Andreia Leal Pereira
Estruturas
fibrosas
de
PCL/Quitosano
para
engenharia de tecidos
Fibrous scaffolds from PCL/Chitosan blends for
tissue engineering
Andreia Leal Pereira
Estruturas
fibrosas
de
PCL/Quitosano
para
engenharia de tecidos
Fibrous scaffolds from PCL/Chitosan blends for
tissue engineering
Dissertação apresentada à Universidade de Aveiro para cumprimento dos requisitos necessários à obtenção do grau de Mestre em Materiais e Dispositivos Biomédicos, realizada sob a orientação científica da Doutora Maria Helena Figueira Vaz Fernandes, Professora associada do Departamento de Engenharia de Materiais e Cerâmica da Universidade de Aveiro e do Doutor Samuel Guieu, Investigador do Departamento de Química da Universidade de Aveiro.
“A vida é mesmo assim. Vai pegar em ti e levar-te para onde achar que lhe fazes mais falta, onde sentir que lhe fazes mais diferença. “
o júri
presidente Prof. Doutor José Maria da Fonte Ferreira
professor associado com agregação da Universidade de Aveiro
Prof. Doutora Ana Luísa Daniel da Silva
Investigadora auxiliar da Universidade de Aveiro
Prof. Doutora Maria Helena Figueira Vaz Fernandes
agradecimentos Ao longo deste trabalho tive o apoio e ajuda de várias pessoas a quem devo um sincero agradecimento.
Em primeiro lugar gostaria de agradecer aos meus orientadores Dra. Maria Helena Figueria Vaz Fernandes e Dr. Samuel Guieu, pela paciência, pelos ensinamentos, pelo apoio e por todo a motivação dada ao longo deste projeto. Às técnicas Marta Ferro e Maria Celeste Azevedo e ao Dr. José Carlos Almeida e Dra. Erika Davim pela ajuda e disponibilidade prestadas, principalmente nas alturas de maior urgência.
À Dra. Nathalie Barroca por me ter posto em contacto com a técnica TISA, assim como pela ajuda prestada no início de todo este projecto.
À Dra. Margarida Costa por toda a disponibilidade e ajuda na realização das análises de micro-CT.
Ao André Girão, Diana Teixeira, Filipa Rodrigues, Frederico Ribeiro, Karolina Rolinska, Manuel de Barros e Tânia Carvalho, pelos bons momentos passados no laboratório, pela amizade e partilha de conhecimentos nas mais variadas áreas.
Às minhas colegas de casa Ana Rita Bandarra Nunes, Ana Beatriz Barata Pinto e Cardoso Bandeira, Beatriz Rocha Araújo e Jéssica Alexandra de Sá Antunes por todos os momentos partilhados nesta vida académica, que serão preciosamente conservados na minha memória.
E por último, à grande mulher e minha mãe, por todo o amor e suporte, pelos sacrifícios que fez para que pudesse concluir esta jornada e por me demonstrar que por mais difícil que sejam os obstáculos, com esforço e dedicação, tudo se consegue alcançar. E vá, também ao meu irmão (garoto).
palavras-chave Scaffolds nanofibrosos 3D electrospinados, Policaprolactona, Quitosano, Engenharia de tecidos
resumo O desenvolvimento de estruturas artificiais (scaffolds), que imitem, o mais perfeitamente possível, a matriz extracelular, e que auxiliem na regeneração dos tecidos vivos, tem sido uma das principais áreas de intervenção em engenharia de tecidos. Arquiteturas nanofibrosas bidimensiomais podem ser obtidas por electrofiação (electrospinning), enquanto que estruturas tridimensionais são muito difíceis de obter diretamente pelo mesmo método. Posto isto, um grupo de investigadores, recentemente desenvolveu uma técnica chamada Thermally Induced Self-Agglomeration (TISA) que permite transformar membranas bidimensionais obtidas por electrofiação em estruturas tridimensionais. Este trabalho teve como objetivo, produzir e caracterizar, membranas por electrofiação de uma mistura de PCL/quitosano, para a seguir convertê-las em estruturas 3D por TISA, seguida de liofilização. Os produtos obtidos foram scaffolds 3D nanofibrosos com crescentes quantidades de quitosano (10, 15 e 20%), altamente porosos (>90%) com poros interconectados de variados tamanhos. Módulos de compressão indicaram compatibilidade para engenharia de tecidos da cartilagem. Os resultados mostraram que os scaffolds apresentavam alta similaridade tanto na morfologia como nas suas propriedades com a matriz extracelular natural e que por isso, a sua aplicação em engenharia de tecidos deverá ser bastante promissora.
keywords 3D electrospun nanofibrous scaffolds, Polycaprolactone, Chitosan, Tissue engineering
abstract The development of artificial structures (scaffolds), that mimic the extracellular matrix as closely as possible, and that aid in the regeneration of living tissues, has been one of the main areas of study in tissue engineering. Two-dimensional nanofibrous can be obtained by electrospinning, but three-dimensional structures are very difficult to obtain directly by electrospinning. Because of that, a group of researchers recently developed a technique called Thermally Induced Self-Agglomeration (TISA) that allows transforming two-dimensional electrospun membranes into three-dimensional structures. The objective of this work was to produce and characterize electrospun membranes of PCL/chitosan blends, to then convert them into 3D structures by TISA, followed by freeze drying. The obtained products were nanofibrous 3D scaffolds with increasing amounts of chitosan (10, 15 and 20%), highly porous (>90%) and with interconnected pores of different sizes. Compression modulus indicated compatibility for cartilage tissue engineering. The results demonstrated that the obtained scaffolds presented high similarity both in morphology and properties to the natural extracellular matrix. Therefore, its application in tissue engineering should be very promising.
