Generic Films and Fibers from Polysaccharides: 
Chitosan and Alginate 
by 
Ramiz Boy 
 
 
A thesis submitted to the Graduate Faculty of 
Auburn University 
in partial fulfillment of the 
requirements for the Degree of 
Master of Science 
 
Auburn, Alabama 
August 06, 2011 
 
 
 
Keywords: Polysaccharides, chitosan, alginate, biomaterials 
Copyright 2010 by Ramiz BOY 
 
 
Approved by 
Maria L. Auad, Chair, Professor of Polymer and Fiber Engineering 
Roy Broughton, Jr., Professor Emeritus of Polymer and Fiber Engineering 
Peter Schwartz, Professor of Polymer and Fiber Engineering 
 
 
ii 
 
 
 
Abstract 
Chitosan and alginate are both extracted from natural resources, and both are 
biocompatible, biodegradable, and hydrophilic. Thus, chitosan-alginate biopolymer 
mixtures have the potential to form new functional polymers towards better 
performance, which can be utilized in different applications. Chitosan-alginate polymer 
films and fibers were prepared. Possible interactions and the effects of each component 
were investigated. 
Miscibility, thermal stability, mechanical, and morphological properties of 
chitosan-alginate films were studied. We found that the film with ratio of 60:40 chitosan 
to alginate has the highest modulus and the highest tensile strength. The film with 80% 
chitosan content shows antibacterial activity. Furthermore, we developed a novel 
method to form wet-spun chitosan-alginate fibers with no additive and analyzed their 
mechanical and morphological properties. Roughness of the fibers increased with more 
chitosan content which reversely affected the tenacity. 
 
 
 
 
 
 
 
iii 
 
  
 
Acknowledgments 
I would like to express my sincere thanks to my advisor, Dr. Maria L. Auad, for 
her guidance and patience throughout this work. I am grateful to Dr. Roy Broughton, Jr. 
for his contribution and critical suggestions. I would also thank Dr. Peter Schwartz for 
being committee members and reviewing my research. Thanks to the Department of 
Polymer and Fiber Engineering for providing support and welcoming environment. I also 
would like to mention that this research would not have been possible without the 
support of Ministry of Turkish National Education. 
I am most grateful to my family who have supported me and provided motivation 
all along my life. I would like also to thank all my friends who have helped me along this 
research. 
 
 
 
 
 
 
 
 
 
 
iv 
 
 
 
Table of Contents 
 
Abstract ............................................................................................................................ii 
Acknowledgments ........................................................................................................... iii 
List of Tables .................................................................................................................. vii 
List of Figures ................................................................................................................ viii 
List of Abbreviations ........................................................................................................ x 
Chapter 1 Inroduction ...................................................................................................... 1 
1.1 Literature Review ................................................................................................... 1 
1.1.1 Polysaccharides ............................................................................................... 1 
1.1.2 Polysaccharide Mixtures .................................................................................. 5 
1.1.2.1 Binary mixtures .......................................................................................... 5 
1.1.2.2 Phase separated gels ................................................................................ 7 
1.1.3 Polyelectrolyte Complexation ........................................................................... 8 
1.1.4 Polysaccharides as Biometarials ..................................................................... 9 
1.1.4.1 Alginate ..................................................................................................... 9 
1.1.4.2 Chitosan .................................................................................................. 15 
1.2 Research Objectives ............................................................................................ 21 
    References ................................................................................................................ 22 
Chapter 2 Characterization of Chitosan-Alginate Films ................................................. 26 
2.1 Introduction .......................................................................................................... 26 
2.2 Experimental ........................................................................................................ 31 
v 
 
2.2.1 Materials ........................................................................................................ 31 
2.2.2 Methods and Techniques............................................................................... 31 
2.2.2.1 Nuclear magnetic resonance (NMR) spectroscopy ................................. 31 
2.2.2.2 Zeta (?) potential measurements ............................................................. 32 
2.2.2.3 Film preparation....................................................................................... 33 
2.2.2.4 Light transmission properties ................................................................... 34 
2.2.2.5 Infrared spectroscopy characterization .................................................... 34 
2.2.2.6 Thermogravimetric analysis (TGA) .......................................................... 35 
2.2.2.7 Differential scanning calorimetry (DSC) ................................................... 35 
2.2.2.8 Dynamical mechanical analysis (DMA) ................................................... 35 
2.2.2.9 Scanning electronic microscopy analyses ............................................... 36 
2.2.2.10 Mechanical properties ........................................................................... 36 
2.2.2.11 Antibacterial testing - Zone of inhibition test .......................................... 36 
2.3 Result and Discussion.......................................................................................... 37 
2.3.1 NMR study for the determination of the degree of deacetylation ................... 37 
2.3.2 Zeta (?) potential measurements ................................................................... 38 
2.3.3 Light transmission properties ......................................................................... 39 
2.3.4 Infrared spectroscopy .................................................................................... 41 
2.3.5 Thermal analysis ............................................................................................ 44 
2.3.6 Morphology of the films .................................................................................. 50 
2.3.7 Mechanical properties .................................................................................... 52 
2.3.8 Antibacterial testing - Zone of inhibition test .................................................. 56 
2.4 Conclusion ........................................................................................................... 59 
vi 
 
    References ................................................................................................................ 61 
Chapter 3 Chitosan and Alginate Wet Spun Fibers ....................................................... 64 
3.1 Introduction .......................................................................................................... 64 
3.2 Experimental ........................................................................................................ 67 
3.2.1 Materials ........................................................................................................ 67 
3.2.2 Methods and Techniques............................................................................... 68 
3.2.2.1 Chitosan solution preparation .................................................................. 68 
3.2.2.2 Ultra-Sonication ....................................................................................... 68 
3.2.2.3 Determination of the average molecular weight of chitosan .................... 68 
3.2.2.4 Preparation of the sodium-alginate spinning solution .............................. 70 
3.2.2.5 Formation of the alginate-chitosan fibers ................................................. 70 
3.2.2.6 Diameter of fibers and the linear density ................................................. 73 
3.2.2.7 Total nitrogen analysis ............................................................................. 74 
3.2.2.8 Morphology analyses ............................................................................... 74 
3.2.2.9 Mechanical properties ............................................................................. 74 
3.3 Result and Discussion.......................................................................................... 75 
3.3.1 Average molecular weight of chitosan ........................................................... 75 
3.3.2 The chitosan content of fibers ........................................................................ 76 
3.3.3 Morphological characteristics of fibers ........................................................... 77 
3.3.4 Mechanical properties .................................................................................... 81 
3.4 Conclusion ........................................................................................................... 83 
    References ................................................................................................................ 84 
 
vii 
 
 
 
List of Tables 
 
Table 2. 1 Chitosan-alginate film compositions ............................................................. 33 
Table 2. 2 Chitosan chemical shifts of proton at room conditions in CD3COOD/D2O .... 38 
Table 2. 3 FTIR bands of chitosan, alginate and 60:40 CH:AL films with assignments . 42 
Table 2. 4 Mechanical properties of chitosan-alginate films .......................................... 55 
 
Table 3. 1 Viscosities and molecular weights of chitosan solutions .............................. 76 
Table 3. 2 The chitosan content of fibers ...................................................................... 77 
Table 3. 3 Changes of roughness of the fibers in longitude and in circumference ........ 78 
 
 
 
 
 
 
 
 
 
 
 
 
viii 
 
 
 
List of Figures 
 
Figure 1. 1 Structure of D-glucose .................................................................................. 2 
Figure 1. 2 Cellulosic chain formed from condensed D-glucose ..................................... 3 
Figure 1. 3 Types of binary polysaccharide gel-structure: Polysaccharide A (?), 
polysaccharide B (?) (a) single polymer network containing the second polymer within 
the gel (swollen network), (b) interpenetrating networks, (c) phase-separated networks, 
(d) coupled network ......................................................................................................... 6 
Figure 1. 4 Chemical structure of alginates: (I) Alginate monomers, (II) chain 
conformation, (III) block distribution............................................................................... 10 
Figure 1. 5 Brown seaweed ........................................................................................... 11 
Figure 1. 6 Egg-box structure of an alginate gel formed by gelation of Ca2+ ions ......... 12 
Figure 1. 7 Crustaceans ................................................................................................ 15 
Figure 1. 8 Chemical structure of chitin ......................................................................... 16 
Figure 1. 9 Chemical structure chitosan ........................................................................ 17 
 
Figure 2. 1 Structure of chitosan ................................................................................... 27 
Figure 2. 2 Structure of alginate .................................................................................... 29 
Figure 2. 3 Structural scheme of chitosan?sodium alginate ion complex formation 
reaction between anion group (?COO-) of sodium alginate and the protonated cation 
group (?NH3+) of chitosan ............................................................................................. 30 
Figure 2. 4 Illustration of film preparation process ......................................................... 34 
Figure 2. 5 1H NMR spectrum of chitosan (DD = 80%), in CD3COOD/D2O, at 400 MHz
 ...................................................................................................................................... 37 
Figure 2. 6 Zeta potential of the solutions of chitosan, alginate and their mixture ......... 39 
ix 
 
Figure 2. 7 Light transmission of different composition of films ..................................... 40 
Figure 2. 8 50:50 CH:AL film ......................................................................................... 41 
Figure 2. 9 FTIR spectra of chitosan (100:0), alginate (0:100) and 60:40 CH:AL films . 42 
Figure 2. 10 TGA thermograms of chitosan (100:0 CH:AL), alginate (0:100 CH:AL) and 
50:50 CH:AL films ......................................................................................................... 45 
Figure 2. 11 DSC thermograms of chitosan (100:0 CH:AL), alginate (0:100 CH:AL), and 
50:50 CH:AL films. a) first run b) second run ................................................................ 46 
Figure 2. 12 Dynamic temperature ramp test of films: a) E? (storage modulus), b) tan ? 
versus temperature ....................................................................................................... 48 
Figure 2. 13 SEM pictures of a, b) 100:0 CH:AL, c, d) 50:50 CH:AL and e, f) 0:100 
CH:AL films ................................................................................................................... 51 
Figure 2. 14 Mechanical properties of chitosan-alginate films: a) Modulus, b) Yield 
strength, c) Deformation at break, d) Stress at break .................................................... 54 
Figure 2. 15 Representative picture of inhibitory effect of the selected films: 100:0, 
60:40, 80:20 and 0:100 CH:AL films.............................................................................. 57 
 
Figure 3. 1 An illustration of the fiber formation ............................................................. 71 
Figure 3. 2 Fiber produced with non-treated chitosan solution ...................................... 72 
Figure 3. 3 The increase of the diameters of fibers with the retention time ................... 73 
Figure 3. 4 Huggins and Kraemer plot of non-treated chitosan solution ........................ 75 
Figure 3. 5 Huggins and Kraemer plot of 20 min. sonicated chitosan solution .............. 75 
Figure 3. 6 Optical microscope pictures of fibers; a) 1h, b) 3h, c) 6h, d) 12h, e) 24h 
retention time ................................................................................................................ 79 
Figure 3. 7 Representative SEM images of surface of fibers coagulated with increasing 
retention time in chitosan solution; a, b) 1h, c, d) 12h, e, f) 24h .................................... 80 
Figure 3. 8 a) Tenacity, b) Elongation at break and c) Young?s modulus of fibers with 
increasing retention time ............................................................................................... 82 
 
x 
 
 
 
