Development of electrospun dressings for infected wounds

Swinburne University of Technology

Faculty of Science, Engineering and Technology

Development of

electrospun dressings

for infected wounds

A thesis submitted for the degree of

Doctor of Philosophy

By

Martina Abrigo

January 17, 2016

Abstract

Chronic non-healing wounds show delayed and incomplete healing pro-

cesses and in turn, expose patients to a high risk of infection. Promising can-

didates for treating these wounds are polymeric micro/nanofibrous meshes,

but the interactions that occur between bacteria and fibres with different

morphological and physico-chemical properties need to be better understood.

In the present work, an electrospinning apparatus was designed and fabri-

cated to manufacture micro/nanofibrous polystyrene meshes with controlled

morphology. Different chemical functionalities were generated on the surface

of the meshes through plasma polymerisation of four monomers (acrylic acid,

allylamine, 1,7-octadiene and 1,8-cineole).

The influence of fibre size and surface chemistry on the attachment and pro-

liferation of Escherichia coli, Pseudomonas aeruginosa, and Staphylococcus

aureus was investigated using a combination of techniques, including viabil-

ity assays, and confocal and scanning electron (SEM) microscopy.

Fibre diameter close to the bacterial length induced the highest proliferation

rates, while nanofibres were found to cause conformational changes of rod

shaped bacteria, limiting the colonisation process.

Fibre wettability, surface charge and chemistry were found to influence the

ability of E.coli cells to transfer, attach and proliferate onto and within the

meshes. The hydrophilic amine rich coating showed the highest proportion

of viable cells transferred from underlying agar cultures. The same chemistry

was also found to attract P.eruginosa cells cultured in tissue engineered mod-

els of human skin. These models were developed by co-culturing skin cells

(fibroblasts and keratinocytes) in human skin grafts to reproduce wounds

at different depths and degrees of severity. The transfer of fibroblasts and

i / 223

keratinocytes from the wound models onto the plasma polymerised meshes

was investigated, since skin cell transfer and ingrowth into the dressing has

to be prevented to avoid wound reopening upon dressing removal. The octa-

diene coating induced the least degree of fibroblast removal, while the acrylic

acid and allylamine chemistries remained partially adhered within the wound

models.

The significant innovative contribution of this research work exists in the de-

sign, development and in-vitro testing of various solutions that can address

some of the major challenges associated to chronic wound care. Results sug-

gest that fibre diameter and surface chemistry could be strategically tuned

for controlling the bacterial load in the wound bed. Depending on the type

and severity of the wound to be treated, various surface chemistry options

were found successful for preventing skin cell transfer and ingrowth.

ii / 223

Acknowledgements

I want to take the opportunity to sincerely thank the people who sup-

ported me throughout my research journey and shared with me unforgettable

moments.

I want to start thanking Swinburne University of Technology for giving me

the opportunity to undertake my doctoral degree, and for the efficient ser-

vices provided to international students, that have been extremely valuable

to make the best out of my overseas experience in Australia.

Thank you to my supervisor, Prof. Sally McArthur, for being always present,

guiding me towards the most rewarding directions, transmitting me her en-

thusiasm and passion for science and encouraging me to take the best deci-

sions for my career and personal future.

Thanks to Prof. Peter Kingshott for the knowledge and experience he has

shared with me to overcome the most challenging obstacles I have encoun-

tered during my research.

Thank you to Andrew Moore for his significant contribution to the instal-

lation of the first electrospinning machine at Swinburne University, which

would not have been possible without his creativity and talent.

Special thanks to Prof. Sheila MacNeil and Prof. Ian Douglas for welcoming

me at The University of Sheffield and making me become part of their pres-

tigious research groups.

Thanks to Dr. Anthony Bullock for guiding me through the world of cell

biology while teaching me some authentic English humour. A special thank

goes to Dr. Marc Daigneault who supported and helped me during all my

time at Sheffield and became a good friend.

Thanks to Dr. Scott Wade, Dr. Thomas Ameringer, Dr. Michelle Dunn,

iii / 223

Dr. Mya Hlaing, Dr. Mirren Charnley, Dr. Nick Reynolds and all the other

scientists and researchers at Swinburne University with whom I have shared

moments of my research journey. Special thanks to A/Prof. Paul Stoddart

for mentoring me and for the Christmas in July parties, which I will dearly

remember.

Thanks to Hannah and Dori, for all the fun we had together and the support

we gave one another.

Chiara and Benoit, thank you for the good time spent together, the dinners,

the travels, the talks about the future. Thank you for being true friends.