i
List of figures ... iii
List of Tables ... v
List of Abbreviations ... vii
Chapter I - State of the Art ... 1
1.1 Introduction ... 3
1.2 Scaffolds ... 4
1.2.1 Structure and morphology ... 5
1.2.2 Composition ... 5
1.2.3 Biological parameters ... 6
1.2.4 Manufacturing technologies ... 7
1.3 Electrospinning ... 8
1.3.1 Description ... 8
1.3.2 Electrospinning processing parameters ... 9
1.3.3 Polymeric nanofibers in electrospinning ... 12
1.4 Polycaprolactone ... 13 1.4.1 Properties ... 13 1.4.2 Electrospinning of PCL ... 14 1.5 Chitosan ... 15 1.5.1 Properties ... 15 1.5.2 Electrospinning of chitosan ... 17 1.6 Electrospinning of PCL/Chitosan ... 19 1.7 Three-dimensional architectures ... 20
1.8 Thermally induced self-agglomeration ... 24
1.9 Objectives ... 25
Chapter II - Materials and Methods ... 29
2.1 Materials and Reagents ... 31
2.2 Preparation of solutions ... 31
2.3 Production of PCL/Chitosan composite membrane by electrospinning ... 32
2.4 Manufacture of a 3D nanofibrous scaffold ... 33
2.4.1 Conversion of membranes into small pieces ... 33
2.4.2 Fabrication of 3D PCL/chitosan scaffolds ... 34
ii
2.5.1 Scanning electron microscope (SEM) analysis ... 35
2.5.2 Viscosity measurements of the solution ... 35
2.5.3 Fourier transform infrared (FTIR) spectroscopic measurements ... 35
2.5.4 Wettability Tests ... 35
2.5.5 Mechanical properties ... 36
2.5.6 Differential scanning calorimetry (DSC) measurements ... 37
2.5.7 Porosity... 37
2.5.8 Micro-computed tomography (μCT) analysis ... 37
2.5.9 Swelling tests... 38
Chapter III - Results and discussion ... 41
3.1 Electrospinning ... 43
3.1.1 Optimization of the operating conditions ... 43
3.1.2 Properties of the solution... 44
3.1.3 Physical-chemical characterization of PCL/Chitosan membranes... 50
3.2 TISA ... 57
3.2.1 TISA optimization ... 57
3.2.2 Morphologies and properties of the scaffolds ... 59
Chapter IV – Conclusions and Perspectives ... 67
References ... 71
iii
Figure 1 - A typical tissue engineering cycle. ... 4
Figure 2 – Different forms of polymeric scaffolds for tissue engineering. ... 5
Figure 3 - Different types of polymeric nanofibers with different sizes.[A] Poly(1,4 butylene succinate) (PBSu); [B] PHBV-PBSu; [C] Chitosan-PVA; [D] PLGA; [E] PLA; [F] PHBV. ... 8
Figure 4 - Schematic representation of a typical electrospinning system. ... 9
Figure 5 - Effect of voltage variation on Taylor cone formation. ... 10
Figure 6 - SEM images of electrospun nanofibers from different polymer concentration solutions. ... 11
Figure 7 - Structure of PCL. ... 13
Figure 8 - Ring opening polymerization of e-caprolactone to polycaprolactone. ... 13
Figure 9 - Structure of Chitosan. ... 16
Figure 10 - SEM images of PCL/Chitosan nanofibers with an AA/FA solvent ration of 3/7 (a) and 5/5 (b). ... 20
Figure 11 - SEM images of scaffolds of chitosan with (a) 50% PCL and (b) 75% PCL. ... 22
Figure 12 - SEM micrographs of scaffolds sections (a) PCL/CH25, (b) PCL/CH50, (c) PCL/CH75. ... 22
Figure 13 - SEM micrographs of (a) PCL and (b) PCL/CH50 porous scaffolds. ... 23
Figure 14 - SEM microphotographs of (a) CH, (b) PCL/CH75 and (c) PCL/CH50 fiber-mesh scaffolds. ... 23
Figure 15 – A photo of 3D nanofibrous scaffold of PCL with respective SEM images. .... 25
Figure 16 - Components of FLUIDNATEK LE-10 electrospinning equipment. ... 32
Figure 17 – A scheme of the steps followed in the conversion of membrane into small pieces. ... 34
Figure 18 – A scheme of the steps followed in the fabrication of 3D PCL/chitosan scaffolds. ... 34
Figure 19 - Contact angle measurement diagram. ... 36
Figure 20 – Images of the equipment and the assembly carried out in the tensile tests. ... 36
Figure 21 – SEM images of PCL10/CH10 membranes at 13 cm (A, B) and 15 cm (C, D) distance from needle tip to collector... 44
Figure 22 – SEM micrographs of PCL/CH membranes of (A, B) PCL6/CH10, (C, D) PCL10/CH10 and (E, F) PCL10/CH15 using a solvent ratio of 1/1. ... 45
Figure 23 – SEM micrographs of PCL/CH membranes of (A, B) PCL10/CH10, (C, D) PCL10/CH15 and (E, F) PCL10/CH20 using a solvent ratio of 1/2. ... 46
iv
Figure 24 – Viscosity of solutions as a function of the chitosan concentration for 8 and 10 wt.% PCL. (dashed lines serve to guide the eye) ... 47 Figure 25 – SEM images showing the morphologies of PCL/chitosan blend nanofibers electrospun from the solutions with PCL/chitosan concentrations of respectively (A) 8wt. %/10%, (B) 8wt. %/15%, (C) 8wt. %/20%, (D) 10wt. %/10%, (E) 10wt. %/15% and (F) 10wt. %/20%. ... 48 Figure 26 – Average fiber diameter of PCL/chitosan blend nanofibers. ... 49 Figure 27 – SEM micrographs showing the morphologies of (A) pure PCL (14wt. %) nanofibers and (B) PCL/chitosan (10wt. %/20%) blend nanofibers. ... 49 Figura 28 – SEM images showing the identical morphologies of PCL/chitosan blend nanofibers electrospun from solutions with PCL/chitosan concentrations of (A) 8wt. %/10%, (B) 8wt. %/15% and (C) 10wt. %/20%. ... 50 Figure 29 – (A) FTIR spectra of PCL pellets, PCL (10 wt. %)/chitosan (20%) membrane and chitosan powder, (B) an enlargement of the spectra from 1550 to 2000 cm-1. ... 51 Figure 30 – Contact angle images of (A) pure PCL and (B) PCL (10wt. %)/CH (20%) membranes using DI Water. ... 52 Figure 31 – Stress-Strain curves of PCL, PCL/CH10, PCL/CH15 and PCL/CH20 membranes. ... 53 Figura 32 – DSC (A) first and (B) second heating scans of PCL pellets, chitosan powder and the several membranes of PCL/CH. ... 55 Figure 33 – SEM images showing the nanofibers ground in (A) ethanol, (B) water and (C) ethanol/water (50/50) mixture. ... 57 Figure 34 – SEM micrographs of final scaffolds with (A) 2 minutes and (B) 5 minutes of heat treatment. ... 59 Figure 35 – Final structure obtained after TISA process with the PCL10/CH20 formulation. ... 59 Figure 36 – SEM images showing the morphologies of (A1, A2 and A3) pure PCL, (B1, B2 and B3) PCL/CH10, (C1, C2 and C3) PCL/CH15 and (D1, D2 and D3) PCL/CH20 of 3D electrospun nanofibrous scaffolds obtained by TISA (different magnifications). ... 60 Figure 37 – Representative 3D micro-CT images of PCL/CH10 scaffold of (A) total morphology and of (B-D) selected small areas. ... 62 Figure 38 – Swelling ratio of the different formulations after 1h, 6h, 24h, 30h and 48h. ... 63 Figure 39 – Compressive modulus of the different scaffolds formulations. ... 64
v
Table 1 - Concentrations of polymer solutions. ... 31
Table 2 - Conditions of the electrospinning parameters. ... 33
Table 3 - Water contact angle of electrospun membranes. ... 53
Table 4 - Mechanical properties of electrospun membranes. ... 54
Table 5 - Melting temperature values of PCL and various PCL/CH contents. ... 56
Table 6 – Features of the different electrospun 3D nanofibrous scaffolds. ... 61
vii 2D Bidimensional 3D Tridimensional AA Acetic acid CH Chitosan DI Deionized Water
DSC Differential scanning calorimetry
FA Formic acid
FTIR Fourier transform infrared spectroscopy HA Hydroxyapatite
PBSu Poly (1,4 butylene succinate) PCL Poly (-caprolactone)
PEO Poly (ethylene oxide)
PHBV Poly(3-hydroxybutyrate-co-3-hydroxyvalerate) PLA Poly (lactic acid)
PLGA Poly (lactide-co-glycolic acid) PLLA Poly (L-lactic acid)
PU Polyurethane PVA Poly (vinyl alcohol)
SEM Scanning electron microscope TE Tissue engineering
TFA Trifluoroacetic acid
TIPS Thermally induced phase separation TISA Thermally induced self-agglomeration Tm Melting temperature
1
3
1.1 Introduction
In recent years, medicine paradigm has evolved from the concept of repair to the regeneration approach. In this process, there is the replacement or regeneration of cells, tissues or organs in order to restore their normal functions.1 Regeneration may occur through stimulation of the body's repair mechanisms or by replacing the damaged zone with artificial tissues and organs. This has given rise to the development of tissue engineering (TE), the main area that creates artificial substitutes for restoring damaged tissue. The main commonly used substitutes are allografts, xenografts and autografts. But lack of donors, limited amount of material available, immune rejection and transfer of pathogens, in this type of grafts, led to tissue engineering to introduce a new alternative.2 Within this context one of the most promising strategies in TE is to create structures that can allow cell growth and proliferation, for further implantation and integration in the body. Thus construction of scaffolds or 3D structures with high porosity has shown to be a very convenient option.2
Figure 1 depicts the five major steps by which a typical tissue engineering process occurs:3 a) Obtaining cells from the patient;
b) Multiplication in a culture;
c) Implantation of the cells in the scaffolds; d) Growth of the cells inside the scaffolds; e) Placement of the final structure in the patient.