List of Abbreviations 
 
CH Chitosan 
AL Alginate 
MMW Medium Molecular Weight 
LMW Low Molecular Weight 
?  Zeta Potential 
NMR Nuclear Magnetic Resonance Spectroscopy 
UV-VIS Ultra Violet-Visible Spectroscopy 
FTIR Fourier Transform Infrared 
TGA Thermal Gravimetric Analysis 
DSC Differential Scanning Calorimetry 
DMA Dynamic Mechanical Analysis 
SEM Scanning Electron Microscopy 
E? Storage modulus 
MPa Megapascal 
kDa Kilodalton 
ASTM American Society for Testing and Materials 
 
 
1 
 
 
 
CHAPTER ONE 
INTRODUCTION 
 
1.1 LITERATURE REVIEW 
1.1.1 Polysaccharides 
Applications of polymers have been exploited in almost all disciplines of 
medicine, from complex tailored implants to drug delivery materials and practical 
devices situated outside the body (Dumitriu, 2002). Because every medical application 
is designed with its particularly specialized needs, a wide array of materials including 
high biocompatibility but distinct chemical and physical characteristics must be present 
(Dumitriu, 2002). 
Most of the carbohydrate materials in nature exist in the form of polysaccharides 
(d?Ayala et al., 2008). They exhibit ideal biocompatibility and biodegradability, which are 
the fundamental characteristics for polymers exerted as biomaterials (Dumitriu, 2002). 
In addition, polysaccharides are not only involved in those substances just composed of 
glycosidically linked sugar structures, but also molecules that contain polymeric 
saccharide backbones linked via covalent bonds to amino acids, peptides, proteins, 
lipids, and other structures (d?Ayala et al., 2008). They possess various properties not 
available in other naturally occurring polymers, such as proteins and nucleic acids 
2 
 
(Dumitriu, 2002). Lately, several classes of polysaccharides have been applied for 
specific properties of antivirals, antitumorals, and gene modulators (Dumitriu, 2002).  
Polysaccharides, also known as glycans, are composed of mono-saccharides 
and their derivatives (d?Ayala et al., 2008). If a polysaccharide comprises only one type 
of monosaccharide molecule, it is called homo-polysaccharide, or homoglycan, while 
the ones which have more than one type of monosaccharide are hetero-
polysaccharides (d?Ayala et al., 2008). The well-known component of polysaccharides is 
D-glucose (Figure 1.1) ? besides, D-fructose, D-galactose, L-galactose, D-mannose, L-
arabinose, and D-xylose are also common (d?Ayala et al., 2008). 
 
 
Figure 1. 1 Structure of D-glucose 
 
Some monosaccharide derivatives occurred in polysaccharides contains the 
amino sugars (D-glucosamine and D-galactosamine) as well as their derivatives (N-
acetylneuraminic acid and N-acetylmuramic acid), and simple sugar acids (glucuronic 
and iduronic acids). Homo-polysaccharides are often identified for the sugar unit they 
have; therefore, glucose homo-polysaccharides are called glucans, whereas mannose 
homo-polysaccharides are mannans. Polysaccharides vary in the character of their 
3 
 
component monosaccharides, in the length of their chains and in the amount of chain 
branching that occurs. Even though any sugar residue possesses only one anomeric 
carbon and also can form only one glycosidic linkage with hydroxyl groups on other 
molecules, each sugar residue carries several hydroxyls, one or more of which may be 
an acceptor of glycosyl substituents. This enables them to form branched structures that 
separates polysaccharides from proteins and nucleic acids, which occur only as linear 
polymers (d?Ayala et al., 2008).  
The significant functions of polysaccharides in nature are either storage or 
structural purposes (d?Ayala et al., 2008). The prevalent storage polysaccharide in 
plants is starch, which occurs in two types: ?-amylose and amylopectin. Structural 
polysaccharides display properties which differ considerably from those of the storage 
polysaccharides, while the structures of these two classes are similar. The structural 
polysaccharide cellulose (Figure 1.2) is the most abundant natural polymer on earth. 
Cellulose, exists in the cell walls of almost all plants as well as marine algae, is one of 
the main components, which gives physical structure and strength. Cellulose of animal 
origin derived from marine, tunicin, extracted from tunicates ? invertebrate sea animals 
? is a choice of material, since it is highly crystalline (Cima et al., 1996, d?Ayala et al., 
2008, Krishnan, 1975). 
 
 
Figure 1. 2 Cellulosic chain formed from condensed D-glucose 
4 
 
Algal origin polysaccharides consist of alginates, agar and carrageenans. Agar 
(or agar-agar) is an unbranched polysaccharide acquired from some species of red 
algae?s cell wall, primarily from the genuses Gelidium and Gracilaria, or seaweed, 
largely utilized as gelatin and thickener in food industry, and as a gel for electrophoresis 
in microbiology. It is chemically formed by galactose sugar molecules; that is the 
primary structural support for the algae?s cell walls. Carrageenans are polysaccharides 
of galactan with alternating 1,3- and 1,4-linked galactose residues, which appear 
between the cellulosic plant structure of seaweeds. They are consumed in the food 
processing industry for their gelling, thickening, and stabilizing properties (d?Ayala et al., 
2008). 
Following cellulose, chitin is the second most common organic compound in 
nature. Chitin is widely distributed in marine invertebrates, insects, fungi, and yeast but 
does not occur in higher plants and higher animals (d?Ayala et al., 2008, Ehrlich et al., 
2007). 
Among polysaccharides that perform structural functions in nature, in this 
literature review, we will mainly focus on alginate and chitin?s derivative chitosan. They 
are substantial not only as plentiful resources, but mainly for their appealing biological 
properties and potential in such fields mentioned above. 
 
 
 
5 
 
1.1.2 Polysaccharide Mixtures 
1.1.2.1 Binary mixtures 
Research on binary mixtures of polysaccharides has garnered much attention for 
more than two decades (Morris, 1998). A multitude of combinations of mixed 
polysaccharides make it impractical to analyze all such research. However, it is practical 
to classify binary mixtures in some simple kinds of gel (Morris, 1998). When two 
polysaccharides are blended together and then the mixture becomes gelled, a number 
of gel structure form, varying the chemistry of components, the rate and extent of 
polymer mixing, and the process of gelation (Cairns et al., 1987). Classification of these 
gels enables one to comprehend the elemental interrelationships among the properties 
of mixed gels and their constituents (Morris, 1998).  This also offers to anticipate which 
kinds of analysis is needed to determine the structures of these gels (Morris, 1998).  
These gel structures can be divided into four types (Cairns et al., 1987): swollen 
networks, phase separated networks, interpenetrating networks, and coupled networks 
(Figure 1.3). Swollen networks occur while one of the constituents of the mixture is 
forming a network and the other constituent occupies within and swells the network 
(Figure 1.3(a)). Interpenetrating networks are composed of two separated networks and 
each of them is dispersed in the total volume and interpenetrating through each other 
(Figure 1.3(b)). Phase separated networks are formed through mixing incompatible 
polysaccharides (Morris, 1998). This kind of gelation results in a composite or filled gel 
(Figure 1.3(c)). The coupled network is the most intriguing binary gel system, in which 
new heterotypic junction zones result on binding of two distinct polysaccharides (Figure 
1.3(d)). 
6 
 
 
Figure 1. 3 Types of binary polysaccharide gel-structure: Polysaccharide A (?), 
polysaccharide B (?) (a) single polymer network containing the second polymer within 
the gel (swollen network), (b) interpenetrating networks, (c) phase-separated networks, 
(d) coupled network 
 
The models exhibited in Figure 1.3 are likely to occur in a binary mixture. 
However, a given type of gel network should not be virtually always expected (Morris, 
1998). For instance, two different polysaccharides can form an interpenetrating network 
at certain concentration of both polysaccharides but at very high ratios of the 
constituents, or at low enough concentrations of the two polymers, one or both of the 
constituents might be inadequate to create a network in all over sample volume (Morris, 
1998). 
7 
 
1.1.2.2 Phase separated gels 
Phase separated mixed gels usually comprise a continual phase and a 
supporting phase having inclusions in one another (Durrani and Donald, 1994). These 
systems exhibit a phase inversion at a given concentration at which the main 
constituent in the continual and supporting phases switch over.  
Although mixed gels contain a high ratio of solvent (water), conventional polymer 
blends and mixed gels have some parallelism and functional similarities. This analogy 
inspired Clark et al. (1983) to develop a model which correlates a relationship between 
the modulus of a phase separated mixed gel and the volume fraction of the two 
component polymers in each phase. The model based on Takayanagi's approach 
(1963) was to calculate the modulus of a binary polymer blend (Clark et al., 1983). 
A major problem in the analysis of phase separated biopolymer networks is the 
detection of any phase separation, unless separation is formed on a relatively bulk scale 
through the inclusion of two different layers (Durrani and Donald, 1994). It is unlikely to 
detect independently phase separated regions in a clear macroscopic single phase 
system either by visual inspection or by optical microscopy when the refractive indices 
of the two polymers in solution are not clearly distinctive (Durrani and Donald, 1994). 
 
 
 
  
8 
 
1.1.3 Polyelectrolyte Complexation 
These complexes are composed of the assembly of two or more compatible 
polymers and may result from supramolecular interactions such as electrostatic effects, 
hydrophobic forces, hydrogen bonding, van der Waals forces, or combinations of these 
interactions (Dumitriu, 2002, Scranton et al., 1992, Tsuchida et al., 1982). The 
mechanism of complexation has a strong effect on the polymer solubility, rheology, 
conductivity, and turbidity in solutions.(Dumitriu, 2002, Scranton et al., 1992).  
Polyelectrolyte complexes are mainly formed by electrostatic forces. 
Poly(cation)/poly(anion) pairs are constituted by proton transfer from acidic polymers to 
compatible basic polymers. Besides interpolymer complexes, research on intrapolymer 
complexes of polyampholytes has been done (Scranton et al., 1992). In these systems, 
polymer complexes may occur between reversely charged components of one polymer 
chain, or more specifically on the same monomer unit of that polymer (Dumitriu, 2002). 
Finally, multivalent ions or polyelectrolyte itself may bind other polyelectrolytes in 
solution to form gels or coacervates (Chatterjee et al., 1987, Dumitriu, 2002). In 
addition, charge density, solvent, ionic strength, pH, and temperature also affect the 
formation and stability of complexation (Dumitriu, 2002). Insolubility of complexes is 
dependent upon the high charge density of polyelectrolytes (Dumitriu, 2002). Properties 
of the polyelectrolyte complexes are mainly tuned by the molecular weight of polymers 
including their cooperative interactions, concerted interactions, steric compatibility, and 
micro-environmental effects (Dumitriu, 2002, Kubota and Kikuchi, 1998). 
 
 
9 
 
1.1.4 Polysaccharides as Biometarials 
1.1.4.1 Alginate  
Alginate, a water-soluble linear binary copolymer, is isolated from brown 
seaweed and consists of alternating blocks of 1?4 linked ?-L-guluronic acid (G) and ?-
D-mannuronic acid (M) residues with randomly alternating composition and sequence 
(Draget et al., 2009, George and Abraham, 2006). Figure 1.4 displays the structures of 
mannuronic and guluronic acid residues and the links between these residues in 
alginate. Due to the particular shapes of the monomers and their manner of linkage in 
the polymer, the geometries of the G-block parts, M-block parts, and alternating parts 
are significantly discrete (George and Abraham, 2006). Specifically, the G-blocks are 
clasped whereas the M-blocks possess a shape proposed to be an extended ribbon. A 
diamond shaped hole results in if two G-block regions are aligned side by side. The 
dimensions of so-called hole are ideal for the collective binding of calcium ions. The 
homo-polymeric regions of ?-D-mannuronic acid blocks and ?-L-guluronic acid blocks 
are inter-dispersed with regions of alternating structure (?-D-mannuronic acid??-L-
guluronic acid blocks) (Haug and Larsen, 1962, Haug et al., 1967). The arrangement 
and size of the orders and the molecular weight determine the physical properties of the 
alginates. 
10 
 
 
Figure 1. 4 Chemical structure of alginates: (I) Alginate monomers, (II) chain 
conformation, (III) block distribution. 
 
