipase-
responsive phase inversion of emulsions. J. Agric. Food Chem. 2020, 68 (33), 8890-
8899.
3. Inger, E.; Sunol, A. K.; Sahiner, N., Catalytic activity of metal-free amine-
modified dextran microgels in hydrogen release through methanolysis of NaBH4. Int.
J. Energy Res. 2020, 44 (7), 5990-6001.
4. Suraiya, A. B.; Hun, M. L.; Truong, V. X.; Forsythe, J. S.; Chidgey, A. P.,
Gelatin-based 3D microgels for in vitro T lineage cell generation. ACS Biomater. Sci.
Eng. 2020, 6 (4), 2198-2208.
5. Youn, H. G.; Je, J. Y.; Lee, C. M.; Yoon, S. D., Inulin/PVA biomaterials using
thiamine as an alternative plasticizer. Carbohydr. Polym. 2019, 220, 86-94.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
97
6. Ji, Y., Microgels prepared from corn starch with an improved capacity for
uptake and release of lysozyme. J. Food Eng. 2020, 285, 110088.
7. An, C.; Liu, W.; Zhang, Y.; Pang, B.; Liu, H.; Zhang, Y.; Zhang, H.; Zhang, L.;
Liao, H.; Ren, C.; Wang, H., Continuous microfluidic encapsulation of single
mesenchymal stem cells using alginate microgels as injectable fillers for bone
regeneration. Acta Biomater. 2020, 111, 181-196.
8. Shende, P.; Pathan, N., Potential of carbohydrate-conjugated graphene
assemblies in biomedical applications. Carbohydr. Polym. 2020, 117385.
9. Vilela, C.; Figueiredo, A. R. P.; Silvestre, A. J. D.; Freire, C. S. R., Multilayered
materials based on biopolymers as drug delivery systems. Expert Opin. Drug Delivery
2017, 14 (2), 189-200.
10. Lemke, P.; Moerschbacher, B. M.; Singh, R., Transcriptome analysis of
solanum tuberosum genotype RH89-039-16 in response to chitosan. Front. Plant Sci.
2020, 11 (1193).
11. Tian, B. R.; Liu, Y. M.; Liu, J. Y., Smart stimuli-responsive drug delivery
systems based on cyclodextrin: A review. Carbohydr. Polym. 2021, 251.
12. Le, T. H.; Kim, Y.; Yoon, H., Electrical and electrochemical properties of
conducting polymers. Polymers 2017, 9 (4).
13. Imran, S. M.; Salman, A.; Shao, G. N.; Haider, M. S.; Abbas, N.; Park, S. S.;
Hussain, M.; Kim, H. T., Study of the electroconductive properties of conductive
polymers-graphene/graphene oxide nanocomposites synthesized via in situ emulsion
polymerization. Polym. Compos. 2018, 39 (6), 2142-2150.
14. Sun, Y.; Ding, X. J.; Zhang, X. Q.; Huang, Q. Z.; Lin, B. P.; Yang, H.; Guo, L.
X., Indeno 1,2-b fluorene-based novel donor-acceptor conjugated copolymers:
Electron acceptor engineering and polymer backbone planarization. High Perform.
Polym. 2018, 30 (2), 192-201.
15. Yuan, M.; Minteer, S. D., Redox polymers in electrochemical systems: From
methods of mediation to energy storage. Curr. Opin. Electrochem. 2019, 15, 1-6.
16. Liu, Y.; McGrath, J. S.; Moore, J. H.; Kolling, G. L.; Papin, J. A.; Swami, N.
S., Electrofabricated biomaterial-based capacitor on nanoporous gold for enhanced
redox amplification. Electrochimica Acta 2019, 318, 828-836.
17. Mergel, O.; Schneider, S.; Tiwari, R.; Kuhn, P. T.; Keskin, D.; Stuart, M. C. A.;
Schottner, S.; Kanter, M. d.; Noyong, M.; Caumanns, T.; Mayer, J.; Janzen, C.; Simon,
U.; Gallei, M.; Wöll, D.; van Rijn, P.; Plamper, F. A., Cargo shuttling by
electrochemical switching of core-shell microgels obtained by a facile one-shot
polymerization. Chem. Sci. 2019, 10 (6), 1844-1856.
18. Janoschka, T.; Martin, N.; Martin, U.; Friebe, C.; Morgenstern, S.; Hiller, H.;
Hager, M. D.; Schubert, U. S., An aqueous, polymer-based redox-flow battery using
non-corrosive, safe, and low-cost materials. Nature 2015, 527 (7576), 78-81.
19. (a) Yamamoto, T.; Kimura, T., Preparation of π-conjugated poly(hydroquinone-
2,5-diyl) and poly(p-benzoquinone-2,5-diyl) and their electrochemical behavior.
Macromolecules 1998, 31 (8), 2683-2685; (b) Yamamoto, T.; Kimura, T.; Shiraishi,
K., Preparation of π-conjugated polymers composed of hydroquinone, p-benzoquinone,
and p-diacetoxyphenylene units. Macromolecules 1999, 32 (26), 8886-8896.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
98
20. He, J.; Zhang, A.; Zhang, Y.; Guan, Y., Novel redox hydrogel by in situ gelation
of chitosan as a result of template oxidative polymerization of hydroquinone.
Macromolecules 2011, 44 (7), 2245-2252.
21. Allan, G. G.; Peyron, M., Molecular weight manipulation of chitosan I: kinetics
of depolymerization by nitrous acid. Carbohydr. Res. 1995, 277 (2), 257-272.
22. Qin, W.; Wang, Y.; Fang, G.; Wu, T.; Liu, C.; Zhou, D., Evidence for the
generation of reactive oxygen species from hydroquinone and benzoquinone: Roles in
arsenite oxidation. Chemosphere 2016, 150, 71-78.