4
Figure 1 - A typical tissue engineering cycle.
1.2 Scaffolds
In tissue engineering, a scaffold is a structure which exhibits a biomimetic environment capable of promoting or repressing physiological responses, and degrading as a new natural tissue grows.4
In our body tissues are organized in structures composed of protein fibers interlaced with each other, with diameters in the nanometer range, where the cells grow and develop, called the extracellular matrix. Scaffolds arose as an attempt to recreate the extracellular matrix.5 A scaffold has as main objective to provide a structure with surface and volume suitable for the attachment of the cells as well as for their growth. To get a perfect scaffold, this should gather some characteristics such as:
1) High porosity with interconnected pores (for cell migration as well as nutrient transport);
2) Fibers with diameters in the nanometer range, to be suitable for the cells attachment; 3) Mechanical properties and flexibility appropriate to the site of implantation;
5
4) Biocompatibility and biodegradability with a controllable rate to match cell growth in vitro and in vivo;
5) Simple manufacturing allowing reproducibility of the method.4
1.2.1 Structure and morphology
When scaffolds are produced, one of the factors requiring more accurate control is structure and morphology. These should be as close as possible to the tissue to mimic, because aspects such as degradation rate, mechanical properties or even cell adhesion depend on them.6 The architecture of a scaffold should have a network of interconnected pores with at least 90% porosity, for a good vascularization, nutrient transport and waste products out of the scaffold. The surface area should also be large for providing enough space for cell attachment and growth.7,8
Currently, there are scaffolds with several different structures, and so there is a major and better application to all areas of the human body. Figure 2 shows some examples: (A) three dimensional porous scaffolds; (B) fibrous matrix and (C) porous microspheres.9
Figure 2 – Different forms of polymeric scaffolds for tissue engineering. 9
1.2.2 Composition
In tissue engineering materials from different classes may be used, namely polymers (natural or synthetic), ceramics and composites.7
Polymers are the most widely used, due to their enormous diversity, as well as their easy processability. Within the natural ones the most common are polysaccharides (cellulose, chitin, chitosan, dextrose) and proteins (collagen, silk, gelatin, elastin). The natural polymers
6
usually exhibit antibacterial properties and good biocompatibility, being therefore more propitious to a good cell adhesion. Synthetic polymers have high mechanical properties and good flexibility. Some of the most used examples are poly (lactic acid) (PLA), poly (-caprolactone) (PCL), poly (ethylene oxide) (PEO), poly (lactide-co-glycolic acid) (PLGA).10 Regarding ceramics the mostly used are bioactive glass (BG), tricalcium phosphate (TCP), hydroxyapatite (HA) and coralline. These have chemical properties similar to the ones of bone, but mechanical properties do not always match. Bioceramics frequently exhibit high compressive and deformation resistance and low ductility. However the bioactivity, bio-efficiency and osteoconductivity of ceramics are properties of considerable interest for their biomedical applications.7,8
More recently composites have emerged as valuable combinations, of two or more materials, put together to enhance their advantages. They can be combined in different shapes and proportions, as polymer-polymer or even polymer-ceramic. The advantages presented are excellent biocompatibility, acceptable rate of degradation, improved mechanical properties and cell adhesion.7,11
1.2.3 Biological parameters
The most important requirement when thinking about implanting scaffolds in the human body is biocompatibility. If the device developed to implant in the body induces a serious inflammatory response, reaction or even toxicity, all previous work will not be of much use.7 Biodegradable and bioresorbable are also important. A scaffold should degrade at a rate of time that allows the new tissue to form. When it is degraded, the waste products should be nontoxic so that the organism can eliminate them without any problem.7
Bioactivity is also a very important requirement particularly in bone-related situations. Bioactivity is the ability of the material to interact with living tissues. Here the material is well accepted by the cells, promoting vascularization and thus a greater cellular growth, avoiding rejection problems.7
7 1.2.4 Manufacturing technologies
A number of manufacturing technologies have been used to fabricate scaffolds.7
Some of the techniques include gas foaming, thermally induced phase separation, particle leaching, sol-gel, freeze-drying, solvent casting and electrospinning. The main disadvantages associated to these technologies are the difficult structure/shape control (porosity, pore size, etc.). 7
Additive manufacturing techniques emerged in order to improve these drawbacks. This technology works by adding layers of material until the final 3D system is obtained. 3D printing, fused deposition modeling, bioprinting, selective laser sintering and stereolithography are some of the existing technologies.7
One of the most interesting areas is related with the production of scaffolds with nanofibrous structures, as they demonstrate enormous potential for various applications, including filters, biomaterials, biosensors, nanocomposites, drug delivery systems, tissue engineering.12 Some of the most prominent methods to make nanofibers are self-assembly, phase separation and electrospinning.12,13
The self-assembly technique consists of a disorganized system of individual and pre-existing components, which organize to form the desired patterns and functions, by non-covalent interactions. It is a simple technique, performed in solution, which consequently entails some disadvantages to the final product, such as poor mechanical properties, difficulty in controlling pore shape and size inside the scaffold and time consuming manufacturing.13,14 Thermally induced phase separation (TIPS) is another technique for forming nanofibers. In this method the polymer solution separates in two phases: a polymer-poor phase and a polymer-rich phase. When the solvent is extracted, only the polymer rich phase remains which is then frozen and later lyophilized. The result is an open nanometric pore foam. It has as disadvantages, a nanofibers processing that is slow and has little control on the diameter and fiber orientation.5,13,14
Electrospinning was the method of choice in this work, and for this reason it will be described with more detail in the next section.
8
1.3 Electrospinning
1.3.1 Description
At the beginning of the 21st century the capacity of electrospinning to manufacture nanofibers was rediscovered, as well as its application in tissue engineering. But this technique already existed. It was first observed in 1897 by Rayleigh, later studied by Zeleny in 1914, and patented by Formhals in 1934.15
With this technique it is possible to produce nanofibers with speed, simplicity, at low cost, using a wide variety of polymers and with diameters in the order of the micrometers down to the nanometers (Fig. 3). In the context of tissue engineering a major advantage of the electrospinning technique is related to the fact that the obtained fibers are very similar in terms of arrangement and average diameter to the extracellular matrix, in particular to those of collagen.15,16
Figure 3 - Different types of polymeric nanofibers with different sizes.[A] Poly(1,4 butylene succinate) (PBSu); [B] PHBV-PBSu; [C] Chitosan-PVA; [D] PLGA; [E] PLA; [F] PHBV. 17
Electrospinning is based on the application of high electrostatic forces to produce fibers. It consists of three components: a syringe pump, a high voltage source and a collector (Fig. 4).16 First a polymer solution is placed in the syringe of the pump, which will drain to the tip
9
of the needle. The solution remains on the tip of the needle, in a drop shape, by surface tension. Then a voltage between 1 and 30 kV is applied to the solution, which causes a repulsion of charges inside the solution. When the charge repulsion force overlaps the surface tension, one or more jets of the drop are fired, depending on the intensity of the electric field. There is the formation of a Taylor cone when a jet leaving the drop becomes convex and uniform. As the jet travels to the normally rotatable collector, the solvent evaporates and the polymer arranges into fibers which are collected on the collector in the form of a membrane.10,14,18
Figure 4 - Schematic representation of a typical electrospinning system.16
1.3.2 Electrospinning processing parameters
There are several parameters that can be changed to facilitate the processing by electrospinning, as well as to improve the characteristics of the obtained nanofibers.19 Parameters such as flow rate, distance between the needle tip and the collector and the voltage are related to the diameter and morphology of the fibers. In addition, the environmental conditions (temperature and humidity) and the properties of the solution (viscosity, conductivity, concentration, surface tension and homogeneity) mainly determine how electrospinning will run.10 The various parameters to control in electrospinning are detailed below.