Regarding their molecular weights, alginates, similar to other polysaccharides, 
are polydisperse (Draget et al., 2009). This may be induced by polysaccharides which 
are indirectly coded for by the DNA, but are separated by polymerase enzymes and 
extraction may cause depolymerization. As a result of their polydispersity, an average 
value of the molecular weight is taken. 
11 
 
In terms of the source of alginates, commercial ones are derived from three 
species of brown marine algae (George and Abraham, 2006). These are Laminaria 
hyperborean, Ascophyllum nodosum, and Macrocystis pyrifera; in which alginates 
constitute up to 40% of the dry weight (Smidsr?d and Skjaok-Brk, 1990, Sutherland, 
1991). The structural composition and its sequential order might alter dependent on 
seasonal and ambient growing conditions (Haug, 1964, Indergaard and Skj?k-Br?k, 
1987). Alginate occurs as a mixed salt of various cations encountered in the seawater 
such as Mg2+, Sr2+, Ba2+, and Na+. Bacterial alginates have also been extracted from 
Azotobacter vinelandii and several Pseudomonas species. 
 
 
Figure 1. 5 Brown seaweed 
 
The process of alginate extraction is basically a procedure of ion-exchange 
(Sime, 1990). The seaweed (Figure 1.5) is milled and washed. Then through a strong 
alkali and heat treatment, the extraction continues with the solution of sodium alginate. 
Alginate is precipitated in that solution with the addition of calcium chloride forming 
calcium alginate. A further acid treatment is applied to convert calcium alginate into 
alginic acid which finally can be treated to produce various forms of commercial 
alginates (Sime, 1990). For example,  
12 
 
1) Treatment with sodium carbonate or other bases to fabricate alginate salts 
2) Reaction with propylene oxide to fabricate propylene glycol alginate 
The process of alginate?s gelation can be performed under an extraordinarily mild 
conditions and by using non-toxic reactants (George and Abraham, 2006). The most 
substantial property of alginate is its ability to form gels by reaction with multivalent 
cations such as Ca2+. Hyrogels as well as beads, fibers and films of alginate can be 
made by extruding a solution of sodium alginate into a divalent cross-linking solution, 
such as Ca2+, Sr2+, or Ba2+ (George and Abraham, 2006). Monovalent cations and Mg2+ 
ions do not cause gelation (George and Abraham, 2006, Rees and Welsh, 1977). The 
gelation and cross-linking of the polymers are mostly accomplished through the 
replacement of sodium ions on the guluronic acids with the divalent cations, and the 
packing of these guluronic groups to form the characteristic egg-box structure shown in 
Figure 1.6. 
 
 
Figure 1. 6 Egg-box structure of an alginate gel formed by gelation of Ca2+ ions 
 
13 
 
The segments of poly-L-guluronate display an intensified binding of the divalent 
cations over a chain length of 20 monomers (Sime, 1990, Smidsr?d and Skjaok-Brk, 
1990). This indicates that gelation mechanism occurs in a cooperative manner in the 
gelation process in which binding sites are already arranged in an ordered array and 
binding of one ion helps to bind the next. These impacts are not seen for poly-D-
mannuronic acid or alternating sequences (Morris et al., 1973, Sime, 1990). 
Each alginate chain dimerizes to form joints with many other chains and 
therefore gel networks are formed (Dupuy et al., 1994, George and Abraham, 2006). So 
the calcium reactivity of alignment is the result of calcium-induced dimeric association of 
the G-block regions (George and Abraham, 2006). These hydrogels that are similar to 
solids in maintaining their forms and resisting stress possess 99?99.5% water with rest 
being alginate (George and Abraham, 2006). Upon the amount of calcium involved in 
the system, these inter-chain connections can be either impermanent or permanent 
(George and Abraham, 2006). Impermanent connections are acquired by low amounts 
of calcium, giving rise to highly viscous, thixotropic solutions (George and Abraham, 
2006). Precipitation or gelation results from permanent connections of the chains at 
higher calcium amounts (George and Abraham, 2006). A great deal of research has 
proved that the chemical composition, molecular size, the hydrogel forming kinetics and 
the cation have a significant impact on some of its functional properties including 
porosity, swelling behavior, stability, biodegradability, gel strength, and the gel's 
immunological characteristics and biocompatibility (George and Abraham, 2006).  
 Alginate and its gels have been used in numerous and markedly different 
applications. It has historically been most commonly exerted in the food industry (Augst 
14 
 
et al., 2006). It has also been utilized as a stabilizer and an emulsifier, e.g., in low-fat fat 
substitutes, on account of its intrinsic properties and the interactions of alginate with 
proteins, fats, or fibers (Augst et al., 2006). Alginate is also found utility as a gelling 
agent for food products, since alginate?pectin mixtures gel independently of sugar 
content, and its use can bear low-calorie products (Augst et al., 2006). Alginate is 
chemically modified with propylene glycol to allow its application under acidic conditions 
(Draget et al., 2005, Glicksman, 1987), and to bind flaked, chunked, or milled foodstuffs. 
It is also a shear-thinning agent in the textile/paper-printing industry due to its high hue 
yield and print levelness (Glicksman, 1987, McLachlan, 1985). In the pharmaceutical 
industry, it is widely applied as an excipient for drugs (Liew et al., 2006), a dental 
impression material (Ashley et al., 2005), and a wound dressing (Matthew et al., 1995). 
As a biomaterial, alginate possesses many advantageous features such as 
biocompatibility and nonimmunogenicity. They are possibly because of its hydrophilicity 
(Shapiro and Cohen, 1997). A significant advantage is its easy gelling behavior, which 
induces entrapping of various substances with minimum trauma (Klock et al., 1997). A 
chemically modified alginate is currently used clinically as a drug delivery vehicle for 
proteins that promote regeneration of mineralized tissue (Bratthall et al., 2001) and as a 
carrier for transplanted cells (Bent et al., 2001). One of its disadvantages is that 
enzymes cannot naturally break it down in mammals and hence has poorly controlled 
degradation (Lansdown and Payne, 1994). In addition to those, cells do not naturally 
adhere to alginate (Smetana Jr, 1993). There has been impressive research over the 
past decade to handle these problems and to widen the utility of alginate hydrogels 
(Augst et al., 2006). 
15 
 
1.1.4.2 Chitosan 
Chitosan is a positively-charged polysaccharide that is extracted from chitin 
which is the main structural polymer in arthropod exoskeletons (Muzzarelli, 1977, 
Muzzarelli et al., 1986). Since its molecular weight is relatively high, between 300-1000 
kDa, depending on the source of chitin and extraction method, applications composed 
of this polymer have high tensile modulus and thus can be used in a wide array of 
application (Dvir et al., 2005). 
Chitin is the second most abundant polysaccharide in nature, exceeded only by 
cellulose. This renewable resource is found in numerous naturally occurring organisms 
such as fungi, yeast, algae and is the principal component in the exoskeleton of sea 
crustaceans (Figure 1.7), such as shrimp and crab as well as in the cuticle and 
ovipositors of insects (Hudson and Jenkins, 2002). 
  
 
Figure 1. 7 Crustaceans 
 
Chitin possesses a homogeneous chemical composition composed of 1-4 linked 
2-acetamido-2-deoxy-?-D-glucopyranose (Figure 1.8). 
16 
 
 
Figure 1. 8 Chemical structure of chitin 
 
While chitin is naturally available in great amounts through many sources, 
chitosan is only accessible in nature in limited quantities, such as in some fungi: mucor 
rouxi, aspergillis nidulans (Hudson and Jenkins, 2002). The chitosan used in industrial 
and research applications is generally extracted from chitin by the use of chemical or 
enzymatic treatments of the shells of Arthropoda (shrimp or crab) secured from the 
waste products of the crabbing and shrimping industries. Several chemical, mechanical, 
and enzymatic methods have been developed to acquire purified chitosan. The 
common method to extract chitosan is the chemical treatment of shell fish wastes from 
the crab and shrimp industries. First, the shell is demineralized in dilute hydrochloric 
acid at room temperature. Second, it is deprotenized in warm dilute caustic. This forms 
a partially deacetylated chitin which finally is further deacetylated to chitosan. Through 
deacetylation process of chitin, chitosan with 70% to 95% deacetylation degree is 
obtained (Hudson and Jenkins, 2002). 
Chitosan, a partially deacetylated derivative of chitin, has at least 50% of the free 
amine groups. Therefore, it has a varying chemical structure consisting of both 1-4 
17 
 
linked 2-acetamido-2-deoxy-?-D-glucopyranose as well as 2-amino-2-deoxy-?-D-
glucopyranose (Figure 1.9).  
 
 
Figure 1. 9 Chemical structure chitosan 
 
As the process conditions control the extent of deacetylation that occurs, the 
properties of chitosan are mainly affected by the conditions under which it is processed. 
The degree of deacetylation (DD) sets the amount of free amino groups in the polymer 
backbone. The free amino groups provide the positive charge to chitosan. The 
functionality of chitosan which allows it to be a highly reactive polysaccharide is 
provided by the amino group and the hydroxyl groups together. The positive charge of 
chitosan enables it to make a great number of electrostatic interactions with negatively 
charged molecules. The extent of functional groups generated by deacetylation and the 
processing conditions support side group attachment, which then affects crystallinity 
that directly links to chitosan?s ability to solubilize in acidic aqueous solutions, an 
important phenomena of the processability of chitosan (Hudson and Jenkins, 2002, 
S?enel and McClureb, 2004). 
18 
 
Some biological properties including biodegradability, lack of toxicity, anti-fungal 
effects, wound healing acceleration, and immune system stimulation of chitosan make it 
promising material for the needs in medical applications (Di Martino et al., 2005, 
Hudson and Jenkins, 2002, Khor and Lim, 2003, S?enel and McClureb, 2004). The 
biological and chemical properties of chitosan provide the potential to make bonds with 
particular materials such as cholesterols, fats, proteins, metal ions, and even tumor 
cells. Thus enables chitosan to be utilized as a chelating agent in many applications (Li 
et al., 1992). 
The degradation of chitosan can be accomplished through several methods. 
Owing to its degradation rate that is inversely proportional to the degree of crystallinity 
and consequently the amount of deacetylation, degradation can be engineered by 
controlling the amount of deacetylation that takes place during processing. Chitosan can 
be thermally degraded at temperatures above 280?C and also polymer chains 
decompose fast. Enzymatic degradation is an important means of handling the 
decomposition of chitosan. A wide range of hydrolytic enzymes, such as lysozyme (the 
primary enzyme responsible for degradation of chitosan), encountered in the lyphoid 
systems of humans and animals, can be exerted to naturally degrade chitosan (Di 
Martino et al., 2005, Hudson and Jenkins, 2002). The degradation of chitosan within the 
body leads to the release of amino-sugars which can be easily processed and 
metabolize through the organic system. Degradation is a substantial property to 
comprehend so that the processing and the end applications can be engineered 
accordingly (S?enel and McClureb, 2004). 
19 
 