I would like to deeply thank Trevor, Sue, Murray and Ruth for making me

feel part of their families when I was missing mine very much. Thank you

for your help and your friendship.

Thank you to Francesca, Charlotte, Irene and Cristina, my best friends, be-

cause nothing will ever change among us, wherever will we be. Special thanks

to Giulia, who has always been next to me, thanks for your advice and your

encouragement.

My warmest thoughts go to my family.

Nonni Marina, Pierino, Rosa, e Mauro, thank you for being always present

in my life and for your interest in all what I do. I missed you every day

more in the past years. Thanks to Carla, Elena, Claudio, Corrado, Roberto,

Laura, e Alfredo, for the invaluable support and the sweet welcome you give

me every time I come back.

Thanks Michelle, Claude and all Lapierre family for the great holiday time

spent together, and for caring much about me.

Florian, thank you for being next to me every day, with your smile, your

energy and your patience. Thanks for believing in me, for helping me be

strong and go through hard times, and for sharing with me many unforget-

iv / 223

table joyful moments. Thanks for being my anchor.

Mamma e Papa, thank you for encouraging me to follow my dreams, for

being my most important confidants, for accompanying me through every

choice, success, and difficulty. Thanks for teaching me how to grow up, how

to achieve my goals without ever forgetting my roots. Enrica, thanks for

being with me whenever I need, for listening to me, advising me, making me

laugh, and for sharing your feelings and fears. I know that regardless the

distance and the time that flies, we will always be together, counting on each

other.

v / 223

Declaration

I, Martina Abrigo, declare that the work presented in this thesis is, to the

best of my knowledge and belief, original, except as acknowledged in the text,

and that the material has not been submitted, either in whole or in part, for

another academic award at this or any other university.

I acknowledge that I have read and understood the Universitys rules, require-

ments, procedures and policy relating to my higher degree research award

and to my thesis. I certify that I have complied with the rules, requirements,

procedures and policy of the University.