10 Voltage
The applied voltage influences the formation of the Taylor cone, which in turn influences the diameter of the nanofibers. At a low voltage, a large droplet is formed at the tip of the needle from which a jet free of beads is expelled. As the voltage increases, the drop size decreases and when the voltage is too high there is no Taylor cone formation and the presence of many beads can be observed (Fig. 5). This means that the higher the voltage, the more beads will be found. The voltage is also reflected in the diameter of the fibers. The higher the applied voltage, the less will be the diameters of the fibers. Therefore it is strictly necessary to find a suitable voltage for a stable electrospinning process.18
Figure 5 - Effect of voltage variation on Taylor cone formation.16
Flow Rate
The flow rate is regulated by the syringe pump. This parameter influences the rate of material transferred and the velocity of the jet. In general, the lower the flow rate, the smaller the fiber diameters. Typically, a low flow rate is desired, so that the solvent has time to evaporate. When the rate is high, the diameters are larger, but the formation of droplets is also observed. The flow rate should follow the formation of the fibers, in order to obtain a stable process.15,18
11 Distance between the needle and the collector
The variation of the distance between the needle and the collector is related to the electroelastic force, the evaporation of the solvent and the formation of beads. At a greater distance, the fibers are thinner, because the solvent is better evaporated. At a shorter distance, the solvent has less time to evaporate, and the fibers become thicker. Also, the smaller the distance the greater the probability of observing beads.16,18
Polymer concentration
The concentration of the polymer influences very important parameters in the electrospinning process, such as the viscosity and the surface tension. The surface tension is important in the concentration, because when the concentration is low, the polymer fibers break into droplets before reaching the collector. If the concentration increases, the viscosity increases, and the number of beads decreases (Fig. 6). Therefore, a suitable polymer concentration is required to performed the electrospinning.14,16
12 Solvent
When choosing the solvent, care must be taken to ensure that it is highly volatile, because when the fibers travel from the needle to the collector, it is crucial to have time to evaporate the solvent. Another important parameter is the electrical conductivity. If the conductivity of the solvent is high, it forms fibers of small diameter. On the contrary, if the solvent has low conductivity, the polymer solution tends to form droplets during the process.16
Molecular weight
Molecular weight influences rheological and electrical properties such as viscosity, surface tension, conductivity and dielectric strength. The higher the molecular weight of the polymer, the larger the diameter of the fibers. When the molecular weight is too low, the polymer forms tangles in the solution, and it becomes difficult to transform the polymer chains into fibers.15
Ambient conditions
Environmental conditions such as temperature and humidity are very important parameters for electrospinning. They will influence the rate of the solvent evaporation during the travel of the fibers from the syringe tip to the collector. If the ambient conditions are not relatively identical to those of the experimental achievements, even if the processing parameters are the same, success of electrospinning will be difficult either because it will not be performed or because similar results will not be obtained. Therefore, both temperature and humidity are important parameters to check.18
1.3.3 Polymeric nanofibers in electrospinning
There is a large number of polymers that can be used to produce nanofibers, but for tissue engineering the number is restricted due to the biological requirements they must have. However, the most commonly used are chitosan, gelatin and collagen in the class of natural
13
polymers and PCL, poly (L-lactic acid) (PLLA), poly (lactic-co-glycolic acid) (PLGA), polyurethane (PU) and poly (vinyl alcohol) (PVA) as synthetic polymers.13
In this work, PCL and chitosan were chosen to prepare electrospun blend membranes, being therefore more detailed below.
1.4 Polycaprolactone
1.4.1 Properties
Poly (-caprolactone) (PCL) is a linear aliphatic polyester (Fig. 7), widely used in tissue engineering. It has a melting point of approximately 60 °C, and a glass transition temperature of -60ºC, depending on the degree of crystallinity and molecular weight.21
Figure 7 - Structure of PCL.
PCL can be produced industrially by 2 methods: 6-hydroxycaproic (6-hydroxyhexanoic) acid condensation, or ring opening polymerization of the -caprolactone with a catalyst (Fig. 8).22,23
14
PCL is biocompatible, bioinert, of controllable biodegradation, stable under environmental conditions, available in large quantities and a low cost polymer. Due to its hydrophobic and semi-crystalline nature, and also to its mechanical properties, it has a low degradation rate (2-4 years depending on its molecular weight).21,24,25 Its degradation occurs by the hydrolysis of the ester linkage in physiological media, such as the human body, hence the great interest in using it as a biomaterial.25
In addition to being miscible with other polymers such as poly (vinyl chloride) or poly (acrylonitrile butadiene styrene), it is also mechanically compatible with polyethylene, poly (vinyl acetate) and others polymers.22
PCL has been approved by the Food and Drug Administration (FDA) as a polymer that can be used in the human body, and since then it has been present in numerous applications such as devices for delivering drugs, sutures and scaffolds.21
Comparing PCL to other aliphatic polymers, their physical properties have attracted great interest in researchers. For example, PCL does not exhibit isomers such as PLA, making it easier to work with. The melting point, the biological degradation, and rheological properties are also parameters that allow the scaffold to be manufactured by diverse methods, making them even more attractive. Nevertheless, its hydrophobic nature results in low wettability and poor biological interactions, not allowing the surface to adsorb the amount of protein required for good cell attachment. To overcome this disadvantage, one of the solutions has been changing the surface either chemically or physically, or by coating their surface.21
1.4.2 Electrospinning of PCL
Electrospun PCL has been used in tissue engineering for various areas such as: bone regeneration, cardiovascular tissue, skin tissue and neural regeneration.26 In the production of PCL fibers, it is important to choose a determined molecular weight range and specific solvents, in order to easily reproduce the electrospinning. Preferably 80 kDa has been the PCL molecular weight most used, but as it recently stopped being produced, an alternative may be 70-90 kDa. As for the solvent, there are many options for dissolving the PCL. The most frequently used is dimethylformamide (DMF), but chloroform, methanol,
15
hexafluoroisopropanol (HFIP), tetrahydrofuran (THF) and dichloromethane (DCM) are also very common.21
Most PCL electrospun membranes are obtained through a combination of polymers, reagents or materials, which improve the properties of the final product. In bone tissue engineering several types of PCL composites have been reported.
Patlolla et al. (2010)27, for example, created a polymer-ceramic composite, using hydroxyapatite, β-tricalcium phosphate and PCL. In the electrospinning process, the solvents used were methylene chloride (MC) and MC + DMF, to manufacture the MC-composite and the MC-DMF composite, respectively. The MC-composite showed a bimodal fiber diameter distribution, with pore sizes 79.6 ± 67 μm, whereas in the MC + DMF-composite the fiber diameters were uniform and with pores around 7 μm. In the distribution of hydroxyapatite and tricalcium phosphate only the MC + DMF-composite showed a good dispersion in the composite. In the end, only the MC-composite supported cell proliferation.