Due to chitosan?s many attractive properties such as biodegradability, natural 
origin, abundance, reactivity, it has many areas of application including: medical, 
agricultural, food processing, nutritional enhancement, cosmetics, and waste and water 
treatment (Hudson and Jenkins, 2002, S?enel and McClureb, 2004). 
The biodegradability, profusion, lack of toxicity, and natural origin of chitosan 
make it used reliably in agricultural applications without concerns of pollution, 
discarding, or negative effect to consumers if eaten (Li et al., 1992). Some of the 
applications within agricultural use are seed coating, leaf coating, fertilizer, and time 
released drug or fertilizer responses where chitosan has found utility (Li et al., 1992). 
The applications in these areas include raising the harvest by enhancing germination, 
rooting, leaf growth, seed yield, and soil moisture retention, while decreasing the 
occurrence of fungal infections and diseases (Li et al., 1992). 
The functional groups and natural chelating properties of chitosan are utilized 
extensively in wastewater treatment by inducing for the binding and removal of metal 
ions such as copper, lead, mercury, and uranium from wastewater (Hudson and 
Jenkins, 2002). The breakdown of food particles that contain protein and removal of 
dyes and other negatively charged solids from wastewater streams and processing 
outlets are the other applications for chitosan (Li et al., 1992). 
Several valuable applications within the food industry such as binding with and 
removing certain elements, particles, and materials such as dyes and fats from foods 
come from its chelating properties and high functionality (Li et al., 1992). The 
antimicrobial and antifungal properties occurring in chitosan make it efficient during the 
storage and preservation of food (Hudson and Jenkins, 2002, Li et al., 1992). 
20 
 
Because chitosan functions in numerous shapes, there is a great deal of use and 
interest within the medical industry for uses such as drug delivery, wound healing, 
orthopedic, periodontal, and tissue engineering (Khor and Lim, 2003). It has been 
greatly modified and used for drug delivery systems in oral, parenteral, transdermal, 
ocular, and nasal applications especially due to its low toxicity with outstanding 
mucoadhesion properties (Paul and Sharma, 2000). Chitosan gels and films displayed 
sustained release of different drugs efficient for wound dressings and colon treatments 
(Hudson and Jenkins, 2002). Wound dressing of chitosan can stimulate some various 
physiological activities (Muzzarelli, 1997). On the wound surface, they induce 
macrophage activity to discharge lysozyme and human chitinase, enzymes which 
separates chitin-based pathogens to prevent infection (Hudson and Jenkins, 2002). 
Because of chitosan?s excellent fiber and film forming ability, it has been blended with 
other polymers to control permeability and to improve blood compatibility in membrane 
applications for hemodialysis patients (Rathke and Hudson, 1994).  
 
 
 
 
 
 
 
 
 
21 
 
1.2 RESEARCH OBJECTIVES 
There are two main objectives associated with this research: 
The first objective of this work is to understand and evaluate the problems 
associated with the composition of chitosan and alginate. Different forms of the 
composition will be evaluated and its quality will be assessed as possible biomaterial 
applications for different areas. 
The second objective is to obtain a basic understanding of the thermal, 
mechanical, morphological, and antibacterial properties of the each component as well 
as their compositions. 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
22 
 
 
  
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25 
 
 
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26 
 
 
 
CHAPTER TWO 
CHARACTERIZATION OF CHITOSAN-ALGINATE FILMS 
 
2.1 INTRODUCTION 
Recently, the growing interest in the development and the application of new 
materials from renewable sources shows a trend away from synthetic polymers and 
towards natural polymers in different areas such as food science, medicine, and 
cosmetics (Neto et al., 2005). This tendency has certainly occurred in food packaging 
due to the growing public health concern, the reducing environmental impact, safety, 
and convenience (Pereda et al., 2008). The biodegradability and biocompatibility are the 
main functional advantages of natural polymers in food packaging. An advantageous 
fact of the biodegradable packaging materials is that through biodegradation and 
decomposition, they could be also used in secondary applications such as fertilizer or 
soil conditioner, providing a better harvest. Even though it is slightly expensive, 
biodegradable packaging is the premise of user friendly and eco-friendly packaging 
(Srinivasa and Tharanathan, 2007). 
 Among various food packaging materials, polysaccharide based films have 
drawn considerable attention. Different types of polysaccharides and their derivatives 
have been utilized as biodegradable film-forming matrixes (Rhim and Ng, 2007). Along 
the process of polysaccharide film formation, the separation of polymer segments and 
27 
 
reorganization of the polymer chain into a film matrix or gel occur through the 
evaporation of a solvent that generates hydrophilic and hydrogen bonding (Rhim and 
Ng, 2007). 
 Chitosan, one of the commonly used polysaccharides, is a cationic biopolymer 
that has unique nutritional, biomedical, and functional properties (Takahashi et al., 
1990, Pereda et al., 2008). It has been used in many application areas, such as 
agriculture, waste water treatment, food industry, and biomedicine. The commercially 
available form of chitosan is mostly the deacetylated chitin, which is the second 
abundant polysaccharide on earth after cellulose and is extracted from waste products 
of the shellfish industry (Xu et al., 2005).  
 
 
Figure 2. 1 Structure of chitosan 
 
Unlike chitin, there are reactive amino and hydroxyl groups on chitosan 
backbone that can be used for chemical reactions and salt formation (Figure 2.1). 
These groups, due to their hydrophilicity, have an important impact on permeability and 
diffusion of water through chitosan films (Meng et al., 2009). From earlier studies, 
28 
 
chitosan has been recognized to carry effective film forming properties, chemical 
resistance, and high permselectivity for water (Meng et al., 2009). The neutralization of -
NH3+ sites into -NH2 in chitosan caused to the vanishing of the ionic repulsions between 
polymer chains; therefore, physically cross-linked formation related to hydrogen 
bonding, hydrophobic interactions and crystallite formation were preferred (Meng et al., 
2009). This property has been applied on edible films by forming electrostatic 
interactions with anionic groups in an acid environment (Xu et al., 2005). 
 Alginate is the main structural component in marine brown algae. The 
intercellular alginate gel matrix provides plants both tenacity and resilience (Draget et 
al., 2009). It is known as potential biopolymer film or coating material because it has 
unique and well examined colloidal properties, such as thickening, stabilizing, 
suspending, and film and gel forming properties (Zactiti and Kieckbusch, 2006). It has 
been widely used in industry including uses in agriculture, medicine, pharmacy, food 
processing, and construction. As extracted from brown algae, alginate is a water-soluble 
linear polysaccharide and consists of alternating blocks of 1?4 linked ?-L-guluronic (G) 
and ?-D-mannuronic (M) acid residues (Figure 2.2) (George and Abraham, 2006). The 
order and the chemical configuration of the M and G blocks are dependent on the 
biological source, growth and seasonal conditions (George and Abraham, 2006). 
 
29 
 
 
Figure 2. 2 Structure of alginate 
 
Several studies on chitosan based films have been reported, such as chitosan-
starch (Lazaridou and Biliaderis, 2002, Xu et al., 2005, Zhai et al., 2004), chitosan-
pullulan (Lazaridou and Biliaderis, 2002), chitosan-gelatin (Arvanitoyannis et al., 1998, 
Chatterjee et al., 1987, Kolodziejska and Piotrowska, 2007), chitosan-PVA 
(Arvanitoyannis et al., 1997), and chitosan-amylose starch (Cervera et al., 2004). 
However, according to our literature review, studies on chitosan-alginate films are rarely 
found. By processing these films, advantages might be found from the electrostatic 
interactions through the -NH3+ groups of chitosan, as a cationic polymer, with anionic 
groups, such as -COO- groups, present in alginate or polyanions (Pereda et al., 2008). 
These interactions illustrated in Figure 2.3 play an important role in the formation of 
polymeric complexes (Pereda et al., 2008). 
30 
 
 
Figure 2. 3 Structural scheme of chitosan?sodium alginate ion complex formation 
reaction between anion group (?COO-) of sodium alginate and the protonated cation 
group (?NH3+) of chitosan 
 
 Chitosan and sodium alginate are both natural polymers, and both have 
biocompatibility, biodegradability, and hydrophilicity. Therefore, chitosan-sodium 
alginate polymer mixtures have the potential to produce new functional polymers with 
31 
 
good performance, which can be used in various applications. In this chapter, chitosan-
alginate polymer films were prepared and miscibility, thermal stability, mechanical 
morphological properties of the films were studied by infrared (FTIR) spectra, scanning 
electron microscopy (SEM), thermogravimetric analyzer (TGA), differential scanning 
calorimeter (DSC) and dynamic mechanical analyzer (DMA). In addition to these 
analyses, antibacterial properties of the films were tested through zone inhibition test. 
 
2.2 EXPERIMENTAL 
2.2.1 Materials 
Commercial medium molecular weight (MMW) chitosan (average molecular 
weight (Mw) = 400,000 Da and deacetylation degree (DD) 80%) was supplied by 
Sigma-Aldrich Corp., St. Louis, MO. Commercial sodium alginate and glacial acetic acid 
(CH3COOH) were purchased from Fisher Scientific, Pittsburgh, PA.  For nuclear 
magnetic resonance test, deuterated acetic acid (CD3COOD) and deuterium oxide 
(D2O) were also purchased from Fisher Scientific. For zeta potential measurements, 
sodium hydroxide (NaOH) and hydrochloric acid (HCl) were purchased from Sigma-
Aldrich Corp., St. Louis, MO. 
2.2.2 Methods and Techniques 
2.2.2.1 Nuclear magnetic resonance (NMR) spectroscopy 
The molecular structure and properties of chitosan is extensively dependent 
upon the degree of deacetylation (Hirai et al., 1991). As a result, it was essential to 
determine precisely the degree of deacetylation. For this determination, two methods 
have been reported in literature besides NMR spectroscopy: (i) Colloid titration 
32 
 
evaluating amino residue (Terayama, 1952), and (ii) Elemental analysis and 
transmission ratio of infrared spectrum evaluating carbon/nitrogen ratio (Miya et al., 
1980, Sannan et al., 1978). The effect of high moisture absorbency of chitosan cannot 
be completely removed with these methods above and also a great amount of sample is 
to be consumed in the case of colloid titration; therefore, 1H NMR spectroscopy is 
selected. In 1H NMR, small amount of sample (~5mg) is needed and water sorption can 
be eliminated (Hirai et al., 1991). 
 1H NMR spectra of chitosan was performed on a Bruker AV-400 spectrometer, 
operating at 400 MHz. Medium molecular weight chitosan was dissolved in 2 wt% 
CD3COOD/D2O solution at room conditions. After dissolution, approximately 1 ml of the 
chitosan solution was transferred to a 5 mm NMR tube. The sample tube was inserted 
in the magnet and allowed to reach thermal equilibrium by waiting 10 minutes before 
performing the experiment. Integration boundaries of obtained spectrum were adjusted 
manually by the analyst?s judgment. 1H chemical shifts were expressed in ppm 
downfield from the signal for sodium 3-(trimethylsilyl) propane sulfonate (TSP) as an 
external reference. 
2.2.2.2 Zeta (? ) potential measurements 
Zeta potential measurements of chitosan, alginate and their mixture solutions 
were performed with a Particle Sizer Analyzer NICOMP at 25?C. Measurements were 
performed as a function of pH in buffered solutions. The measurements were carried 
out in triplicate. All the values were measured with a relative accuracy of 1%. 
33 
 
2.2.2.3 Film preparation 
Solution casting is the chosen method for the preparation of films. Chitosan 
solutions (1% w/v) were prepared by adding chitosan powder in acetic acid solution (1% 
v/v). Sodium alginate solution (1% w/v) was prepared by dissolving alginate powder in 
distilled water. Both solutions were filtered through a 325 mesh stainless steel woven 
disk filter. 
Different proportions of chitosan and sodium alginate solutions (100:0, 20:80, 
40:60, 50:50, 60:40, 80:20, 0:100 CH:AL) were prepared by manual stirring, then 
poured into glass Petri dishes (? = 9 cm) and finally kept in the laboratory condition 
(21?C, 50% RH) for 24 hours to remove air bubbles. After that the films were dried at 
constant temperature (30?C) overnight in a convection oven, resulting in films with 
thicknesses between 45 and 55 ?m. After the drying stage, the films were peeled off 
from the glass plates and kept in a desiccator at ambient conditions. 
 