Martina Abrigo

Industrial Research Institute Swinburne

Faculty of Science, Engineering and Technology

Swinburne University of Technology

Dated this day, January 17, 2016

vi / 223

Nomenclature

AFM Atomic force microscopy

CFU Colony-forming unit

CTAB Cetyltrimethylammonium bromide

CTAB cetyltrimethylammonium bromide

CV Crystal violet

DED De-epidermised dermis

H&E Haematoxylin Eosin

HTAB Hexadecyltrimethylammonium bromide

HV High voltage

MQ Milli-Q water

MTS 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium

MTT 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide

MW Molecular weight

N − C Needle-collector distance

PBS Phosphate buffered saline

PI Propidium iodide

ppAAc Plasma polymerised acrylic acid

ppAAm Plasma polymerised allylamine

ppCo Plasma polymerised 1,8-cineole

ppOct Plasma polymerised 1,7-octadiene

PS Polystyrene

SDS Sodium dodecyl sulfate

SEM Scanning electron microscopy

vii

STS Split Thickness Skin

XPS X-ray photoelectron spectroscopy

viii / 223

Contents

Nomenclature vii

Introduction 1

1 Literature review 7

1.1 The structure and function of human skin . . . . . . . . . . . 8

1.1.1 Physiological wound healing . . . . . . . . . . . . . . . 9

1.1.2 Chronic wounds . . . . . . . . . . . . . . . . . . . . . . 11

1.2 Wound dressings . . . . . . . . . . . . . . . . . . . . . . . . . 14

1.3 Nanofibrous meshes . . . . . . . . . . . . . . . . . . . . . . . . 22

1.3.1 The electrospinning techniques . . . . . . . . . . . . . 23

1.3.2 Control over the morphology of electrospun fibres . . . 25

1.4 Electrospun meshes as wound dressings . . . . . . . . . . . . . 29

1.5 Controlling biological interaction with electrospun meshes . . . 38

1.5.1 How Do Bacteria Respond to Nanofibrous Meshes? . . 38

1.5.2 Role of fibre size and surface chemistry . . . . . . . . . 40

1.6 Surface modification strategies . . . . . . . . . . . . . . . . . . 41

1.7 Biological responses to plasma polymerised surfaces . . . . . . 45

1.7.1 Bacterial interactions with plasma polymerised surfaces 45

1.7.2 Skin cell interactions with plasma polymerised surfaces 46

ix

CONTENTS

1.8 In vitro Wound Models . . . . . . . . . . . . . . . . . . . . . . 48

1.9 Aims & objectives . . . . . . . . . . . . . . . . . . . . . . . . . 51

2 Experimental methods and techniques 53

2.1 Electrospinning . . . . . . . . . . . . . . . . . . . . . . . . . . 54

2.1.1 Electrospinning apparatus . . . . . . . . . . . . . . . . 54

2.1.2 Fibre fabrication . . . . . . . . . . . . . . . . . . . . . 58

2.2 Plasma polymerisation . . . . . . . . . . . . . . . . . . . . . . 61

2.3 Bacterial culture techniques . . . . . . . . . . . . . . . . . . . 64

2.4 Cell culture techniques . . . . . . . . . . . . . . . . . . . . . . 66

2.5 Wound models . . . . . . . . . . . . . . . . . . . . . . . . . . 69

2.5.1 De-epidermisation of STS . . . . . . . . . . . . . . . . 70

2.5.2 Decellularisation of STS . . . . . . . . . . . . . . . . . 71

2.5.3 Model of superficial partially de-epidermised wounds . 71

2.5.4 Model of superficial de-epidermised wounds . . . . . . 72

2.5.5 Model of deep wounds . . . . . . . . . . . . . . . . . . 73

2.5.6 3-Dimensional deep infected wound . . . . . . . . . . . 74

2.6 Characterisation . . . . . . . . . . . . . . . . . . . . . . . . . . 76

2.6.1 Physico-chemical characterisation . . . . . . . . . . . . 77

2.6.2 Biological characterisation . . . . . . . . . . . . . . . . 79

3 Electrospinning of polystyrene meshes 85

3.1 Optimisation of electrospinning parameters . . . . . . . . . . . 86

3.2 Electrospinning of nanofibres . . . . . . . . . . . . . . . . . . . 100

3.3 Characterisation of electrospinning apparatus performance . . 106

3.4 Electrospinning of aligned fibres . . . . . . . . . . . . . . . . . 109

3.5 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111

x / 223

CONTENTS

4 Plasma polymerisation of electrospun meshes 113

4.1 Characterisation of plasma polymerised meshes . . . . . . . . 114

4.1.1 Surface morphology of plasma polymerised meshes . . . 114

4.1.2 Surface chemistry of plasma polymerised meshes . . . . 116

4.1.3 Aging study on ppAAm coating . . . . . . . . . . . . . 122

4.2 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 124

5 Interactions of wound bacteria with electrospun meshes 127

5.1 Bacterial colonisation of electrospun meshes . . . . . . . . . . 129

5.2 Influence of fibre diameter on bacterial behaviour . . . . . . . 133

5.3 Influence of fibre surface chemistry on bacterial behaviour . . 146

5.3.1 Bacterial transfer onto ppAAm coated meshes . . . . . 154

5.4 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . 156

6 Skin and bacterial cells transfer onto electrospun meshes 159

6.1 Cell transfer studies . . . . . . . . . . . . . . . . . . . . . . . . 160

6.2 Wound models . . . . . . . . . . . . . . . . . . . . . . . . . . 168

6.2.1 Superficial partially de-epidermised wound . . . . . . . 170

6.2.2 Superficial de-epidermised wound . . . . . . . . . . . . 173

6.2.3 Deep wound . . . . . . . . . . . . . . . . . . . . . . . . 175

6.2.4 3-Dimensional deep infected wound . . . . . . . . . . . 178

6.3 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . 185

7 Conclusions 189

7.1 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189

7.2 Outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 191

References 193

xi / 223

Appendix 217

Research Achievements 219

List of Figures

1.1 Schematic representation of the layered structure of the human skin. 8

1.2 (a) Application fields of electrospun nanofibres targeted by US patents

and (b) potential application of electrospun polymeric nanofibres [1]. 26

1.3 Schematic representation of electrospinning basic set up. (a) Image

of Taylor cone forming at the spinneret during the electrospinning

process [2]; (b) Image of polymeric filament forming from Taylor

cone and moving toward the collector [3]. . . . . . . . . . . . . . . 27

1.4 Schematic representation of electrospinning collectors: (a) planar

collector for non-woven meshes; (b) square frame for unidirectional

oriented fibres; (c) cylindrical collector for tubular oriented fibres;