Another widely used combination is PCL/gelatin. In the study by Gautam et al. (2013)28 using chloroform/methanol to dissolve PCL and acetic acid for gelatin, uniform fibers were obtained, where the morphology of the fibers depended on the percentage of PCL in the gelatin. Cell adhesion tests indicated high viability to apply in tissue engineering.
The PCL/chitosan combination in the formation of fibers by electrospinning is also referred in the literature. It was first investigated with the aim of restoring neuronal tissues, and was later used for bone tissue engineering.29 A few examples of electrospun PCL nanofibers from this work will be presented later.
1.5 Chitosan
1.5.1 Properties
Chitosan is a semicrystalline polymer formed from 2-acetamido-2-deoxy-β-D-glucopyranose (chitin) that turns into 2-amino-2-deoxy-β-D-2-acetamido-2-deoxy-β-D-glucopyranose (chitosan) (Fig. 9).30 It is mainly obtained from shells of shrimps and crabs.31 Due to the difficulty of
16
dissolving chitin in common solvents, chitosan appeared as an alternative to chitin. Chitosan is soluble in acidic solutions and has a larger application area.32
Figure 9 - Structure of Chitosan.
The process to obtain the chitosan begins by treating the remains of the crustaceans to obtain the chitin. Then, the acetamido groups at the C-2 position of chitin are converted to amino groups (NH2). This process is called deacetylation.33
Chitosan is economically cheap because it is a natural polymer abundant in nature and requires only a very simple processing.30 The characteristics that define the quality and the area of application of chitosan are mainly the molecular weight, degree of deacetylation, surface area and particle size.
Molecular weight
The molecular weight varies according to the number of monomers in the polymer, and it is typically 20 to 1200 kDa. Depending on the molecular weight, chitosan can be classified as low (LMWC), medium (MMWC) and high (HMWC). 34,35 It is one of the most important characteristics, as it affects properties such as viscosity and solubility.
Degree of deacetylation (DD)
The degree of deacetylation is the percentage of free amino groups, reported to the total number of nitrogen atoms (which means: n(amino)/[n(amino) + n(acetamido)]). The greater the number of amine groups, the greater the degree of deacetylation, and consequently the more cationic and soluble the polymer will be.36
17 Surface area
The surface area is related to porosity, pore volume and pore size distribution of the chitosan particles. It is known that chitosan powder or flakes have low surface area. Because chitosan is a non-porous material, some modifications are performed to increase its surface area. These properties are not relevant for the project, because the morphology of the chitosan will be transformed into nanofibers.
Chitosan is biodegradable, biocompatible, bio-renewable, has high activity against bacteria and fungi and still binds to toxic metal ions.37,38 Due to these properties, chitosan has gained interest in biomedical applications. A variety of products is fabricated such as films, gels, fibers or nanofibers, microparticles, scaffolds.32 Nanofibers are among the most studied. The most commonly used techniques for producing chitosan nanofibers are electrospinning, phase separation, sol-gel and self-assembly.38
1.5.2 Electrospinning of chitosan
Chitosan raised a great interest in electrospinning, but because of its polycationic character in acid medium, due to the presence of many amino groups in its structure, the formation of nanofibers is very difficult when this technique is used.39 However, some nanofibers of pure chitosan have already been generated by electrospinning using solvents such as acetic acid (90 wt.%), trifluoroacetic acid (TFA)/dichloromethane (DCM) or formic acid (FA).32
When the solvent used is acetic acid there is a great contradiction in the information reported. According to Geng et al. (2005)40 and Vrieze et al. (2007)41, nanofibers were obtained using 90 wt% acetic acid. But in another article42, using the same protocol, no results were obtained. Examining further the foregoing articles, it was found that the fibers obtained were only for the first half hour of electrospinning, then the jet became so unstable that it did not allow the nanofibers to be obtained. The explanation is that due to the high volatility of the acetic acid, after some time the chitosan fibers begin to deposit at the tip of the needle, causing the jet to become increasingly distorted and the chitosan fibers do not reach the collector. Another disadvantage in using acetic acid is that only nanofibers are obtained, not a membrane, because the process can only work for a short period of time.40–42
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In the literature, the nanofibers produced using TFA43 were achieved due to two reasons: a) the TFA forms salts with the amino groups, leading to the breaking of the rigid interactions of the chitosan molecule, allowing the electrospinning and b) during electrospinning, the high volatility of the TFA (boiling point 71,8 ºC), allows the solidification of the polymer to be fast, forming better nanofibers.43
With TFA, the best results were obtained using a solution containing 8 wt% chitosan. But even so, the nanofibers presented small beads on the fibers. The alternative found was to mix TFA with DCM as organic solvent. The best ratio was TFA/DCM (70/30)44. From the point of view of biomedical applications, TFA is a toxic solvent and dangerous for the environment, thus its use should be limited.39
Although it is possible to produce nanofibers of pure chitosan, the difficulty involved in its manufacture, and its weak mechanical properties, led to a new type of approach. The alternative was to combine chitosan with synthetic polymers such as polyvinyl alcohol (PVA), polyethylene oxide (PEO), polylactic acid (PLA), polycaprolactone (PCL), etc. 37,39 PVA has been widely mixed with chitosan, due to its good fiber forming characteristics. The addition of PVA allows the formation of fibers with diameters between 20-100 nm, with a uniform distribution of both polymers throughout the membrane.37,45
PEO is a biocompatible polymer extensively used in cartilage repair. It is a polymer widely used to aid in electrospinning as it increases electrical conductivity. For example, in 2011, Pakravan et al.39 were able to electrospun highly deacetylated chitosan (97.5%) in 50% aqueous acetic acid with 10% by weight PEO to form nanofibers of 60 to 80 nm in diameter.37,39
The chitosan/PLA combination was first electrospun in 2009, by Jia Xu46, using TFA as co-solvent. With the introduction of PLA, the beads disappear gradually, but the diameter of the fibers decreased. The results of Fourier transform infrared spectroscopy (FTIR) demonstrated interaction between chitosan and PLA.37,46
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1.6 Electrospinning of PCL/Chitosan
Chitosan has been widely used in biomedical applications, but its poor mechanical properties have led to the need of mixing it with other types of polymers. PCL is a synthetic polymer with good mechanical properties, in addition to being biocompatible, biodegradable and non-toxic. The fact that it is very hydrophobic and induces poor cell adhesion may be overcome by blending it with another polymer, chitosan for example. As chitosan and PCL complement each other, it is expected that mixing the polymers will provide the adequate environment for the attachment, growth and differentiation of cells.
There are several reports available of PCL/chitosan nanofibers.