Table 2. 1 Chitosan-alginate film compositions 
 Volume (ml) 
Chitosan-
Alginate       
(CH:AL) (%) 
Chitosan   
(1.0 % w/v) 
Alginate    
(1.0 % w/v) 
100:0 40 0 
80:20 32 8 
60:40 24 16 
50:50 20 20 
40:60 16 24 
20:80 8 32 
0:100 0 40 
34 
 
 
Figure 2. 4 Illustration of film preparation process 
 
2.2.2.4 Light transmission properties 
Light transmission tests were performed in a UV-VIS Spectrophotometer (UV-
2450 Shimadzu Scientific) under wavelength 400~750 nm. Each dry film (3cm?1cm) 
was fixed onto the wall of the sample holder. Film transparency was determined by 
measuring the amount of light transmitted through the films relative to air. 
2.2.2.5 Infrared spectroscopy characterization 
Infrared spectrums of the films were measured by attenuated total reflection 
method (ATR) using a Thermo Nicolet 6700 Fourier transform infrared spectrometer 
connected to a PC with OMNIC software analysis.  The thickness of the films used was 
35 
 
about 45-55 ?m. All spectrums were recorded between 400 and 4000 cm-1 over 32 
scans with a resolution of 4 cm-1. 
2.2.2.6 Thermogravimetric analysis (TGA) 
TGA thermograms of the films were measured on a Thermogravimetric Analysis 
(TA) Instrument Q500. All analyses were carried out with ~5 mg samples in platinum 
pans under a dynamic nitrogen atmosphere between 25?500?C at a heating rate of 
10?C/min under nitrogen purge. 
2.2.2.7 Differential scanning calorimetry (DSC) 
Differential scanning calorimetric test was performed using a Differential 
Scanning Calorimeter TA Instruments DSC 2920. Samples (~5 mg) were placed in 
aluminum pans with lids. The test included two runs; first run from -50 to 180?C and the 
second one from -50 to 280?C with a scanning rate of 10?C/min. 
2.2.2.8 Dynamical mechanical analysis (DMA) 
DMA measurements were performed on a Dynamic Mechanical Analyzer (DMA 
RSA3 TA Instruments) at 1 Hz and a heating rate of 2?C/min. DMA curves were 
obtained in two cycles of heating run: the first one of heating up to 110?C then holding 
there for 10 minutes followed by cooling to 30?C and the second one of heating up to 
250?C. The samples were cut 20.0 mm long, 5 mm wide and about 0.04 mm thick. The 
samples were not clamped during the first heating and subsequent cooling process in 
order to avoid residual stress.  
36 
 
2.2.2.9 Scanning electronic microscopy analyses 
The fractured surfaces of films pre-chilled in liquid nitrogen were studied by 
scanning electron microscope (SEM, Cambridge 360). Samples were sputtered with 
gold prior to SEM observation. 
2.2.2.10 Mechanical properties 
During this research, tensile tests were performed in order to evaluate the 
mechanical response of films. 
Tensile tests: Film thicknesses were measured using a 0?25 mm automatic 
micrometer, with a resolution of 0.01 mm. The reported values are the average of 10 
readings taken randomly on each film sample. Tensile tests were performed according 
to the ASTM D1708-93 (1993) with a Dynamic Mechanical Analyzer (DMA RSA3 TA 
Instruments) in multiple extension mode at room temperature (23?C ? 2?C). A film strip 
(dimensions 2.0 cm?0.5 cm) was clamped between grips and stretched by top grip at 
rate of 0.05 mm/sec. The yield strength (?y), elongation at break (?b) and elastic 
modulus (E) were calculated as described in ASTM D638-94b (1994). 
2.2.2.11 Antibacterial testing - Zone of inhibition test 
In vitro antibacterial behavior of the 100:0, 60:40, 80:20, 0:100 CH:AL films were 
evaluated by inhibition zone experiments against Gram-negative bacterium (Escherichia 
coli). The films were cut into 2x2 cm2 and then placed on nutrient agar plates coated 
with the bacteria with a concentration of approximately 105 CFU/ml. Finally, the plates 
were incubated at 37?C for 24 hours, and the inhibition zone was measured. 
 
 
37 
 
2.3 RESULT AND DISCUSSION 
2.3.1 NMR study for the determination of the degree of deacetylation 
The 1H-NMR spectrum of chitosan is shown in Figure 2.5. HOD band appeared 
due to dissolution of chitosan in 2 wt% CD3COOD/D2O and its proton which is not 
bonded to chitosan resonates at 4.62 ppm. All other chemical shifts of chitosan protons, 
also similarly reported by Hirai et al. (1991) and Varum et al. (1991), are shown in Table 
2.2.   
 
 
Figure 2. 5 1H NMR spectrum of chitosan (DD = 80%), in CD3COOD/D2O, at 400 MHz   
 
 
38 
 
Table 2. 2 Chitosan chemical shifts of proton at room conditions in CD3COOD/D2O 
Proton Chemical Shift (ppm) 
H1 5.36 
H2 2.95 
H3, 4, 5, 6, 6' 3.73, 3.69, 3.56, 3.5 
N-acetyl 1.85 
 
Deacetylation degree (DD) of chitosan is calculated from Equation (1) (Hirai et 
al., 1991) by using the integral intensity,     , of this CH3 residue, and the total of 
integral intensities,       , of H2, H3, H4, H5, H6, and H6' protons. 
    ( )  *  (               )+     Equation (1)    
Through the NMR spectra of chitosan (           ,               )  the 
deacetylation degree (DD) was calculated 79% which is almost the same degree as the 
provider?s claim, ~80%. 
2.3.2 Zeta (? ) potential measurements 
Zeta potential of 0.1 w/v% chitosan and 0.1 w/v% alginate solutions and their 
mixture of 50:50 with the same concentrations were measured with 0.1 pH unit of a 
chosen value with 0.1 or 1 M NaOH or HCl solution. The results are shown in Figure 
2.6.  
 
39 
 
 
Figure 2. 6 Zeta potential of the solutions of chitosan, alginate and their mixture 
 
Chitosan begins precipitating above pH 6.6 and its zeta potential declines to zero 
at pH 7?0.1 which is its isoelectric point indicating no net charge. Below its precipitation 
point, chitosan has high positive zeta potential confirming the cationicity of the polymer. 
However, alginate dissolved in water carries negative charge at pH 7?0.1. Below that 
point, the net charge of alginate solution decreases and reaches zero at pH 3.0?0.1. 
The 50:50 solution of chitosan and alginate has a negative charge above pH 4.3 
indicating the excess negative charge on account of alginate. In acidic medium, the 
mixture solution carries almost no net charge which most likely occurred due to 
electrostatic interaction between cationic chitosan and anionic alginate. 
2.3.3 Light transmission properties 
The transparency of prepared samples is reported in Figure 2.7. The light 
transmission of all the films increases with the wavelength. 
40 
 
 
Figure 2. 7 Light transmission of different composition of films 
 
In the range of visible light spectrum (400 ? 750 nm), chitosan (100:0 CH:AL) film 
exhibited the best light transmission and alginate (0:100 CH:AL) film displayed a very 
close degree of light transmission to chitosan. This occurred due to the intrinsic 
yellowish hue of alginate. However, the films prepared by means of different 
composition (80:20, 60:40, 50:50, 40:60 CH:AL) exhibited lower light transmission 
property than pure films. Possible ionic complexation between chitosan and alginate 
induced an impact on the transparency of the composite films, thus lessened their light 
transmission ratio. In addition, 80:20 CH:AL film exhibited the worst property due to the 
precipitation of the alginate. Nevertheless, except 80:20 CH:AL film, the light 
transmission ratio of all films is still above 86%, indicating high transparency. 
41 
 
 It can be also clearly observed from Figure 2.8 that the 50:50 CH:AL film is highly  
transparent, which is already confirmed by light transmission measurements. 
 
 
Figure 2. 8 50:50 CH:AL film 
 
2.3.4 Infrared spectroscopy 
FTIR spectra of 60:40 CH:AL film and pure samples are observed in Figure 2.9. 
The characteristic absorption bands at 1637 and 1542 cm-1 represent the amide I and 
amide II band respectively on chitosan films (Table 2.3) (Lawrie et al., 2007). For 
alginate, the absorption bands at 1592 cm-1 and 1407 cm-1 represent the anti-symmetric 
and symmetric stretching vibrations of -COO- (Table 2.3). Lawrie et al. (2007) also 
reported similar results on the absorption bands of chitosan and alginate. 
42 
 
 
Figure 2. 9 FTIR spectra of chitosan (100:0), alginate (0:100) and 60:40 CH:AL films 
Table 2. 3 FTIR bands of chitosan, alginate and 60:40 CH:AL films with assignments 
100:0 CH:AL      
Vibration (cm-1) Assignment 
3350-3000 O-H and N-H stretch 
2875 C-H stretch 
1637 Amide I 
1542 N-H bending from amine and amide II 
1405 -CH2 bending 
1376 CH3 symmetrical deformation 
1151 Antisymmetric C-O-C stretch and C-N stretch 
1064-1018 Skeletal vibration of C-O stretching 
43 
 
0:100 CH:AL      
Vibration (cm-1) Assignment 
3600-3000 (broad) O-H stretch 
2930 C-H stretch 
1592 Antisymmetric COO- stretch 
1407 Symmetric COO- stretch 
1295 Skeletal vibration 
1081-1024 Antisymmetric C-O-C stretch 
60:40 CH:AL      
Vibration (cm-1)  Assignment 
3500-3000 (broad)  O-H and N-H stretch  
2923 C-H stretch  
1597 Antisym COO- stretch  
1540 N-H bending from amide II 
1400 Sym COO- stretch or -CH2 bending 
1305 skeletal vibration  
1149 Antisym C-O-C stretch and C-N stretch 
1064-1024 Skeletal vibration of C-O stretching  
 
The absorption bands of 60:40 CH-AL film reveal similar characteristics with both 
chitosan (100:0 CH:AL) and alginate (0:100 CH:AL) films. The O-H and -NH2 groups in 
chitosan may form hydrogen bonds with -C=O and -OH groups of alginate. The 
absorption band at 1542 cm-1 in chitosan shifted to 1540 cm-1 and 1592 cm-1 in alginate 
shifted to 1590 cm-1 after alginate reacted with -NH2 groups via hydrogen bonds (Table 
2.3). Due to the same probable bonding, two similar peaks, one in chitosan at 1405 cm-1 
44 
 
and the other in alginate at 1407 cm-1 overlap each other in the 60:40 CH:AL sample 
(Table 2.3). The composite film also shows characteristic absorption bands between 
3000 cm-1 and 3600 cm-1, which represent the -OH and -NH2 groups in free as well as in 
amide form in chitosan (Meng et al., 2009). Additionally, ionic interactions most likely 
occurred between chitosan and alginate. The cationic       group in chitosan and 
anionic       group in alginate are capable of forming complexes with opposite 
charges. Therefore, these intermolecular interactions result in forming polyelectrolyte 
complex (Meng et al., 2009). 
2.3.5 Thermal analysis 
Figure 2.10 and 2.11 show the TGA and DSC thermograms of 50:50, 100:0, 
0:100 CH:AL films. For all samples, weight loss took place in two stages. The first one 
starts at ~50?C with a weight loss of 10% and reaches a maximum above 100?C. This 
stage is assigned to water evaporation. 
45 
 
 
Figure 2. 10 TGA thermograms of chitosan (100:0 CH:AL), alginate (0:100 CH:AL) and 
50:50 CH:AL films 
 
The second stage for chitosan occurs at 300?C with a weight loss of above 40%. 
This result is similar to the findings of Nieto et al. (1991) and Tirkistani (1998). For 
alginate, this stage takes place at 250?C with a weight loss of ~35%. ?aykara et al. 
(2005) and Parhi et al. (2006) reported similar results for alginate. This stage 
corresponds to the decomposition (thermal and oxidative) of chitosan and alginate, 
vaporization and elimination of volatile products. According to Nieto et al. (1991), 
pyrolysis of polysaccharides starts by a random split of the glycosidic bonds, followed 
by a further decomposition forming acetic and butyric acids and a series of lower fatty 
acids, where C2, C3 and C6 predominate. 
 