(d) Non-woven fibres fabricated using collector (a); (e) aligned fibres

that can be fabricated using collectors (b) or (c). . . . . . . . . . . 28

1.5 Approaches for surface modification of electrospun fibres: (a) plasma

polymerisation; (b) wet chemical method; (c) surface graft poly-

merisation; and (d) co-electrospinning [4]. . . . . . . . . . . . . . . 43

2.1 Schematic representation of the assembly of the components consti-

tuting the electrospinning set up. . . . . . . . . . . . . . . . . . . . 56

xiii

LIST OF FIGURES

2.2 Photograph of the rotating mandrel, used to collect fibres aligned

along one direction. The black and red cables connect the motor to

a power supply, while the green cable ensures the connection of the

collector to the ground. . . . . . . . . . . . . . . . . . . . . . . . . 57

2.3 Photograph of the electrospinning set up after installation. The red

cable terminating with a crocodile clip connects the needle to the

HV power supply, positioned on the top of the safety box. The

needle is held in place by a perspex support connected to a wood

system that allows to move the needle horizontally and vertically.

The teflon tube connecting the syringe to the needle passes through

a hole drilled in the safety box. . . . . . . . . . . . . . . . . . . . . 58

2.4 Photograph of the plasma polymerisation reactor. The pressure in

the reactor chamber is brought to 1x10−3 mbar through the vacuum

pump. The needle valve constitutes the inlet for the volatilised

liquid monomer into the chamber. The plasma is generated when

an electric field at radio frequency (13.56 MHz) is ignited through

the electrode, producing a glow discharge that ionises a fraction of

the molecules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62

2.5 Schematic representation of the plasma polymerisation process of

the electrospun meshes. When the monomer is introduced in the

chamber, the ignition of the electric field generates electrons, ions,

free radicals, photons and molecules in both ground and excited

states. The reactive species impinge on the surface of the substrate

creating reactive sites within the plasma zone which are available

for the covalent attachment of other species. . . . . . . . . . . . . . 63

2.6 Schematic representation of the bacterial agar culture experiment

designed to investigate the transfer of bacterial cells onto and within

electrospun meshes. . . . . . . . . . . . . . . . . . . . . . . . . . . . 65

xiv / 223

LIST OF FIGURES

2.7 Schematic illustrating the well plate transfer experiment: (a) mesh

exposed to a confluent layer of primary human dermal fibroblasts.

A metal grid was used to hold the mesh in contact with the culture;

(b) mesh exposed to a confluent layer of primary human dermal

keratinocytes. A metal ring was used to hold the mesh in contact

with the culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70

2.8 Schematic illustrating the preparation of the tissue engineered skin

models of: (a) superficial partially de-epidermised wound; (b) su-

perficial de-epidermised wound; (c) deep wound. . . . . . . . . . . 72

2.9 Image processing steps for the quantification of bacterial cells at-

tached onto the meshes: (a) projection of the z-stack image along

the z-axis; (b) application of threshold to isolate bacteria from fibres

and noise; (c) particle counting and outlines. Scale bar µm. . . . . 81

3.1 SEM images of PS fibres electrospun from 35% w/v solution in

chloroform at different magnification. Scale bar: (a) 10 µm; (b) 2

µm; and (c) 1 µm. . . . . . . . . . . . . . . . . . . . . . . . . . . . 90

3.2 SEM images of PS fibres electrospun from 35% w/v solution in DMF

at different magnifications. Scale bar: (a) 10 µm; (b) 2 µm; and (c)

1 µm. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91

3.3 Photograph of the electrospun mesh obtained from PS solution in

DMF (35% w/v). Rhodamine was added to DMF (1% w/v) prior

electrospinning for imaging purposes. After 5 minute electrospin-

ning a mesh with approximate square shape, surface of about 30 x

30 cm2 and thickness of 1-2 mm was obtained. Scale bar 10 mm. . 92

3.4 (a) Bright microscopy, (b) SEM and (c) AFM images of fibres elec-

trospun from 35% w/v PS solution in DMF. Scale bar (a) 50 µm;

(b) 10 µm; and (c) 100 nm. . . . . . . . . . . . . . . . . . . . . . . 93

xv / 223

LIST OF FIGURES

3.5 SEM images of PS meshes electrospun from solutions in DMF at

different concentrations: (a) C = 10% w/v, Φ = 300 ± 200 nm; (b)