23. Butwong, N.; Srijaranai, S.; Glennon, J. D.; Luong, J. H. T., Cysteamine capped
silver nanoparticles and single-walled carbon nanotubes composite coated on glassy
carbon electrode for simultaneous analysis of hydroquinone and catechol.
Electroanalysis 2018, 30 (5), 962-968.
24. Liu, W.; Li, X.; Wang, S.; Fang, F.; Wang, X.; Hou, Q., Nanocomposites
derived from licorice residues cellulose nanofibril and chitosan nanofibril: Effects of
chitosan nanofibril dosage on resultant properties. Int. J. Biol. Macromol. 2020, 165,
2404-2411.
25. Mania, S.; Ryl, J.; Jinn, J.-R.; Wang, Y.-J.; Michałowska, A.; Tylingo, R., The
production possibility of the antimicrobial filaments by co-extrusion of the PLA pellet
with chitosan powder for FDM 3D printing technology. Polymers 2019, 11 (11), 1893.
26. Acemi, A., Polymerization degree of chitosan affects structural and
compositional changes in the cell walls, membrane lipids, and proteins in the leaves of
Ipomoea purpurea: An FT-IR spectroscopy study. Int. J. Biol. Macromol. 2020, 162,
715-722.
27. Chen, S. Y.; Han, Y.; Jian, L.; Liao, W.; Zhang, Y.; Gao, Y., Fabrication,
characterization, physicochemical stability of zein-chitosan nanocomplex for co-
encapsulating curcumin and resveratrol. Carbohydr. Polym. 2020, 236, 116090.
28. Mohanadas, D.; Tukimin, N.; Sulaiman, Y., Simultaneous electrochemical
detection of hydroquinone and catechol using poly(3,4-
ethylenedioxythiophene)/reduced graphene oxide/manganese dioxide. Synth. Met.
2019, 252, 76-81.
29. Minisy, I. M.; Salahuddin, N. A.; Ayad, M. M., In vitro release study of
ketoprofen-loaded chitosan/polyaniline nanofibers. Polym. Bull. 2020.
30. Zhang, C.-W.; Li, F.-Y.; Li, J.-F.; Li, Y.-L.; Xu, J.; Xie, Q.; Chen, S. Y.; Guo,
A., Novel treatments for compatibility of plant fiber and starch by forming new
hydrogen bonds. J. Cleaner Prod. 2018, 185, 357-365.
31. Trammell, S. A.; Lowy, D. A.; Seferos, D. S.; Moore, M.; Bazan, G. C.;
Lebedev, N., Heterogeneous electron transfer of quinone-hydroquinone in alkaline
solutions at gold electrode surfaces: Comparison of saturated and unsaturated bridges.
J. Electroanal. Chem. 2007, 606 (1), 33-38.
32. Wild, U.; Hübner, O.; Himmel, H.-J., Redox-active guanidines in proton-
coupled electron-transfer reactions: real alternatives to benzoquinones? Chem. Eur. J.
2019, 25 (70), 15988-15992.
33. Kiss, V.; Lehoczki, G.; Ősz, K., Mathematical description of pH-stat kinetic
traces measured during photochemical quinone decomposition. Photochem. Photobiol.
Sci. 2017, 16 (4), 519-526.
2. Redox-Active Supramolecular Poly(hydroquinone)-Chitosan Microgels
99
34. Bourbonnais, R.; Leech, D.; Paice, M. G., Electrochemical analysis of the
interactions of laccase mediators with lignin model compounds. Biochim. Biophys.
Acta, Gen. Subj. 1998, 1379 (3), 381-390.
35. Rupley, J. A., The effect of urea and amides upon water structure. J. Phys. Chem.
1964, 68 (7), 2002-2003.
36. Gerken, J. B.; Stamoulis, A.; Suh, S.-E.; Fischer, N. D.; Kim, Y. J.; Guzei, I.
A.; Stahl, S. S., Efficient electrochemical synthesis of robust, densely functionalized
water soluble quinones. Chem. Commun. 2020, 56 (8), 1199-1202.
37. Roman, D. L.; Ostafe, V.; Isvoran, A., Deeper inside the specificity of lysozyme
when degrading chitosan. A structural bioinformatics study. J. Mol. Graphics Modell.
2020, 100, 107676.
38. Kim, S.; Fan, J.; Lee, C.-S.; Lee, M., Dual functional lysozyme–chitosan
conjugate for tunable degradation and antibacterial activity. ACS Applied Bio Materials
2020, 3 (4), 2334-2343.
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3. Polyaniline-Chitosan Microgels
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3. Polyaniline-Chitosan Microgels
3. Polyaniline-Chitosan Microgels
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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
3. Polyaniline-Chitosan Microgels
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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
3. Polyaniline-Chitosan Microgels
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
3. Polyaniline-Chitosan Microgels
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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
3. Polyaniline-Chitosan Microgels
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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
3. Polyaniline-Chitosan Microgels
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 Experimental Section
3.2.1 Materials
All reagents were purchased commercially and used without further
purification except chitosan. Chitosan of medium molecular weight (75-85%
3. Polyaniline-Chitosan Microgels
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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
3. Polyaniline-Chitosan Microgels
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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
3. Polyaniline-Chitosan Microgels
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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
3. Polyaniline-Chitosan Microgels
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
3. Polyaniline-Chitosan Microgels
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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 Results and Discussion
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
3. Polyaniline-Chitosan Microgels
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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.
3. Polyaniline-Chitosan Microgels
114
Scheme 2. Mechanism of polyaniline polymerization49.