46 
 
 
 
Figure 2. 11 DSC thermograms of chitosan (100:0 CH:AL), alginate (0:100 CH:AL), and 
50:50 CH:AL films. a) first run b) second run 
47 
 
The composite (50:50 CH:AL) film shows the characteristic of both chitosan and 
alginate films. The second stage for this film starts at 204?C and reaches a maximum at 
380?C. It undergoes a broad range of decomposition due to its components. Therefore, 
this binary polysaccharide includes the thermal history of chitosan and alginate and its 
decomposition stage can be attributed to the same explanation expressed by Nieto et 
al. (1991). 
The first heating (Figure 2.11(a)) process on DSC thermogram shows a large 
endothermic peak above 100?C indicating solvent evaporation (mainly water) of all 
films. This is also in agreement with the TGA results. The second heating (Figure 
2.11(b)) process was applied after all solvent traces were eliminated. Its thermogram 
shows degradation of all films above 200?C. For example, the 50:50 mixture film starts 
degrading around 204?C which is also the same temperature found in TGA results.  
Because no glass transition is observed on calorimetric data dynamic mechanical 
analysis (DMA) was performed for films. DMA tests were performed using two heating 
processes. The first heating was applied in order to eliminate the effect of residual 
water. After holding the samples at 110?C for ten minutes, no color change observed. 
However, all samples shrunk almost 10% in every dimension. The shrinkage can be 
attributed to the [10%] weight loss observed on TGA. After the first heating, second 
heating was applied with 0.01% strain. Figure 2.12 displays the dynamic mechanical 
spectra on storage modulus, E?, and tan ? of samples, acquired from the DMA 
measurement during the second heating process.  
 
48 
 
                                    
 
Figure 2. 12 Dynamic temperature ramp test of films: a) E? (storage modulus), b) tan ? 
versus temperature 
49 
 
A sharp peak at 155?C is observed on tan ? or curve deflection on E? for pure 
chitosan (100:0 CH:AL) sample during the second heating which corresponds to the 
relaxation process of the chitosan network. Similar results were reported by Mucha and 
Pawlak (2005). Furthermore, no related transition occurred on pure alginate. Although 
chitosan and alginate have similar structures based on glycosidic bonds, alginate did 
not exhibit any transition other than molecular degradation after 195?C. Mitchell (1998) 
explains this with chain mobility which in biopolymers occurs less often than most 
synthetic polymers. Two reasons stand out for this in polysaccharides. First, steric 
hindrances intensively affect the open angles of rotation on the glycosidic bond, the 
geometry of which mostly provides that restriction. Second, some intra chain attractive 
interactions, particularly hydrogen bonding, will also limit the mobility. Because glass 
transition depends on chain mobility, a clear transition less likely occurs in immobilized 
polymers through other mechanisms (Mitchell, 1998). 
It can be clearly noticed that there is a tendency on both graphs at 155?C through 
the rise of chitosan content in each composite film. The peak becomes much clearer as 
the percentage of the chitosan in composite films increases. Therefore, composite films 
exhibit a transition due to the chitosan segment. 
It is remarkable that the storage moduli (E?) of all composite films increased 
around one order of magnitude compared to pure films. This most likely occurred 
though stronger interaction between chitosan and alginate after the solvent evaporation. 
When water induced hydrogen bonding replaced with more ionic interaction between 
the two polymers, the binary polysaccharide network became more compact and more 
binding were obtained.  
50 
 
2.3.6 Morphology of the films 
The surface morphology of the chitosan alginate composite films was observed 
with scanning electron microscopy to verify the compatibility of the mixtures of alginate 
and chitosan. Figure 2.13 shows the fractured surface of the pure and composite films.  
The cross section structure of alginate film (0:100 CH:AL) is very rough 
compared to that of chitosan film (Figure 2.13(a, b) and 2.13(e, f)). This difference can 
also be observed in the composite film (Figure 2.13(c, d)) which two biopolymers 
formed mainly a two-phase system. Although the presence of phase separation at a 
microscopic level was detected for the composite films, pronounced interactions 
between two components can be confirmed by their solubility. After the composite films 
are dried, they cannot be dissolved completely in water again because of the polyionic 
complexation. 
 
 
51 
 
 
 
Figure 2. 13 SEM pictures of a, b) 100:0 CH:AL, c, d) 50:50 CH:AL and e, f) 0:100 
CH:AL films 
 
It is well-known that a better mixture could be achieved if the concentration of the 
each solution is reduced accordingly. However, in our study, it was observed that lower 
than 1.0 w/v% concentrations of either of the component solutions creates precipitation 
in one another. Consequently, the resulting films cast from the mixed solutions will 
possess a macroscopically phase separated structure and exhibit an immiscible 
appearance.  
Even though phase separation and layered structure were observed on our 
composite films, the transparency was not affected. This is a result of the refractive 
52 
 
index of both phases which are polysaccharides. They (nChitosan= 1.59, nAlginate= 1.507) 
have similar refractive indexes (Ligler et al., 2001, Cathell and Schauer, 2007). 
2.3.7 Mechanical properties 
In this study, aqueous acetic solution was used as a casting solution. Relatively 
warm air (30?C) evaporates water from the outermost region of the initial mixture to 
induce film to dry and finally form a dense uniform skin. As is also shown in Figure 2.8, 
the 50:50 CH:AL film exhibits a miscible appearance. However, the microscopic 
evidence revealed phase separation of the two polymer networks, with phase inversion 
occurring at each composition. As a result, all films prepared are symmetric and layer-
by-layer phase separated except the 80:20 CH:AL film in which alginate precipitates 
due to lack of interaction with chitosan.  
 The Young?s modulus, deformation at break, yield strength and stress at break of 
the chitosan-alginate composite films with various chitosan content were tested 
according to the ASTM D638-94b (1994) and the results are reported in Figure 2.14 and 
Table 2.4. 
 
53 
 
 
 
54 
 
 
 
Figure 2. 14 Mechanical properties of chitosan-alginate films: a) Modulus, b) Yield 
strength, c) Deformation at break, d) Stress at break 
55 
 
 All the composite films showed similar mechanical behaviors but the one with 
chitosan content of 80% due to the precipitation of alginate. The mechanical results of 
this film on all graphs excluded as an exception to the composite films produced. 
The stress at break, yield strength and Young?s modulus of composite films 
reach a maximum at 60% chitosan content indicating the greatest ionic interaction 
among all composite films. Because tensile tests were conducted without evaporating 
water content of the films, we could not observe any increase on the modulus of 
composite films as we did in their storage moduli. Although phase separation was 
observed, composite films possess hydrogen bonding and ionic interaction through the 
interface of each component. As a result, the toughest composition was obtained in 
60:40 CH:AL film which has nearly the same modulus as chitosan (100:0 CH:AL) and 
alginate (0:100 CH:AL) films. 
 
Table 2. 4 Mechanical properties of chitosan-alginate films 
Films        
CH-AL 
Young's 
Modulus (MPa) 
Deformation                 
at Break (%)  
Yield Strength 
(MPa)  
Stress at 
Break (MPa)  
0:100 3433 ? 302 10.17 ? 2.85 54.93 ? 8.57 83.53 ? 11.07 
20:80 3007 ? 236 10.26 ? 2.04 43.89 ? 3.41 72.64 ? 7.41 
40:60 2323 ? 330 17.0 ? 4.18 36.14 ? 5.37 64.84 ? 12.28 
50:50 2800 ? 210 13.35 ? 2.48 44.23 ? 2.75 67.30 ? 7.25 
60:40 3432 ? 423 10.47 ? 3.24 59.29 ? 7.76 83.44 ? 9.89 
80:20 2404 ? 150 9.28 ? 2.49 32.10 ? 2.08 46.05 ? 3.09 
100:0 3430 ? 168 31.25 ? 5.74 60.27 ? 10.57 92.06 ? 16.70 
  
56 
 
 Chitosan and alginate films show good breaking strength, but alginate film show 
lower deformation at break as shown in Figure 2.14. The tendency with the increase of 
chitosan content between Young?s modulus and deformation at break is slightly 
contrary. 60:40 CH:AL film has low deformation at break along with 20:80 CH:AL and 
0:100 CH:AL films. There is also the same tendency for all films in Young?s modulus, 
yield strength and stress at break. 
 Arvanitoyannis et al. (1998) produced films from a binary polysaccharide network 
composed of 2 w/v% chitosan and 5 w/v% gelatin. They cast phase separated and 
08?0.06 mm thick films which are much thicker than our films. In addition, the 50:50 
composite film has 2050?200 MPa modulus and 4.1?0.5 % deformation at break. In 
comparison with the same ratio of our composite film, we obtained higher values in 
terms of modulus and deformation at break. 
2.3.8 Antibacterial testing - Zone of inhibition test 
The antibacterial behaviors of films with composition of 100:0, 60:40, 80:20 and 
0:100 CH:AL were evaluated and the results were reported whether there is inhibition 
zone of bacterial growth at the film/medium interface. Figure 2.15 shows that the 
chitosan (100:0 CH:AL) film itself and the 80% and the higher content of chitosan 
containing composite films could inhibit the bacterial growth at the interface. 
57 
 
 
 
Figure 2. 15 Representative picture of inhibitory effect of the selected films: 100:0, 
60:40, 80:20 and 0:100 CH:AL films 
58 
 
 Alginate film is taken as a reference sample on account of its non-bioactivity 
against bacteria. Based on the charge potential of the films, the composite films have a 
decreased number of charges as a result of the possible ionic interaction of polycationic 
chitosan and polyanionic alginate. Consequently, the film with a lower content of 
chitosan such as 60% does not exhibit any biocidal activity. Tsai et al. (2002) 
investigated that the antibacterial action of chitosan increases with the increase of 
deacetylation degree (DD) of chitosan. So 80% DD of chitosan is the most apparent 
reason that the expected antibacterial activity vanishes in composite films with higher 
than 50% of chitosan. This effect of DD in chitosan-alginate composition is overcome 
through a greater content of chitosan such as 80% and 90%. 
 The antibacterial mechanism of chitosan is still unknown. However, different 
approaches have been suggested. Rabea et al. (2003) reported that interaction 
between ?NH3+ of chitosan and negatively charged bacterial cell membranes causes 
the leakage of proteinaceous and other intracellular constituents. Chitosan was mostly 
effective on the outermost surface of the bacteria (Rabea et al., 2003). At a low 
concentration (<0.2 mg/mL), the positively charged chitosan likely binds to the anionic 
bacterial surface to induce agglutination, whereas at high concentrations, more positive 
charges might give a final positive charge to the bacterial surfaces to maintain them in 
suspension (Rabea et al., 2003). 
 