In 2009, Bhattarai et al.47, produced electrospun nanofibers using chitosan in TFA, 5wt%, and PCL in trifluoroethanol (TFE), 10wt%, in a ratio of 40/60 (chitosan / PCL). The results demonstrated excellent mechanical and biological properties in vitro. Those nanofibers were created to correct defects in nerve conduction.47
Using DMF and methylene chloride (MC) as solvents, Hong et al. (2011)48 manufactured a PCL/chitosan composite. There was an improvement in mechanical properties, with the Young's modulus increasing by 75% when compared to a pure PCL membrane. It also improved the hydrophilicity and interaction between the cells and the composite.48
In order to correct problems in peripheral nerve regeneration, Prabhakaran et al. (2008)49 designed nanofibrous PCL/chitosan scaffolds with fibers of 630, 450 and 190 nm diameters using HFIP, TFA e DCM. Again the mechanical and biological properties improved substantially when compared to pure chitosan.49
As can be seen, all the solvents used are organic. The use of this type of solvent causes serious problems such as toxicity, environmental impact, safety problems to workers in laboratories and the presence of toxic waste in electrospinning membranes.50 For biological applications, these problems prove to be even more critical and therefore their resolution is highly important. The solution was to create the concept of "Green electrospinning", which consists of the use of benign (potentially less toxic) solvents, for both the body and the environment. These should also allow electrospinning to be easily reproducible.51
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The first approach was to use water as a medium in electrospinning, followed by cross-linking of the membranes.51 However the difficulty of dissolving most polymers in water led to the use of other low toxic solvents such as acetone, acetic acid or formic acid.
Within this context, Shalumon et al. (2011)52 prepared a nanofibrous scaffold with fibers having a diameter of approximately 102 nm using acetone and formic acid as solvents. The obtained nanofibers indicated to be a good system for bone and skin tissues engineering.52 Schueren and Steyaert53,54 used a mixture of AA/FA to study the electrospinning of the PCL/chitosan mixture. In this study, the different proportions of the solvents in the unwinding of electrospinning were examined. The viscosity remained relatively equal, using different proportions of solvents, while the conductivity increased with increasing formic acid proportion. So, to get a stable Taylor Cone, AA/FA ratios chosen should be 3/7 and 5/5. Fiber diameters of respectively 203 ± 44 nm and 367 ± 118 nm were obtained (Fig. 10).54
Figure 10 - SEM images of PCL/Chitosan nanofibers with an AA/FA solvent ration of 3/7 (a) and 5/5 (b).54
It is also suggested that for the occurrence of a stable electrospinning the viscosity of the polymer solution should be between 1250 mPa s and 8000 mPa s.53,54
1.7 Three-dimensional architectures
Tissue engineering aims to rebuild, maintain or improve damaged or non functional tissues in the human body. In the last decade tissue engineering has grown so much that it offers solutions to regenerate almost all organs and tissues.55 For many parts of the body, a simple
21
2D structure, can be enough to support the formation of new tissue. But in others, a complex 3D network is needed for cell migration to take place.
The membranes obtained from electrospinning are 2D scaffolds, much used for example as films. These membranes are a well-compacted layer of nanofibers, which partially may restrict cell infiltration and growth, thereby limiting the connection between cells and organs due to their spatial organization, as well as their application in the human body.24,56
In order to circumvent this problem and better imitate the extracellular matrix, in recent years structures with a further dimension have been developed. Compared to a 2D structure, a 3D architecture provides another direction for cell proliferation, offering better interaction between cells and organs, and allowing long-term cell survival to increase. Several electrospinning approaches have been developed to produce 3D scaffolds of their nanofibers such as multilayering electrospinning, self-assembly, post-processing after electrospinning, template-assisted collection, liquid-assisted collection and adding porogen to electrospinning.57
However, these techniques often produce structures with low porosity, small pores (nanometer or less than 1 μm) and small pore distribution. Fibrous 3D structures must have porosity greater than 95%, fiber size between 1-1000 m and an isotropic structure in order to adequately mimic the extracellular matrix.58
In the case of the PCL/chitosan mixture, there are already scaffolds developed directly by 3D techniques, such as freeze-drying, particle-leaching and wet-spinning.29
With freeze drying, porous scaffolds were obtained using an aqueous solution of 25% acetic acid, and the pore morphology and stability depended on the mass ratio between PCL and chitosan. The mechanism that occurs in this process is a separation of the solid polymer from the solvent, thus forming crystals which, after being sublimated, create a porous network. SEM images (Fig. 11) showed that the pore morphology was open with sizes between 10-100 μm and that PCL appeared as globules in the network. The chitosan and PCL mixture in this study proved to be a promising combination to support cell growth in the 2D and 3D forms.59
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Figure 11 - SEM images of scaffolds of chitosan with (a) 50% PCL and (b) 75% PCL.59
Particle-leaching, using two different solvents, to manufacture 3D scaffolds is also reported in the literature. 59,60
For one study, the authors used a solution of acetic acid (80% v/v) to dissolve PCL and chitosan, to which salt particles were added. After washing the salt particles away, the obtained pores (Fig. 12) presented a square shape for PCL/CH25, sides with right angles for PCL/CH50, and completely irregular forms for PCL/CH75. These results demonstrated that with the same amount of salt particles, the morphology depended on the combination of polymer ratios. The pore sizes obtained varied from 30 to 280 μm and the scaffold had a porosity of approximately 70%.60
Figure 12 - SEM micrographs of scaffolds sections (a) PCL/CH25, (b) PCL/CH50, (c) PCL/CH75.60 In a second study hexafluoro-2-propanol and sodium chloride as porogen were used. Again the proportion of the mixture influenced the morphology of the structure, and the percentage of chitosan should not exceed 50% by weight, in order to maintain the strength of the scaffolds. The internal pores were interconnected with irregular shapes (Fig. 13) and had sizes between 40 and 270 μm.61
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Figure 13 - SEM micrographs of (a) PCL and (b) PCL/CH50 porous scaffolds.61
This technique proved to be more valid in controlling porosity as well as pore size.
At the University of Minho in Portugal, scaffolds were fabricated for cartilage repair through wet spinning. This technique consists of a polymer solution which is extruded directly into a chemical bath, causing the precipitation of fibers and then the solidification.62,28 In this study three polymer mixtures with different proportions in pure formic acid were used. The scaffolds obtained (Fig. 14) had a homogeneous surface distribution with porosity, pore sizes and good interconnectivity suitable for biomedical applications. The combination of 75/25 (chitosan / PCL) has been shown to be the best ratio mixing because of its final chemical, physical and biological properties.29
Figure 14 - SEM microphotographs of (a) CH, (b) PCL/CH75 and (c) PCL/CH50 fiber-mesh scaffolds.29
Although 3D nanofibrous structures are achieved, many improvements still need to be made in porous constructs, mechanical properties and structural features.5
24
1.8 Thermally induced self-agglomeration
Because the production of 3D structures directly from electrospinning is not feasible, we have moved on to a new type of approach. It has begun by transforming the membranes of electrospinning into small and short parts and only then uses them in the construction of 3D scaffolds.
This idea was created by Yang Si 62, who in 2014 manufactured superelastic 3D structures with ultra-low density, combining polyacrylonitrile (PAN)/SiO2-based nanofibers and the fibrous freeze-shaping method.63 Three-dimensional scaffolds were then produced for the first time from electrospun nanofibers, with desired density and shapes. However, its application in tissue engineering was not possible, mainly due to two reasons: the pores are very small, between 10-30 μm, for cell proliferation and the reagents and materials used are toxic to the body.63,64
Then in 2015, Gaigai Duan 64, using poly(methylacrylate(MA)-co-methyl methacrylate(MMA)-co-4-methacryloyloxybenzophenome(MABP), prepared ultralight polymer sponges of electrospun short fibers dispersed in dioxane followed by freeze-drying. The only drawback of this concept is that sponges are not biodegradable, and again, they cannot be used as scaffolds.64,65
Based on previous work, a group of scientists from the University of South Dakota, created a new approach, the thermally induced self-agglomeration (TISA), of nanofibers, to produce electrospun PCL 3D scaffolds.64
Through TISA, it is possible to produce nanofibrous scaffolds with high porosity and hierarchically interconnected pores through a very simple and convenient approach.