3. Polyaniline-Chitosan Microgels
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
3. Polyaniline-Chitosan Microgels
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.
3. Polyaniline-Chitosan Microgels
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.
3. Polyaniline-Chitosan Microgels
118
Fig. 2. FTIR spectra of chitosan and the CH-PANI microgels with different grafting
ratio.
3.3.3 Influence of pH on Microgel Size and Electrophoretic Mobility
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.
3. Polyaniline-Chitosan Microgels
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.
3. Polyaniline-Chitosan Microgels
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.
3. Polyaniline-Chitosan Microgels
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
3. Polyaniline-Chitosan Microgels
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.
3.3.5 Degradation of Microgels
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.
3. Polyaniline-Chitosan Microgels
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.
3. Polyaniline-Chitosan Microgels
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
3. Polyaniline-Chitosan Microgels
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.
3.5 References and Notes
1. Xu, Y.; Ma, Y.; Ji, X.; Huang, S.; Xia, J.; Xie, M.; Yan, J.; Xu, H.; Li, H.,
Conjugated conducting polymers PANI decorated Bi12O17Cl2 photocatalyst with
extended light response range and enhanced photoactivity. Appl. Surf. Sci. 2019, 464,
552-561.
2. Ashizawa, M.; Zheng, Y.; Tran, H.; Bao, Z., Intrinsically stretchable conjugated
polymer semiconductors in field effect transistors. Prog. Polym. Sci. 2020, 100, 101181.
3. Moon, J. M.; Thapliyal, N.; Hussain, K. K.; Goyal, R. N.; Shim, Y. B.,
Conducting polymer-based electrochemical biosensors for neurotransmitters: A review.
Biosens. Bioelectron. 2018, 102, 540-552.
4. Qu, H.; Duan, X., Recent advances in micro detectors for micro gas
chromatography. Sci. China Mater. 2019, 62 (5), 611-623.
5. B. Aziz, S.; H. Hamsan, M.; M. Nofal, M.; San, S.; Abdulwahid, R. T.; Raza
Saeed, S.; Brza, M. A.; Kadir, M. F. Z.; Mohammed, S. J.; Al-Zangana, S., From
cellulose, shrimp and crab shells to energy storage EDLC cells: the study of structural
and electrochemical properties of proton conducting chitosan-based biopolymer blend
electrolytes. Polymers 2020, 12 (7), 1526.
6. Asnawi, A. S. F. M.; B. Aziz, S.; M. Nofal, M.; Hamsan, M. H.; Brza, M. A.;
Yusof, Y. M.; Abdilwahid, R. T.; Muzakir, S. K.; Kadir, M. F. Z., Glycerolized Li+
ion conducting chitosan-based polymer electrolyte for energy storage EDLC device
applications with relatively high energy density. Polymers 2020, 12 (6), 1433.
7. Shifrina, Z. B.; Matveeva, V. G.; Bronstein, L. M., Role of polymer structures
in catalysis by transition metal and metal oxide nanoparticle composites. Chem. Rev.
2020, 120 (2), 1350-1396.
3. Polyaniline-Chitosan Microgels
126
8. Tao, H.; Liu, S.; Luo, J. L.; Choi, P.; Liu, Q.; Xu, Z., Descriptor of catalytic
activity of metal sulfides for oxygen reduction reaction: a potential indicator for
mineral flotation. J. Mater. Chem. A 2018, 6 (20), 9650-9656.
9. Puiggalí-Jou, A.; del Valle, L. J.; Alemán, C., Drug delivery systems based on
intrinsically conducting polymers. J. Controlled Release 2019, 309, 244-264.
10. Zhang, Y.; Xu, P.; Zeng, Q.; Liu, Y.; Liao, X.; Hou, M., Magnetism-assisted
modification of screen printed electrode with magnetic multi-walled carbon nanotubes
for electrochemical determination of dopamine. Mater. Sci. Eng. C 2017, 74, 62-69.
11. Zhao, M.; Song, B.; Pu, J.; Wada, T.; Reid, B.; Tai, G. P.; Wang, F.; Guo, A.;
Walczysko, P.; Gu, Y.; Sasaki, T.; Suzuki, A.; Forrester, J. V.; Bourne, H. R.;
Devreotes, P. N.; McCaig, C. D.; Penninger, J. M., Electrical signals control wound
healing through phosphatidylinositol-3-OH kinase-[gamma] and PTEN. Nature 2006,
442 (7101), 457-460.
12. Shahini, A.; Yazdimamaghani, M.; Walker, K. J.; Eastman, M. A.; Hatami-
Marbini, H.; Smith, B. J.; Ricci, J. L.; Madihally, S. V.; Vashaee, D.; Tayebi, L., 3D
conductive nanocomposite scaffold for bone tissue engineering. Int J Nanomedicine
2014, 9, 167-181.
13. Liu, Y.; Cui, H.; Zhuang, X. L.; Wei, Y.; Chen, X., Electrospinning of aniline
pentamer-graft-gelatin/PLLA nanofibers for bone tissue engineering. Acta Biomater.
2014, 10 (12), 5074-5080.
14. Senapati, S.; Mahanta, A. K.; Kumar, S.; Maiti, P., Controlled drug delivery
vehicles for cancer treatment and their performance. Signal Transduction Targeted
Ther. 2018, 3 (1), 7.
15. Masdarolomoor, F.; Hajizadeh, S.; Arab Chamjangali, M.; Innis, P. C., Novel
approach to the synthesis of polyaniline possessing electroactivity at neutral pH. Synth.
Met. 2019, 250, 121-130.