 
 
 
59 
 
2.4 CONCLUSION 
Our NMR results confirmed that the deacetylation degree of chitosan used is 
~80%. Through zeta potential measurement, we observed that in a mixture solution of 
chitosan and alginate, the net charge of the system decreased due to polyelectrolyte 
complexation. After preparing the composite films, their transparency did not show 
significant change due to the complexation compared to either chitosan or alginate film. 
FTIR spectroscopy of the composite films showed that characteristic absorption bands 
of each one of the polymers shifted which indicated an ionic interaction between 
chitosan and alginate.  
Thermogravimetric analyses of the selected films revealed degradation and 
decomposition of the biopolymers following the evaporation of water. Calorimetric 
analyses were in agreement with TGA analyses but showed no indication of glass 
transition and melting temperature of any of the biopolymers used. On DMA results, a 
transition peak at 155?C of chitosan film was also observed and intensified with 
increase of chitosan content in the composite films. However, none of the results was a 
proof of glass transition and melting temperature. Additionally, the storage moduli of 
composite films increased due to strong polyelectrolyte complexation between the two 
polysaccharides without 10% water content. 
Even though the composite films have a miscible appearance, SEM pictures 
displayed that they are phase separated at a microscopic level. Mechanical behaviors of 
the films revealed that 60:40 CH:AL film was the ideal composition for chitosan and 
alginate components. Finally, antibacterial activity of the composite films was found in 
60 
 
80:20 CH:AL film in which positive charged amine groups are more available to kill the 
bacteria.  
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
61 
 
 
 
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64 
 
 
 
CHAPTER THREE 
CHITOSAN AND ALGINATE WET SPUN FIBERS 
3.1 INRODUCTION 
Fibers produced from renewable resources, in particular polysaccharides, have 
drawn considerable attention for their outstanding biocompatibility, non-toxicity, 
biodegradability and intrinsic bioactivity in applications, such as wound management, 
tissue engineering, and drug delivery (Knill et al., 2004, Tuzlakoglu and Reis, 2008, Liao 
et al., 2005). The number of studies on alginate and chitosan as polysaccharide 
biopolymers has been expanding due to their potential use in biomedical applications. 
Fibers have been widely utilized in biomedical applications on account of their 
typical favorable properties, for instance, large surface area, softness, high porosity, 
and ease of production into many forms (Knill et al., 2004). Several commercial wound 
dressing materials (woven and non-woven dressings and hydrogels) have been 
manufactured from polysaccharides and their derivatives. For example, support and 
compression bandages, absorbents, gauzes, tulle dressings, and wound dressing pads 
are made from woven cellulose fibers (cotton and viscose) (Kennedy et al., 1996, Lloyd 
et al., 1998, Kennedy and Thoriey, 2001, Knill et al., 2004). 
The favorable properties (mentioned earlier) of biodegradable fiber structures 
can be modified for specific applications (Tuzlakoglu and Reis, 2008). For instance, they 
can replace many tissues, such as nerve, muscle, tendon, ligament, blood vessel, bone, 
65 
 
and teeth which already have cannular or fiber-shaped architectures and anti-symmetric 
properties. Therefore, fiber-based structures can generate a myriad of applications in 
tissue engineering. On the other hand, numerous factors must be taken into account in 
the fiber design for such applications, such as ideal fiber diameter and linear mass, 
overall absorbency and the distribution of pore size, effect of fiber orientation on 
response of cells, and effect of degradation on the properties of the structures 
(Tuzlakoglu and Reis, 2008). 
Liao et al. (2005) suggested that polyelectrolyte complex fibers comprised of 
chitosan and alginate can be used to control drug or and protein release. Compared to 
microparticule and bead systems, there are less parameters, such as complexation and 
the drawing ratio of fibers affecting protein entrapment efficacy and release behaviors. 
Besides that, drug loading process does not require complicated process stages. 
However, the molecule of target must possess ionic charge groups in order to obtain 
continuous drug delivery. Overall, the polyelectrolyte complex fibers could be a 
promising drug deliverer with the benefits of high effective encapsulation, high loading 
level, sustained release profile, and continuum of the activity of biomolecules (Liao et 
al., 2005). 
Chitosan based materials (details given in Chapter 1) can be generated as 
powder, films, beads, fibers, and fabrics (Qin and Agboh, 1998). However, materials 
produced from pure chitosan fibers have not been practically applicable as a result of 
their insufficient tensile properties and the relatively high production costs 
(Watthanaphanit et al., 2009). Furthermore, pure chitosan is currently not accessible for 
a mass production of those fibers (Watthanaphanit et al., 2009). Another disadvantage 
66 
 
of chitosan is its poor solubility in water which limits its applicability as a polysaccharide 
drug. Although this drawback can be overcome through chemical modifications (Kurita 
et al., 1998), the complexity of reaction process and the relatively high cost of 
manufacturing final product are usually the main disadvantages (Watthanaphanit et al., 
2009). 
Alginate fibers have also been produced and specifically used in wound 
management (Qin, 2008). Because of alginate?s high water absorbency and insolubility 
generated by the present of divalent cations, such as Ca2+, alginate-based fiber mats 
was produced as a wound dressing application (Aogren, 1996). However, as previously 
stated, alginate does not possess an intrinsic antibacterial property thereby wounds 
may not be sterilized form colonization of microorganisms, which may cause infection 
and hinder healing, through that of fiber mat. Nevertheless, integration of chitosan into 
alginate or vice versa could improve the properties of the fibers with the either one of 
the component. The interaction between these two biopolymers enables the 
development of tailored bio-fibers through their potential to form a polyelectrolyte 
complex via ionic interactions. 
There are some examples of mixture of individually assembled alginate and 
chitosan fibers. Pandit (1998) produced alginate and chitosan staple fibers separately 
and then blended in order to achieve a nonwoven wound dressing. Cole and Nelson 
(1993) employed chitosan as the cationic core for the production of an alginate fiber. 
Tamura et al. (2002) coated of chitosan on alginate filament and Knill et al. (2004) and 
Niekraszewicz et al. (2006) independently obtained a fiber of which alginate is the core 
and chitosan is the skin. Knill et al. (2004) proved that adsorption of chitosan into 
67 
 
alginate fiber or coating onto an alginate filament core could be elevated via 
hydrolization. Due to the better mobility of the hydrolyzed chitosan chains, the amount 
of chitosan incorporation increases (Knill et al., 2004). Nevertheless, coating though 
ionic interactions has been the basis of most of these methods.  
In this part of the project, we developed a novel method of forming wet-spun 
chitosan-alginate hybridized fibers and analyzed their mechanical and morphological 
properties through dynamic mechanical analyzer (DMA), scanning electron microscopy 
(SEM) and optical microscope. Total nitrogen analysis was also included in order to 
calculate the chitosan content of the hybrid fibers. 
 
3.2 EXPERIMENTAL 
3.2.1 Materials 
Commercial low molecular weight chitosan (average Mw=150,000 Da and 
deacetylation degree (DD) 80%) was supplied by Fluka. Commercial sodium alginate 
was purchased from Fisher Scientific. Sodium chloride (NaCl) and glacial acetic acid 
(CH3COOH) was purchased from Fisher Scientific.  
Syringe pump was supplied by Fisher Scientific. Syringes were purchased from 
HSW (Henke Sass Wolf), and needles were supplied by Becton Dickinson. All the 
materials were used as received.  
 
 
68 
 
3.2.2 Methods and Techniques 
3.2.2.1 Chitosan solution preparation 
Low molecular weight (LMW) chitosan powder was introduced into distilled water 
and next acetic acid was then added during the mixing process in order to reach 0.5 
w/v% concentration. The chitosan dissolution process was carried out at a temperature 
of 20?C. The solution was also filtered through a 325 mesh stainless steel woven disk 
filter to remove the undissolved matter. After filtration, the pH of the chitosan solution 
was measured as 4.2 ? 0.1. The chitosan solutions prepared were used after suitable 
fragmentation via sonication for the process of forming the alginate-chitosan fibers.  
3.2.2.2 Ultra-Sonication 
Sonication is a simple technique that uses high-intensity acoustic energy to 
agitate particles in a sample for various purposes. In this project, sonication was used in 
order to reduce the molecular weight of low molecular weight chitosan (?150 kDa). The 
probe of sonicator was dipped into chitosan solution and sonication was applied at high 
amplitude (40%) for a short period of time (20 min).  
3.2.2.3 Determination of the average molecular weight of chitosan 
The average molecular weights of non-treated and sonicated chitosan were 
determined by the viscometric method on the basis of intrinsic viscosity [?]. The 
determination of the intrinsic viscosity as a function of the concentration was performed 
with an Ubbelohde capillary viscometer (? = 0.58 mm) having a solvent flow time of at 
least 100 s. Solutions for viscometric measurements of the molecular weight of chitosan 
were prepared by a standard procedure (Pogodina et al., 1986). First, weighed portions 
of chitosan were dissolved in 0.33 M CH3COOH. Then, NaCl was added up to a 
69 
 
concentration of 0.3 mol/l to suppress the polyelectrolyte effect. The solutions were 
passed through a filter of 325 mesh stainless steel woven disk to remove insoluble 
materials. 
The capillary viscometer was filled with 10 ml of the sample and equilibrated in a 
water bath which was controlled with a thermostat to maintain the temperature at 21?C. 
Each sample was run 5 times. The running times of the solution and solvent were taken 
to calculate the kinematic viscosity. Kinematic viscosity can be converted to dynamic 
viscosity through: 
                     (Equation 3.1) 
In this equation   is the kinematic viscosity in stokes;   is the dynamic viscosity in 
poise; and   is the density in g/cm3. Because the concentrations are low and the density 
of the solution and solvent are adjacent to 1 g/cm3, the stoke (centistoke) is equal to 
poise (centipoise). The solution and solvent viscosities are used to calculate the relative 
viscosity, specific viscosity, reduced viscosity, and inherent viscosity on Equation 3.2, 
3.3, 3.4 and 3.5 (Launay et al., 1986, Chen and Tsaih, 1998). The value of the intrinsic 
viscosity [ ] was determined by the mean intercept of Huggins and Kraemer plots, using 
the Huggins and the Kraemer equation (Equation 3.6 and 3.7). 
Relative viscosity (    )          (Equation 3.2) 
Specific viscosity (   )  (  
 
)      (Equation 3.3) 
Reduced viscosity (    )            (Equation 3.4) 
Inherent viscosity  (    )  (      )      (Equation 3.5) 
70 
 
Huggins equation       [ ]   [ ]      (Equation 3.6) 
Kraemer equation          [ ]    [ ]      (Equation 3.7) 
where t is the running time of the chitosan solution, ts is the running time of the 
solvent, c is the chitosan concentration in [g/dl] and kH and kK are the Huggins and 
Kraemer constants, respectively. 
The values of viscosity average molecular weight (Mv) were calculated according 
to the Mark-Houwink equation: 
[ ]                (Equation 3.8) 
where K and a are constants for measurements, K = 3.41?10-5, a = 1.02 
(Pogodina et al., 1986). These values of the constants were chosen due mainly to 
appropriateness in respect of the deacetylation degree (DD) of chitosan Pogodina et al. 
(1986) used and molecular weight range, 13-193 kDa. 
3.2.2.4 Preparation of the sodium-alginate spinning solution 
The sodium-alginate spinning solution (1% w/v) was prepared by dissolving 
sodium alginate powder in distilled water. The solution then was filtered through a 325 
mesh stainless steel woven disk filter to remove the undissolved matter. After filtration, 
pH of the solution was measured as 7.2 ? 0.1. 
3.2.2.5 Formation of the alginate-chitosan fibers 
The sodium-alginate spinning solution, after filtration, was sucked into a plastic 
syringe with a 10 ml volume. Then, the determined amount of the solution was fed by a 
syringe pump through the selected needle into chitosan solution at a flow rate of 3.0 
71 
 
ml/min. (Figure 3.1). 30 G needle which has 0.16 mm inner diameter was used for the 
process. 
The spinning solution was pressurized into chitosan solution to generate 
chitosan-alginate fiber. The solution gelled after mixing with chitosan solution at the 
needle outlet. After the gels grew into a round piece with ?1 mm3 size, the gelled piece 
was pulled up with a pair of fine tweezers to let the stream of the spinning solution flow 
in a fiber shape.  
 