The TISA consists of a spontaneous agglomeration of the polymer in a gelatinous aqueous suspension followed by freeze drying. The procedure begins with the conversion of PCL nanofibers into small and short pieces, followed by a heat treatment that will promote their self-conglutination in situ. It ends with the removal of water by freeze drying.64
The final scaffold (Fig. 15) is elastic and smooth, having a porosity of approximately 96.4% and macropores with 300 μm of size. In vitro tests indicated high cell viability and bone marrow growth in rats. With these results, the authors demonstrated that the obtained 3D
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structures were functionally reliable for bone regeneration and therefore optimal synthetic extracellular matrix.64
Figure 15 – A photo of 3D nanofibrous scaffold of PCL with respective SEM images.64
In 2017, the same group of scientists created a combined 3D scaffold of PCL/PLA using TISA. Again, it was found that through this technique, highly porous structures, ~95.8%, with interconnected pores, ranging from sub-micrometers to 300 μm, were obtained and provided a good environment for the growth of cranial bone. The binding of PLA to PCL substantially improved the mechanical properties of the scaffold as well as its cell viability.24 Finally, this year, the same researchers produced PCL/HA composite scaffolds for the purpose of better application to the bone.66
1.9 Objectives
The present dissertation aims at the development of a 3D nanofibrous scaffold of a PCL/chitosan blend, through electrospinning followed by TISA, for tissue engineering. In this context, it is specifically envisaged:
i. Production of PCL/chitosan membrane by electrospinning; ii. Manufacture of a 3D nanofibrous scaffold by TISA;
iii. Optimization of the production parameters, both in electrospinning and TISA, in order to obtain the adequate characteristics in the scaffolds;
26
iv. Characterization both chemical and physical of the electrospun membranes, as well as the TISA scaffold.
29
Chapter II - Materials and
Methods
31
2.1 Materials and Reagents
Polycaprolactone (PCL, molecular weight Mn = 80 KDa, Ref. 440744) and medium molecular weight chitosan (75-85% deacetylation degree, Mn = 190 - 300 KDa, Ref. 448877), glacial acetic acid (99.8 v/v %) and formic acid (99-100 v/v %), purchased from Sigma-Aldrich, were the reagents used in the preparation of the polymer solution. In the TISA process, ethanol and gelatin (Ref. 04055) were used. The chemicals/solvents were used without further purification.
2.2 Preparation of solutions
Solutions with different concentrations of PCL and chitosan were prepared in a mixture of acetic acid (AA) / formic acid (FA) solvents with volume ratios of 1/1 and 1/2 (Table 1). The concentration of PCL was expressed as weight percent in the solution (wt. %), while the chitosan was expressed as a percentage relative to the mass of PCL (%). The formulas used to calculate the polymer concentrations were as follows:
) ( ) ( (%) PCL mass chitosan mass Chitosan = (1) ) ( ) ( ) ( ) ( .%) ( solvents mass chitosan mass PCL mass PCL mass wt PCL + + = (2)
Table 1 - Concentrations of polymer solutions.
Solvent AA/FA 1:1 (v/v) AA/FA 1:2 (v/v)
PCL (wt.%) 6 10 8 10
Chitosan (%) 10 10, 15 10, 15, 20 10, 15, 20
Samples were identified by the designation of the polymer followed by the respective percentage in table 1 (i.e. PCL6/CH10, PCL10/CH10 and so on).
32
The polymers solutions were warmed in a water bath at 40 °C under magnetic stirring until complete dissolution (approximately 2 hours). A pure PCL membrane (14% wt.%) was also prepared using AA/FA (1:2), at room temperature under stirring for 4 hours.
2.3 Production of PCL/Chitosan composite membrane by electrospinning
The nanofibrous membranes were produced in the FLUIDNATEK LE-10 (Fig. 16) equipment of the Department of Materials and Ceramics Engineering and the NANON-01A, MECC Co. Ltd. equipment of the TEMA research group. The equipments have the same characteristics, being the only difference the fiber production in the first being horizontal and in the second being vertical.Figure 16 - Components of FLUIDNATEK LE-10 electrospinning equipment.
After preparing the solution, it was placed in a 5 ml syringe and introduced into the infusion pump of the equipment to which a plastic tube with a stainless steel needle is attached to the tip. The distance from the collector to the tip of the needle was adjusted and the collector was covered with aluminum foil. Then the rotational speed of the collector (200 rpm), the flow rate and the voltage were selected. Table 2 summarizes the conditions used in electrospinning, as well as the environmental conditions. The fibers were collected randomly on the aluminum foil until a membrane was obtained which was easily removable.
33
Table 2 - Conditions of the electrospinning parameters.
Distance (cm) 13 and 15 Flow rate (L/h) 400-500
Voltage (kV) 16-30
Temperature (ºC) 20-25
Humidity (%) 30-50
2.4 Manufacture of a 3D nanofibrous scaffold
The membranes obtained by electrospinning were then transformed into 3D scaffolds through TISA. As the information on the technique is very scarce, a whole study on the conditions to be used had to be carried out. All the experimental steps performed with the optimized TISA conditions will be described below.
2.4.1 Conversion of membranes into small pieces
The PCL/chitosan membrane was first cut into pieces of approximately 1cm x 1cm and then soaked in a mixture of ethanol/water (50/50). Subsequently, the mixture was placed in a mortar with liquid nitrogen. When the pieces were frozen, they were ground mechanically and liquid nitrogen was added continuously to aid in the process. The pieces were then milled through a sieve (~ 1mm pore size) to make sure they all had a maximum area of 1mm2. The pieces that did not cross the sieve were again placed in the mortar to repeat the previous step, until the largest number of individualized and short nanofibers could be collected. The obtained mixture was then placed in a glass bottle and kept for 48 hours at room temperature so that the fibers deposited on the bottom of the bottle. The excess liquid at the top of the flask was then removed with the aid of a Pasteur pipette.
34
Figure 17 – A scheme of the steps followed in the conversion of membrane into small pieces.
2.4.2 Fabrication of 3D PCL/chitosan scaffolds
The obtained material was first mixed in an aqueous solution of gelatin (ethanol/DI water/gelatin) (4/2/1), which resulted in a mixture of dispersed polymer pieces in a somewhat viscous solution. This solution was then submerged in a 50 °C water bath for 2 to 5 minutes. During this time the nanofibers will agglomerate and form a 3D structure. Immediately afterwards, the flask was immersed in ice water for 30 minutes, so that the agglomerate did not shrink. The structure obtained was then rinsed 3 times with DI water to remove the gelatin and ethanol residues from the 3D agglomerate. It was then dispersed in DI water and taken to the freezer for 24 hours. The next day the sample was dipped in liquid nitrogen so that all of the water was frozen and freeze dried at room temperature for 24 hours.
35
2.5 Characterization
2.5.1 Scanning electron microscope (SEM) analysis
The morphologies of electrospun membranes and of 3D nanofibrous scaffolds were observed using a scanning electron microscope (SEM, Hitachi S-4100, Japan). The images were taken with an acceleration of 25kV at different magnifications.
Prior SEM analysis, all the samples were sputter coated with a layer of gold to improve their electrical conductivity. The diameters of the fibers were calculated using the software Image J from the SEM images.
2.5.2 Viscosity measurements of the solution
The viscosity of the solutions to be electrospun were measured with a Thermo ScientificTM
HAAKETMviscometer. The viscosity was measured at the cut-off speed of 116 s-1.
2.5.3 Fourier transform infrared (FTIR) spectroscopic measurements
Chemical analyses of the membranes were performed via attenuated total reflectance Fourier transform infrared (ATR-FTIR) in Bruker Tensor 27 FT-IR spectrometer (Bruker corporation). The spectra were recorded between 4000 and 400 cm-1, with a resolution of 4 cm-1 and 256 scans.