16. Wu, D.; Zhong, W., A new strategy for anchoring a functionalized graphene
hydrogel in a carbon cloth network to support a lignosulfonate/polyaniline hydrogel as
an integrated electrode for flexible high areal-capacitance supercapacitors. J. Mater.
Chem. A 2019, 7 (10), 5819-5830.
17. Aliabadi, R. S.; Mahmoodi, N. O., Synthesis and characterization of
polypyrrole, polyaniline nanoparticles and their nanocomposite for removal of azo dyes;
sunset yellow and Congo red. J. Cleaner Prod. 2018, 179, 235-245.
18. Wang, H.; Nie, S.; Li, H.; Ali, R.; Fu, J.; Xiong, H.; Li, J.; Wu, Z.; Lau, W. M.;
Mahmood, N.; Jia, R.; Liu, Y.; Jian, X., 3D hollow quasi-graphite capsules/polyaniline
hybrid with a high performance for room-temperature ammonia gas sensors. ACS Sens.
2019, 4 (9), 2343-2350.
19. Zeng, R.; Luo, Z.; Zhang, L.; Tang, D., Platinum nanozyme-catalyzed gas
generation for pressure-based bioassay using polyaniline nanowires-functionalized
graphene oxide framework. Anal. Chem. 2018, 90 (20), 12299-12306.
20. Kobayashi, T.; Yoneyama, H.; Tamura, H., Polyaniline film-coated electrodes
as electrochromic display devices. J. Electroanal. Chem. 1984, 161 (2), 419-423.
21. (a) Higashimura, H.; Kobayashi, S., Oxidative polymerization. In Encyclopedia
of Polymer Science and Technology, John Wiley & Sons, Inc.: 2002; (b) Gospodinova,
N.; Terlemezyan, L., Conducting polymers prepared by oxidative polymerization:
polyaniline. Prog. Polym. Sci. 1998, 23 (8), 1443-1484.
3. Polyaniline-Chitosan Microgels
127
22. Xiong, S.; Wang, R.; Li, S.; Wu, B.; Chu, J.; Wang, X.; Zhang, R.; Gong, M.,
Electrochromic behaviors of water-Ssoluble polyaniline with covalently bonded acetyl
ferrocene. J. Electron. Mater. 2018, 47 (7), 3974-3982.
23. Pruneanu, S.; Veress, E.; Marian, I.; Oniciu, L., Characterization of polyaniline
by cyclic voltammetry and UV-Vis absorption spectroscopy. J. Mater. Sci. 1999, 34
(11), 2733-2739.
24. Wang, H.; Wen, H.; Hu, B.; Fei, G.; Shen, Y.; Sun, L.; Yang, D., Facile
approach to fabricate waterborne polyaniline nanocomposites with environmental
benignity and high physical properties. Sci. Rep. 2017, 7 (1), 43694.
25. (a) Pud, A. A.; Shapoval, G. S.; Kamarchik, P.; Ogurtsov, N. A.; Gromovaya,
V. F.; Myronyuk, I. E.; Kontsur, Y. V., Electrochemical behavior of mild steel coated
by polyaniline doped with organic sulfonic acids. Synth. Met. 1999, 107 (2), 111-115;
(b) Ayad, M. M.; Salahuddin, N. A.; Minisy, I. M.; Amer, W. A., Chitosan/polyaniline
nanofibers coating on the quartz crystal microbalance electrode for gas sensing. Sens.
Actuators, B 2014, 202, 144-153.
26. Pashaei, S.; Hosseinzadeh, S.; Hosseinzadeh, H., TGA investigation and
morphological properties study of nanocrystalline cellulose/ag-nanoparticles
nanocomposites for catalytic control of oxidative polymerization of aniline. Polym.
Compos. 2019, 40 (S1), E753-E764.
27. Oberhauser, W.; Evangelisti, C.; Tiozzo, C.; Bartoli, M.; Frediani, M.;
Passaglia, E.; Rosi, L., Platinum nanoparticles onto pegylated poly(lactic acid)
stereocomplex for highly selective hydrogenation of aromatic nitrocompounds to
anilines. Appl. Catal., A 2017, 537, 50-58.
28. Ramya, E.; Rajashree, C.; Nayak, P. L.; Narayana Rao, D., New hybrid organic
polymer montmorillonite/chitosan/polyphenylenediamine composites for nonlinear
optical studies. Appl. Clay Sci. 2017, 150, 323-332.
29. Utiye, A. S.; Awasthi, S. K.; Bajpai, S.; Mishra, B. In Electrical and mechanical
characterization of gelatin/poly (aniline) composite films, Adv Mater Proc, 2017; pp
337-341.
30. Atoufi, Z.; Zarrintaj, P.; Motlagh, G. H.; Amiri, A.; Bagher, Z.; Kamrava, S. K.,
A novel bio electro active alginate-aniline tetramer/agarose scaffold for tissue
engineering: synthesis, characterization, drug release and cell culture study. J.
Biomater. Sci., Polym. Ed. 2017, 28 (15), 1617-1638.
31. Sarwar, M. S.; Huang, Q. R.; Ghaffar, A.; Abid, M. A.; Zafar, M. S.; Khurshid,
Z.; Latif, M., A smart drug delivery system based on biodegradable
chitosan/poly(allylamine hydrochloride) blend films. Pharmaceutics 2020, 12 (2), 131.
32. Kumar, S.; Ye, F.; Dobretsov, S.; Dutta, J., Chitosan nanocomposite coatings
for food, paints, and water treatment applications. Appl. Sci. 2019, 9 (12), 2409.
33. Jindal, A.; Mondal, T.; Bhattacharya, J., An in vitro evaluation of zinc silicate
fortified chitosan scaffolds for bone tissue engineering. Int. J. Biol. Macromol. 2020,
164, 4252-4262.