 
Figure 3. 1 An illustration of the fiber formation 
 
Fibers in gel state were retained in chitosan solution for different periods of time 
(1, 3, 6, 12, 24 hours). After collected from coagulation bath, each fiber was soaked in 
distilled water in order to remove the excess chitosan. They were finally left to dry at 
30oC in a convection oven. 
72 
 
If one inquiries the necessity of reducing the molecular weight of chitosan, our 
motive was to incorporate more chitosan into fibers. Initially, the fibers composed of 
non-treated chitosan (Mw = 150 kDa) and sodium alginate were also produced by 
drawing the gelled piece at the tip of the needle unlike the fibers which are formed 
simultaneously in the chitosan solution of reduced molecular weight. The former type of 
fibers was not uniform and had weak points which negatively affected the stability of the 
process of the formation of that fiber (Figure 3.2). This effect occurred most likely due to 
the misalignment of two polymers during micro-fluidic formation. 
 
 
Figure 3. 2 Fiber produced with non-treated chitosan solution 
 
The formation of the latter type of fiber is very similar to that of cross-linked either 
chitosan or alginate fiber. During crosslinking process, for example, sodium alginate 
solution is fed into calcium chloride solution bath whereby sodium alginate is 
precipitated out in filament form as a calcium alginate fiber. The Ca+2 ions (in our case 
73 
 
chitosan polymers) quickly diffuse into the gel solution and crosslink the polymer chains 
of alginate into a gel state through electrostatic force. 
3.2.2.6 Diameter of fibers and the linear density 
 The diameters were measured on Spencer optical microscope made by 
American Optical Company [US]. 10 readings were taken randomly from 10 cm of each 
fiber and the average value of each fiber?s diameter ( ) was calculated and plotted 
against the retention time (Figure 3.3). For the calculation of the linear mass density of 
fibers, denier (Equation 3.9) was chosen as the unit. The density ( ) of fibers is about 
1.6 g/cm3 since their main component is alginate. 
          (     )    (     )         (Equation 3.9) 
 
 
Figure 3. 3 The increase of the diameters of fibers with the retention time 
 
74 
 
3.2.2.7 Total nitrogen analysis 
5 mg of 20 minutes sonicated chitosan powder which was collected by freeze 
drying method and the two fibers with 1h and 24 h retention time were sent to Atlantic 
Microlab, Inc. They performed the analyses by combustion using automatic analyzers. 
The test was aimed to determine the chitosan content of fibers from their nitrogen (N) 
contents. 
3.2.2.8 Morphology analyses 
The fractured cross section of the fibers pre-chilled in liquid nitrogen was studied 
by scanning electron microscope (SEM, Cambridge 360). Samples were sputtered with 
gold prior to SEM observation. Pictures of each fiber were taken under optical 
microscope that is also used for the measurement of diameters of fibers. Longitudinal 
roughness (Ra) and circumferential roughness (Ra) were measured on those pictures by 
an SPM data visualization and analysis computer program [Gwyddion].   
3.2.2.9 Mechanical properties 
Tensile tests were performed using a Dynamic Mechanical Analyzer (DMA RSA3 
TA) Instrument on multiple extension mode, at room temperature (23?C ? 2?C). The 
initial length of fiber sample was 20 mm. It was clamped between grips and stretched by 
top grip at a rate of 0.05 mm/sec. The yield strength (?y), elongation at break (?b) and 
elastic modulus (E) and tenacity were calculated according to the ASTM D-3822-07 
Standard Test Method for Tensile Properties of Single Textile Fibers (2007). 
 
75 
 
3.3 RESULT AND DISCUSSION 
3.3.1 Average molecular weight of chitosan 
Figure 3.4 and Figure 3.5 show Huggins/Kraemer plots of chitosan solutions 
which were used to calculate the [ ] values. Using Mark-Houwink equation (Equation 
3.8), viscosity average molecular weights were obtained (shown in Table 3.1). 
 
 
Figure 3. 4 Huggins and Kraemer plot of non-treated chitosan solution 
 
Figure 3. 5 Huggins and Kraemer plot of 20 min. sonicated chitosan solution 
y = 36.622x + 6.141 
y = 8.5676x + 6.2889 
3
4
5
6
7
8
9
0 0.02 0.04 0.06
Visc
osity
 (d
l/g)
 
Concentration (g/dl) 
Non-treated Chitosan Solution 
Reduced
viscosity
y = 2.7714x + 1.5249 
y = 1.008x + 1.5526 
0
0.5
1
1.5
2
2.5
0 0.05 0.1 0.15 0.2
Visc
osity
 (d
l/g)
 
Concentration (g/dl) 
20' Sonicated Chitosan Solution 
Reduced
viscosity
76 
 
Table 3. 1 Viscosities and molecular weights of chitosan solutions 
 
?int  
intrinsic 
viscosity 
(dl/g) 
Mv  
viscosity average       
(kDa)  
(Experimental 
data) 
Mw  
weight average 
(kDa)  
(provided by Fluka, 
USA) 
Non-treated 
chitosan 
solution 
6.215 143.7 150 
20' sonicated 
chitosan 
solution 
1.54 36.6 - 
 
The viscosity average molecular weight of non-treated chitosan was measured in 
order to confirm the accuracy of the method. The value of Mv of chitosan is very close to 
the value of weight average molecular weight (Mw) of chitosan reported by the selling 
company, Fluka USA (shown in Table 3.1). These experiments confirm the theoretical 
distribution of molecular weights since the various average molecular weights always 
rank in the order: 
  ????    ????    ?????    ????      ???????    ????         (Equation 3.10) 
 
3.3.2 The chitosan content of fibers 
The total nitrogen analysis was performed on chitosan?alginate fibers to 
determine their nitrogen content so as to determine the percentage by weight of 
chitosan in the chitosan-alginate fibers. The results are displayed on Table 3.2.  
 
 
 
77 
 
Table 3. 2 The chitosan content of fibers 
Element 20? sonicated chitosan powder Fiber with 1 hour retention time Fiber with 24 hour retention time 
Nitrogen 
(% w/w) 7.1 0.21 0.33 
Chitosan 
(% w/w) 100 2.96 4.65 
 
With more retention time in chitosan solution, it was already predicted that 
chitosan would penetrate more into the gelled fiber. The result obtained from the total 
nitrogen analyses supports this phenomenon; the fiber with 24 hour retention time 
contains 4.65 w/w% chitosan. 
Tamura et al. (2002) produced a fiber with a highest chitosan content of 0.067 
w/w%. Compared to their study, we were able to incorporate a lowest chitosan content 
of 2.96 w/w% with at least 1 hour retention time in chitosan solution. Even though the 
formation of our fibers was not based on a continuous system, chitosan was not coated 
along the length of the fibers but bonded to alginate polymer.  
3.3.3 Morphological characteristics of fibers 
 It is well studied that the surface properties of scaffolds change the interaction 
between cell and material through modified surface chemistry and roughness (Yilgor et 
al., 2009). As potential fibers for biomedical applications, we measured the roughness 
of fibers on the optical microscope pictures. Table 3.3 shows values of the roughness 
on fibers. These values are quantitative measurement of the roughness which can also 
be observed qualitatively on the optical microscope pictures and the SEM images of 
78 
 
fibers (Figure 3.6 and 3.7). Both longitudinal and circumferential roughness increase 
with gradual retention time in chitosan solution and thus with the growth of the diameter 
of fibers. 
 
Table 3. 3 Changes of roughness of the fibers in longitude and in circumference 
Retention 
Time (h) 
Diameter 
?m 
Ra (nm) 
longitudinal 
Ra (nm) 
circumferential 
1 82.5 29.8 72.4 
3 125 49.1 89.3 
6 130 55.4 93.2 
12 147.5 58.3 107.7 
24 150 69.8 118.5 
 
        
79 
 
        
 
Figure 3. 6 Optical microscope pictures of fibers; a) 1h, b) 3h, c) 6h, d) 12h, e) 24h 
retention time 
 
The diameter and the roughness of fibers (82.5 ? 150 ?m) were dependent upon 
the retention time in chitosan coagulation bath, suggesting more penetration of chitosan 
macromolecules into gel fiber. This was also confirmed with total nitrogen analysis of 
the hybrid fibers. The diffusion of chitosan into gel-state fiber occurred around the fiber 
creating slightly marked curved indentations. Figure 3.7 shows SEM pictures of surface 
80 
 
and cross section of fibers with different retention time. The surface of every fiber was 
characterized by distinct curved indentations, which were the main cause of roughness. 
 
 
 
Figure 3. 7 Representative SEM images of surface of fibers coagulated with increasing 
retention time in chitosan solution; a, b) 1h, c, d) 12h, e, f) 24h 
81 
 
  It is also important to emphasize that there is no coating of one polymer onto 
another. The cross section of fibers (Figure 3.7) shows no sign of layered or two phased 
structure but instead a hybridization of two polymers. It is also safe to assume that 
possible ionic interaction occurred through coupling of chitosan and alginate in the gel 
state of fibers. The coupled network of chitosan and alginate can be formed through 
heterotypic junction zones that bind them together. 
3.3.4 Mechanical properties 
Mechanical properties in terms of tenacity, elongation at break, and Young?s 
modulus of all fibers were investigated and results are shown in Figure 3.8. All plots 
were drawn against the retention time (h) of the prepared samples in chitosan solution. 
 
             
 
82 
 
 
Figure 3. 8 a) Tenacity, b) Elongation at break and c) Young?s modulus of fibers with 
increasing retention time 
 
Clearly, higher retention times, in other words deeper penetration of chitosan into 
fibers, reduced modulus and tenacity in tensile tests, while it largely improved the 
ultimate deformation of fibers. Fibers became less rigid and more flexible and 
stretchable. Alternatively, lower modulus can be explained through the fracture behavior 
of the fiber with 24 hour retention time. Its SEM picture (Figure 3.7(e, f)) shows that its 
cross section is different from other fibers. So it can be concluded that additional 
incorporation of chitosan significantly changed the fracture behavior as well as the 
rigidity of fibers. 
 
 
83 
 
3.4 CONCLUSION  
Fibers containing at least 2.96% of chitosan were obtained by a method for 
forming chitosan-alginate fibers which consisted in introducing alginate spinning solution 
into the chitosan coagulation bath. The formation of chitosan-alginate fibers consists of 
using chitosan solution with a reduced molecular weight, 36.6 kDa which was calculated 
through dilute solution viscometry. 
Chitosan content of fibers completely depends on the retention time in chitosan 
solution. As the time increased so did the chitosan content. This also changed majorly 
morphology and mechanical properties of fibers. 
The diameters of fibers enlarged with more inclusion of chitosan, also affecting 
surface morphology throughout the fiber. Roughness changed visibly and quantitatively 
in a gradual manner towards a higher degree. This can be observed in both optical 
microscopy and SEM pictures and confirmed with roughness measurements.  
Mechanical properties in terms of modulus, tenacity and elongation at break were 
determined. The tensile results showed that the modulus and tenacity values were 
greatest at the lowest content of incorporated chitosan (i.e., 2.96 w/w% chitosan), while 
they decreased with a further increase in the chitosan content. But elongation at break 
values reversibly increased in the same manner of chitosan content. 
 
 
 
 
 
84 
 
 
 
 
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