2.5.4 Wettability Tests
The wettability of the membranes was evaluated using the SL200HT Drop Shape Analytical System. A single drop of volume ~2 μl was poured on the surface. From the video made during the deposition, several images were captured and measured using the software Image J – contact angle. The measured angle is the one formed by the horizontal plane and the tangent line to the profile of the drop of water in the contact zone, as shown in figure 19.
36
Figure 19 - Contact angle measurement diagram.
2.5.5 Mechanical properties
The mechanical properties of the membranes and scaffolds were measured with a Shimadzu MMT-101 N (Shimadzu Scientific Instruments, Japan) (Figure 20) with a load cell of 100 N and a velocity of 1 mm min-1. For the calculation of tensile strength, 5 specimens were analysed. The samples thickness was measured using a micrometer with a precision of 1 μm. For each sample, the load versus cross-head displacement data from initial until maximum elongation of the equipment were measured using a PC data acquisition system connected to the tester.
37
For compression tests, 3 specimens of each sample were analysed. The samples were compressed after a pre-charge of 0.07N.
2.5.6 Differential scanning calorimetry (DSC) measurements
DSC analyses were done using Differential Scanning Calorimetry (DSC) named Shimadzu DSC-50 for PCL pellets, chitosan powder and different compositions of PCL/Chitosan membranes. For the first cycle, the heating range was from room temperature to 110ºC with heating rate of 20ºC/min. Then the second cycle was the cooling run with a cooling rate of 20ºC/min for temperature range from 110ºC to 20ºC. The same procedure was repeated for two times to obtain a total number of four cycles which consist of two heating and two cooling runs. The obtained DSC curves were used to analyze the melting temperature (Tm) of the samples.
2.5.7 Porosity
The porosity was calculated according to the following equation:
% 100 − = V V V Pscaffold p
where Pscaffold is the porosity of scaffold, V is the total volume of scaffold, and Vp is the
volume of nanofibers (i.e., the mass divided by the density, 1.145 g/cm3 for PCL, 1.158 g/cm3 for PCL/CH10, 1.164 g/cm3 for PCL/CH15 and 1.170 g/cm3 for PCL/CH20.
2.5.8 Micro-computed tomography (μCT) analysis
The three-dimensional internal and external morphology, the interconnectivity and the verification of the porosity of the scaffolds were performed by micro-computed tomography (μCT). The μCT imaging was carried out in a ShyScan microtomography model 1174v2 by Brucker Company (Brussels, Belgium) using a pixel size resolution of 14 μm and an exposure time of 7000 ms. The CT system was set with a rotation step of 0.9º, operating at 50 kV and 800 μA. Slices data sets were reconstructed in a NRecon software, and the visualization was performed in SkyScan Data Viewer program. The 3D virtual images were
38
built with CTvox software, and the porosity quantification were assessed through CTan software.
2.5.9 Swelling tests
For the swelling studies, dried scaffolds of each formulation were weighted (Wd) before immersion in distilled water for 48 hours at room temperature. After 15 min, 30 min, 1 h, 6h, 24h, 30h and 48h of immersion, the samples were weighted (wet, Ws). The superficial water was removed prior weighing with paper. The swelling ratio (Q) was obtained using the following equation:
d d s W W W Q=( − )
41
Chapter III - Results and
discussion
43
3.1 Electrospinning
In order to optimize the size and homogeneity of the diameter of the electrospun fibers, a study was carried out on two major variables: the operating conditions used in the electrospinning process and the properties of the solution. Regarding the properties, the parameters under study were the solvent ratio, the viscosity and the concentration. At the same time, the operational conditions were optimized, because both variables are dependent on each other.
3.1.1 Optimization of the operating conditions
In the electrospinning process, some operational conditions must be adjusted in order to optimize the characteristics of the fibers. The flow rate, the voltage, and the distance between the collector and the needle tip were the conditions tested in this work. Due to the complexity of the electrospinning process, each laboratorial test had to undergo an adjustment, in order to ensure the formation of the adequate Taylor cone. For each experiment, the distance was fixed and the values of the flow rate and of the voltage were adapted. Table 2 (in section 2.3) summarizes the interval values used.
Distance between the collector and the needle
Based on previous experimental work, two distances, 13 and 15 cm, were tested. The study was performed for all solution concentrations, but only the mixture of PCL10/CH10 was selected to present here. Figure 21 shows SEM images of the fibers collected at 13 cm (A, B) and 15 cm (C, D) distance from the collector to the needle tip. The production of the membranes proceeded without any adversity for both distances. Observing the images, it is clear that the distribution of the fibers is completely random for both samples and that the membranes contain some or no beads. The average diameters of the fibers were 350 nm and 150 nm for the distances of 13 and 15 cm, respectively. The decrease in the diameter is due to the fact that the fibers have to travel a larger path, thus allowing a greater evaporation of the solvent. This was observed for all samples, but with a decrease much lower.
44
Figure 21 – SEM images of PCL10/CH10 membranes at 13 cm (A, B) and 15 cm (C, D) distance from needle tip to collector.
In the next step (TISA), the membranes should present fibers diameters in a range between 50 nm and 1000 nm (i.e. 1 μm), but more preferably closer to 500 nm.67 Therefore, the nanofibers produced at 13 cm distance were chosen for the following studies.
3.1.2 Properties of the solution
Solvent ratio
Among the solvents commonly used to dissolve the polymers, acetic acid (AA) and formic acid (FA) are the most suitable due to their low toxicity and high reproducibility in electrospinning.54 In the work carried out by Schueren et al.53,54 the relationships 5/5 and 3/7 (AA/FA) were the ones that presented the best results. In the present work, the relations 1/1 (AA/FA) and 1/2 (AA/FA) were tested. In the preparation of both solutions, no difficulty in the dissolution of the polymers has occurred. As presented in table 1, for each solvents ration, different polymer concentrations were prepared.
45
For the experiments with the ratio 1/1 (AA/FA) the electrospinning process was unstable, with a bad formation of the Taylor cone. The solution of PCL6/CH10 produced non-individualized fibers with very small diameters, approximately 50 nm (Fig. 22 (A) and (B)). In order to increase the diameter, the concentration of the solution was increased to PCL10/CH10. The results obtained (Fig. 22 (C) and (D)) were significantly better, with the average fibers diameter increasing to 400 nm, and the fibers having a random and individualized morphology. With a higher ratio of chitosan, PCL10/CH15, the diameter decreased to 200 and the fibers showed some beads (Fig. 22 (E) and (F)), being therefore not suitable for TISA process.
Figure 22 – SEM micrographs of PCL/CH membranes of (A, B) PCL6/CH10, (C, D) PCL10/CH10 and (E, F) PCL10/CH15 using a solvent ratio of 1/1.
For the ratio, 1/2 (AA/FA), the increase of the formic acid amount is expected to lead to a more stable electrospinning process. As acetic acid has a low dielectric constant (6.6 0) and formic acid a high dielectric constant (57.2 0), increasing the content of formic acid helps counteract the distribution of the electric field that is applied.68 All electrospinning performed with this solvent ratio occurred much more easily, with a noticeable formation of the Taylor's cone. From the SEM images (Fig. 23), it was observed that the concentration PCL10/CH10, which worked for the ratio 1/1, also worked for the ratio 1/2. The diameter and morphology of the fibers remained in the same range for this new solvent ratio. When the concentration was increased, the electrospinning was possible and fibers without beads