34. Sun, M.; Wang, T.; Pang, J.; Chen, X.; Liu, Y., Hydroxybutyl chitosan centered
biocomposites for potential curative applications: a critical review. Biomacromolecules
2020, 21 (4), 1351-1367.
3. Polyaniline-Chitosan Microgels
128
35. Ali Khan, Z.; Jamil, S.; Akhtar, A.; Mustehsan Bashir, M.; Yar, M., Chitosan
based hybrid materials used for wound healing applications-A short review. Int. J.
Polym. Mater. Polym. Biomater. 2020, 69 (7), 419-436.
36. Yezer, I.; Demirkol, D. O., Cellulose acetate–chitosan based electrospun
nanofibers for bio-functionalized surface design in biosensing. Cellulose 2020, 27 (17),
10183-10197.
37. Campos, E. V. R.; Oliveira, J. L.; Fraceto, L. F., Poly(ethylene glycol) and
cyclodextrin-grafted chitosan: from methodologies to preparation and potential
biotechnological applications. Front. Chem. 2017, 5 (93).
38. Ulutürk, C.; Alemdar, N., Electroconductive 3D polymeric network production
by using polyaniline/chitosan-based hydrogel. Carbohydr. Polym. 2018, 193, 307-315.
39. Moutsatsou, P.; Coopman, K.; Georgiadou, S., Biocompatibility assessment of
conducting PANI/chitosan nanofibers for wound healing applications. Polymers 2017,
9 (12), 687.
40. Mohammadi, B.; Pirsa, S.; Alizadeh, M., Preparing chitosan–polyaniline
nanocomposite film and examining its mechanical, electrical, and antimicrobial
properties. Polym. Polym. Compos. 2019, 27 (8), 507-517.
41. Torvi, A.; Naik, S.; Kariduraganavar, M., Development of supercapacitor
systems based on binary and ternary nanocomposites using chitosan, graphene and
polyaniline. Chem. Data Collect. 2018, 17-18, 459-471.
42. Sadoughi, F.; Mansournia, M. A.; Mirhashemi, S. M., The potential role of
chitosan-based nanoparticles as drug delivery systems in pancreatic cancer. IUBMB
Life 2020, 72 (5), 872-883.
43. Abdul Khalil, H. P. S.; Adnan, A. S.; Yahya, E. B.; Olaiya, N. G.; Safrida, S.;
Hossain, M. S.; Balakrishnan, V.; Gopakumar, D. A.; Abdullah, C. K.; Oyekanmi, A.
A.; Pasquini, D., A review on plant cellulose nanofibre-based aerogels for biomedical
applications. Polymers 2020, 12 (8), 1759.
44. (a) Zouaoui, F.; Bourouina-Bacha, S.; Bourouina, M.; Jaffrezic-Renault, N.;
Zine, N.; Errachid, A., Electrochemical sensors based on molecularly imprinted
chitosan: A review. TrAC, Trends Anal. Chem. 2020, 130, 115982; (b) Khalid, M.;
Honorato, A. M. B.; Varela, H.; Dai, L., Multifunctional electrocatalysts derived from
conducting polymer and metal organic framework complexes. Nano Energy 2018, 45,
127-135.
45. Kumar Meena, L.; Rather, H.; Kedaria, D.; Vasita, R., Polymeric microgels for
bone tissue engineering applications-a review. Int. J. Polym. Mater. Polym. Biomater.
2020, 69 (6), 381-397.
46. Allan, G. G.; Peyron, M., Molecular weight manipulation of chitosan I: kinetics
of depolymerization by nitrous acid. Carbohydr. Res. 1995, 277 (2), 257-272.
47. Marcasuzaa, P.; Reynaud, S.; Ehrenfeld, F.; Khoukh, A.; Desbrieres, J.,
Chitosan-graft-polyaniline-based hydrogels: elaboration and properties.
Biomacromolecules 2010, 11 (6), 1684-1691.
48. (a) Tiwari, A.; Shukla, S. K., Chitosan-g-polyaniline: a creatine
amidinohydrolase immobilization matrix for creatine biosensor. Express Polym. Lett.
2009, 3 (9), 553-559; (b) Tiwari, A.; Gong, S., Electrochemical synthesis of chitosan-
co-polyaniline/WO3⋅nH2O composite electrode for amperometric detection of NO2 gas.
Electroanalysis 2008, 20 (16), 1775-1781.
3. Polyaniline-Chitosan Microgels
129
49. (a) Wei, Y.; Jang, G. W.; Chan, C. C.; Hsueh, K. F.; Hariharan, R.; Patel, S. A.;
Whitecar, C. K., Polymerization of aniline and alkyl ring-substituted anilines in the
presence of aromatic additives. J. Phys. Chem. 1990, 94 (19), 7716-7721; (b) Wei, Y.;
Hariharan, R.; Patel, S. A., Chemical and electrochemical copolymerization of aniline
with alkyl ring-substituted anilines. Macromolecules 1990, 23 (3), 758-764; (c) Wei,
Y.; Tang, X.; Sun, Y.; Focke, W. W., A study of the mechamism of aniline
polymerization. Polym. Sci. Pol. Chem. 1989, 27 (7), 2385-2396.
50. Wu, T.; Li, Y.; Lee, D. S., Chitosan-based composite hydrogels for biomedical
applications. Macromol. Res. 2017, 25 (6), 480-488.
51. Ramadani, A. H.; Ningrum, R. S., Effectiveness of eco-absorbent modified
chitosan membrane from Pila ampullacea as urban water filter to provide healthy
sanitary water in Kediri. IOP Conf. Ser. Earth Environ. Sci. 2019, 308, 012036.
52. Zahiri, M.; Khanmohammadi, M.; Goodarzi, A.; Ababzadeh, S.; Sagharjoghi
Farahani, M.; Mohandesnezhad, S.; Bahrami, N.; Nabipour, I.; Ai, J., Encapsulation of
curcumin loaded chitosan nanoparticle within poly (ε-caprolactone) and gelatin fiber
mat for wound healing and layered dermal reconstitution. Int. J. Polym. Mater. Polym.
Biomater. 2020, 153, 1241-1250.
53. Chawla, S.; Rawal, R.; Pundir, C. S., Fabrication of polyphenol biosensor based
on laccase immobilized on copper nanoparticles/chitosan/multiwalled carbon
nanotubes/polyaniline-modified gold electrode. J. Biotechnol. 2011, 156 (1), 39-45.
54. Xu, X.; Ren, G.; Cheng, J.; Liu, Q.; Li, D.; Chen, Q., Self-assembly of
polyaniline-grafted chitosan/glucose oxidase nanolayered films for electrochemical
biosensor applications. J. Mater. Sci. 2006, 41 (15), 4974-4977.
55. Zengin, H.; Zhou, W.; Jin, J.; Czerw, R.; Smith, D. W.; Echegoyen, L.; Carroll,
D. L.; Foulger, S. H.; Ballato, J., Carbon nanotube doped polyaniline. J. Adv. Mater.
2002, 14 (20), 1480-1483.
56. Guo, B.; Finne-Wistrand, A.; Albertsson, A. C., Facile synthesis of degradable
and electrically conductive polysaccharide hydrogels. Biomacromolecules 2011, 12 (7),
2601-2609.
57. Cabuk, M.; Yavuz, M.; Unal, H. I., Electrokinetic properties of biodegradable
conducting polyaniline-graft-chitosan copolymer in aqueous and non-aqueous media.
Colloids Surf., A 2014, 460, 494-501.
58. Guo, B.; Finne-Wistrand, A.; Albertsson, A.-C., Facile synthesis of degradable
and electrically conductive polysaccharide hydrogels. Biomacromolecules 2011, 12 (7),
2601-2609.
59. D., A., New polymers for special applications. InTech: 2012.
60. (a) Chiang, J.-C.; MacDiarmid, A. G., ‘Polyaniline’: Protonic acid doping of
the emeraldine form to the metallic regime. Synth. Met. 1986, 13 (1), 193-205; (b)
Macdiarmid, A. G.; Chiang, J.-C.; Huang, W.; Humphrey, B. D.; Somasiri, N. L. D.,
Polyaniline: Protonic acid doping to the metallic regime. Mol. Cryst. Liq. Cryst. 1985,
125 (1), 309-318.
61. Ding, H.; Zhong, M.; Kim, Y. J.; Pholpabu, P.; Balasubramanian, A.; Hui, C.
M.; He, H.; Yang, H.; Matyjaszewski, K.; Bettinger, C. J., Biologically derived soft
conducting hydrogels using heparin-doped polymer networks. ACS Nano 2014, 8 (5),
4348-4357.
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4. Dual-Degradable Dextran-Chitosan Microgels
131
4. Dual-Degradable Dextran-Chitosan Microgels
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)
4. Dual-Degradable Dextran-Chitosan Microgels
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,
4. Dual-Degradable Dextran-Chitosan Microgels
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.
4. Dual-Degradable Dextran-Chitosan Microgels
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)-
4. Dual-Degradable Dextran-Chitosan Microgels
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 Experimental Section
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
4. Dual-Degradable Dextran-Chitosan Microgels
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
4. Dual-Degradable Dextran-Chitosan Microgels
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
4. Dual-Degradable Dextran-Chitosan Microgels
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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
4. Dual-Degradable Dextran-Chitosan Microgels
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
4. Dual-Degradable Dextran-Chitosan Microgels
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
4. Dual-Degradable Dextran-Chitosan Microgels
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.
4. Dual-Degradable Dextran-Chitosan Microgels
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4.2.9 Drug Loading and Release Studies
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).
4. Dual-Degradable Dextran-Chitosan Microgels
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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.
4. Dual-Degradable Dextran-Chitosan Microgels
145
4.3 Results and Discussion
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.
4. Dual-Degradable Dextran-Chitosan Microgels
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.
4. Dual-Degradable Dextran-Chitosan Microgels
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.
4. Dual-Degradable Dextran-Chitosan Microgels
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
4. Dual-Degradable Dextran-Chitosan Microgels
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).
4.3.2 Influence of pH on Microgel Size and Electrophoretic Mobility
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
4. Dual-Degradable Dextran-Chitosan Microgels
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.
4. Dual-Degradable Dextran-Chitosan Microgels
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4.3.3 Degradation of Microgels
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.
4. Dual-Degradable Dextran-Chitosan Microgels
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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.
4. Dual-Degradable Dextran-Chitosan Microgels
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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.
4. Dual-Degradable Dextran-Chitosan Microgels
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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.
4. Dual-Degradable Dextran-Chitosan 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.
4. Dual-Degradable Dextran-Chitosan Microgels
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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).
4. Dual-Degradable Dextran-Chitosan Microgels
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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.
4. Dual-Degradable Dextran-Chitosan Microgels
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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
4. Dual-Degradable Dextran-Chitosan Microgels
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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.
4.3.5 Drug Loading and Release Studies
An antibiotic glycopeptide, VM, was selected as a model drug to investigate
the drug loading and triggered release profiles of the obtained biohybrid
4. Dual-Degradable Dextran-Chitosan Microgels
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
4. Dual-Degradable Dextran-Chitosan Microgels
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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
4. Dual-Degradable Dextran-Chitosan Microgels
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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
4. Dual-Degradable Dextran-Chitosan Microgels
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expected to have potential applications as bioactive delivery vehicles for the
release of the drugs at specific sites, such as the colonic region.
4.5 References and Notes
1. Vinod, A.; Sanjay, M. R.; Suchart, S.; Jyotishkumar, P., Renewable and
sustainable biobased materials: An assessment on biofibers, biofilms, biopolymers and
biocomposites. J. Cleaner Prod. 2020, 258, 120978.
2. Neamtu, I.; Rusu, A. G.; Diaconu, A.; Nita, L. E.; Chiriac, A. P., Basic concepts
and recent advances in nanogels as carriers for medical applications. Drug Delivery
2017, 24 (1), 539-557.
3. Jo, Y. K.; Lee, D., Biopolymer microparticles prepared by microfluidics for
biomedical applications. Small 2020, 16 (9), 1903736.
4. Wong, E. H. M.; Rondeau, E.; Schuetz, P.; Cooper-White, J., A microfluidic-
based method for the transfer of biopolymer particles from an oil phase to an aqueous
phase. Lab Chip 2009, 9 (17), 2582-2590.
5. Senapati, S.; Mahanta, A. K.; Kumar, S.; Maiti, P., Controlled drug delivery
vehicles for cancer treatment and their performance. Signal Transduction Targeted
Ther. 2018, 3 (1), 7.
6. Sacco, P.; Borgogna, M.; Travan, A.; Marsich, E.; Paoletti, S.; Asaro, F.; Grassi,
M.; Donati, I., Polysaccharide-based networks from homogeneous chitosan-
tripolyphosphate hydrogels: synthesis and characterization. Biomacromolecules 2014,
15 (9), 3396-3405.
7. Lee, C. F.; Wen, C. J.; Chiu, W. Y., Synthesis of poly(chitosan-N-
isopropylacrylamide) complex particles with the method of soapless dispersion
polymerization. J. Polym. Sci. Pol. Chem. 2003, 41 (13), 2053-2063.
8. Li, Y.; Zhang, Z. S.; van Leeuwen, H. P.; Stuart, M. A. C.; Norde, W.; Kleijn,
J. M., Uptake and release kinetics of lysozyme in and from an oxidized starch polymer
microgel. Soft Matter 2011, 7 (21), 10377-10385.
9. Stepczyńska, M.; Rytlewski, P., Enzymatic degradation of flax-fibers
reinforced polylactide. Int. Biodeterior. Biodegrad. 2018, 126, 160-166.
10. Tiwari, S.; Patil, R.; Bahadur, P., Polysaccharide based scaffolds for soft tissue
engineering applications. Polymers 2019, 11 (1), 1.
11. Kuznetsova, T. A.; Andryukov, B. G.; Besednova, N. N.; Zaporozhets, T. S.;
Kalinin, A. V., Marine algae polysaccharides as basis for wound dressings, drug
delivery, and tissue engineering: A review. J. mar. sci. 2020, 8 (7), 481.
12. Gopinath, V.; Saravanan, S.; Al-Maleki, A. R.; Ramesh, M.; Vadivelu, J., A
review of natural polysaccharides for drug delivery applications: Special focus on
cellulose, starch and glycogen. Biomed. Pharmacother. 2018, 107, 96-108.
13. Dheer, D.; Arora, D.; Jaglan, S.; Rawal, R. K.; Shankar, R., Polysaccharides
based nanomaterials for targeted anti-cancer drug delivery. JoJ. Drug Targeting 2017,
25 (1), 1-16.
4. Dual-Degradable Dextran-Chitosan Microgels
164
14. Sinha, V. R.; Kumria, R., Polysaccharides in colon-specific drug delivery. Int.
J. Pharm. 2001, 224 (1-2), 19-38.
15. Beata Łabowska, M.; Michalak, I.; Detyna, J., Methods of extraction,
physicochemical properties of alginates and their applications in biomedical field-a
review. Open Chem. 2019, 17 (1), 738-762.
16. Han, H.; Zhang, Y.; Jin, S.; Chen, P.; Liu, S.; Xie, Z.; Jing, X.; Wang, Z.,
Paclitaxel-loaded dextran nanoparticles decorated with RVG29 peptide for targeted
chemotherapy of glioma: an in vivo study. New J. Chem. 2020, 44 (15), 5692-5701.
17. Dodero, A.; Brunengo, E.; Alloisio, M.; Sionkowska, A.; Vicini, S.; Castellano,
M., Chitosan-based electrospun membranes: Effects of solution viscosity, coagulant
and crosslinker. Carbohydr. Polym. 2020, 235, 115976.
18. Munair, B.; Hanif, U.; Fazli, W.; Taous, K., Bacterial cellulose-based metallic
green nanocomposites for biomedical and pharmaceutical applications. Curr. Pharm.
Des. 2020, 26, 1-15.
19. Selvakumar, G.; Lonchin, S., Fabrication and characterization of collagen-
oxidized pullulan scaffold for biomedical applications. Int. J. Biol. Macromol. 2020,
164, 1592-1599.
20. Muramatsu, K.; Tajima, Y.; Kaneko, R.; Yanagita, Y.; Hirai, H.; Hiura, N.,
Characterization of poly(L-glutamic acid)-grafted hyaluronan as a novel candidate
medicine and biomedical device for intra-articular injection. J. Biomed. Mater. Res.,
Part A 2017, 105 (11), 3006-3016.
21. Bang, S.; Jung, U.-W.; Noh, I., Synthesis and biocompatibility characterizations
of in situ chondroitin sulfate–gelatin hydrogel for tissue engineering. Tissue Eng.
Regener. Med. 2018, 15 (1), 25-35.
22. Rosellini, E.; Zhang, Y. S.; Migliori, B.; Barbani, N.; Lazzeri, L.; Shin, S. R.;
Dokmeci, M. R.; Cascone, M. G., Protein/polysaccharide-based scaffolds mimicking
native extracellular matrix for cardiac tissue engineering applications. J. Biomed.
Mater. Res., Part A 2018, 106 (3), 769-781.
23. Du, X.; Liu, Y.; Wang, X.; Yan, H.; Wang, L.; Qu, L.; Kong, D.; Qiao, M.;
Wang, L., Injectable hydrogel composed of hydrophobically modified
chitosan/oxidized-dextran for wound healing. Mater. Sci. Eng. C 2019, 104, 109930.
24. Zamboulis, A.; Nanaki, S.; Michailidou, G.; Koumentakou, I.; Lazaridou, M.;
Ainali, N. M.; Xanthopoulou, E.; Bikiaris, D. N., Chitosan and its derivatives for ocular
delivery formulations: Recent advances and developments. Polymers 2020, 12 (7),
1519.
25. Yucel Falco, C.; Falkman, P.; Risbo, J.; Cárdenas, M.; Medronho, B., Chitosan-
dextran sulfate hydrogels as a potential carrier for probiotics. Carbohydr. Polym. 2017,
172, 175-183.
26. Qu, B.; Luo, Y., Chitosan-based hydrogel beads: Preparations, modifications
and applications in food and agriculture sectors – A review. Int. J. Biol. Macromol.
2020, 152, 437-448.
27. Guo, B.; Qu, J.; Zhao, X.; Zhang, M., Degradable conductive self-healing
hydrogels based on dextran-graft-tetraaniline and N-carboxyethyl chitosan as
injectable carriers for myoblast cell therapy and muscle regeneration. Acta Biomater.
2019, 84, 180-193.
4. Dual-Degradable Dextran-Chitosan Microgels
165
28. Mansuroğlu, B.; Kızılbey, K.; Şayan Poyraz, F.; Yurttaş, Z.; Fuerkaiti, S. N.;
Abaoğlu, İ. Y.; Başat, H. N., Chitosan/dextran sulphate sodium hydrogels for wound
healing material: preparation, characterisation and in vitro evaluation. Mater. Technol.
2020, 1-8.
29. De Queiroz Antonino, R. S. C. M.; Lia Fook, B. R. P.; De Oliveira Lima, V. A.;
De Farias Rached, R. Í.; Lima, E. P. N.; Da Silva Lima, R. J.; Peniche Covas, C. A.;
Lia Fook, M. V., Preparation and characterization of chitosan obtained from shells of
shrimp (litopenaeus vannamei boone). Mar. Drugs 2017, 15 (5), 141.
30. Soon, C. Y.; Tee, Y. B.; Tan, C. H.; Rosnita, A. T.; Khalina, A., Extraction and
physicochemical characterization of chitin and chitosan from Zophobas morio larvae
in varying sodium hydroxide concentration. Int. J. Biol. Macromol. 2018, 108, 135-
142.
31. Bakshi, P. S.; Selvakumar, D.; Kadirvelu, K.; Kumar, N. S., Chitosan as an
environment friendly biomaterial-a review on recent modifications and applications.
Int. J. Biol. Macromol. 2020, 150, 1072-1083.
32. (a) Aktuganov, G. E.; Melent’ev, A. I., Specific features of chitosan
depolymerization by chitinases, chitosanases, and nonspecific enzymes in the
production of bioactive chitooligosaccharides (Review). Appl. Biochem. Microbiol.
2017, 53 (6), 611-627; (b) Rathinam, K.; Singh, S. P.; Arnusch, C. J.; Kasher, R., An
environmentally-friendly chitosan-lysozyme biocomposite for the effective removal of
dyes and heavy metals from aqueous solutions. Carbohydr. Polym. 2018, 199, 506-515.
33. Bhavsar, C.; Momin, M.; Gharat, S.; Omri, A., Functionalized and graft
copolymers of chitosan and its pharmaceutical applications. Expert Opin. Drug
Delivery 2017, 14 (10), 1189-1204.
34. Treenate, P.; Monvisade, P., In vitro drug release profiles of pH-sensitive
hydroxyethylacryl chitosan/sodium alginate hydrogels using paracetamol as a soluble
model drug. Int. J. Biol. Macromol. 2017, 99, 71-78.
35. Chu, C. W.; Ryu, J. H.; Jeong, Y.-I. L.; Kwak, T. W.; Lee, H. L.; Kim, H. Y.;
Son, G. M.; Kim, H. W.; Kang, D. H., Redox-responsive nanophotosensitizer
composed of chlorin e6-conjugated dextran for photodynamic treatment of colon
cancer cells. J. Nanomater. 2016, 2016, 4075803.
36. Allan, G. G.; Peyron, M., Molecular weight manipulation of chitosan I: kinetics
of depolymerization by nitrous acid. Carbohydr. Res. 1995, 277 (2), 257-272.
37. De Geest, B. G.; Van Camp, W.; Du Prez, F. E.; De Smedt, S. C.; Demeester,
J.; Hennink, W. E., Degradable multilayer films and hollow capsules via a 'Click'
strategy. Macromol. Rapid Commun. 2008, 29 (12-13), 1111-1118.
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.
4. Dual-Degradable Dextran-Chitosan Microgels
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
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5. Conclusion and Outlook
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
5. Conclusion and Outlook
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,
5. Conclusion and Outlook
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.
5. Conclusion and Outlook
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.
5.1 References and Notes
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
7. List of Publications
[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.
7. List of Publications
174