C = 15% w/v, Φ = 900 ± 200 nm; C = 20% w/v, Φ = 1000 ± 100

nm; C = 30% w/v, Φ = 3000 ± 1000 nm. Scale bar 2 µm. . . . . . 94

3.6 SEM images showing the morphology and size of the beads along

fibres electrospun from (a) 10% w/v; (b) 15% w/v; and (c) 20% w/v

PS solution in DMF. Scale bar 1 µm. . . . . . . . . . . . . . . . . . 95

3.7 Influence of applied voltage and N-C distance on the average fibre

diameter of electrospun PS meshes. . . . . . . . . . . . . . . . . . . 96

3.8 Average fibre diameter of the meshes electrospun from 15 and 20%

w/v PS solutions in DMF before and after the addition of CTAB

and SDS. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103

3.9 SEM micrographs of electrospun meshes obtained from: 15% w/v

PS in DMF with the addition of (a) CTAB; (b) SDS; and 20% w/v

PS in DMF with the addition of (c) CTAB; (d) SDS. The diameter

of the single fibres (in red) is expressed in nm. Scale bar 1 µm. . . 104

3.10 Average fibre diameter of the meshes electrospun from 15% w/v PS

solutions in DMF at a time distance of one month. . . . . . . . . . 107

3.11 Graph showing the weight of the meshes after different times of

electrospinning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 108

3.12 SEM images of PS meshes electrospun on the rotating mandrel at

two rotational speeds: (a) 500 rpm; (b) 2500 rpm. Scale bar 10 µm. 110

4.1 SEM microgaphs of (a) untreated; (b) air plasma treated; (c) ppAAc;

(d) ppCo; (e) ppOct; and (f) ppAAm plasma coated PS fibres. Scale

bar 1 µm . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115

4.2 XPS high-resolution carbon 1s spectra of untreated and plasma

polymerised PS meshes . . . . . . . . . . . . . . . . . . . . . . . . 117

xvi / 223

LIST OF FIGURES

4.3 XPS wide scan spectra of the uncoated PS mesh and the plasma

polymerised meshes . . . . . . . . . . . . . . . . . . . . . . . . . . . 119

4.4 XPS high-resolution carbon 1s spectra of (a) untreated PS and (b)

ppAAc coated meshes with fitted curves . . . . . . . . . . . . . . . 120

4.5 (a) Elemental composition and (b) oxygen/carbon and nitrogen/carbon

atomic ratios of the ppAAm caoted meshes from day 0 until 22 days

after plasma polymerisation . . . . . . . . . . . . . . . . . . . . . . 123

5.1 Electrospun PS meshes stained through the MTT assay. (a) Con-

trol mesh, not exposed to bacterial culture; (b) Mesh exposed to

bacterial culture for 1 hour. Scale bar 1 cm . . . . . . . . . . . . . 129

5.2 E.coli cells onto electrospun PS fibres after incubation for (a) 30

min; (b) 1hr; (c) 2hrs; (d) 4 hrs; and (e) 6 hrs. Scale bar: (a) and

(b) 1 µm; (c), (d) and (e) 2 µm. . . . . . . . . . . . . . . . . . . . 130

5.3 SEM of bacteria colonising PS electrospun mesh. (a) The elongated

bacterium pointed by the red arrow is in the elongation configura-

tion occurring during the binary fission process; (b) two bacterial

cells after cell fission, ready to divide. Scale bar 1 µm. . . . . . . . 131

5.4 Bacterial solution culture experiment. Confocal (a; c; e) and SEM

(b; d; f) images of E. coli cells colonising PS electrospun meshes

with fibre diameter ranges: (a, b) Φ1 = 500 ± 200 nm; (c; d) Φ2 =

1000 ± 100 nm; (e; f) Φ3 = 3000 ± 1000 nm. Scale bar: (a), (c),

and (e) 5 µm; (b), (d), and (f) 2 µm. . . . . . . . . . . . . . . . . . 136

5.5 SEM of single E. coli cells (false coloured in red) adhered onto PS

electrospun fibres with diameter: (a) 0.3 µm (b); 1 µm; (c) 5 µm.

Scale bar 1 µm. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137

5.6 Bacterial agar culture experiment. Crystal violet staining of agar

cultures after mesh removal: (a) E. coli ; (b) P. aeruginosa; and (c)

S. aureus. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139

xvii / 223

LIST OF FIGURES

5.7 Bacterial agar culture experiment. Confocal (a; c; e) and SEM (b;

d; f) images of E. coli cells colonising PS electrospun meshes with

fibre diameter ranges: (a; b) Φ1 = 500 ± 200 nm; (c; d) Φ2 = 1000

± 100 nm; (e; f) Φ3 = 3000 ± 1000 nm. Scale bar: (a), (c), and (e)

5 µm; (b), (d), and (f) 2 µm. . . . . . . . . . . . . . . . . . . . . . 140


d; f) images of P. aeruginosa cells colonising PS electrospun meshes

with fibre diameter ranges: (a; b) Φ1 = 500 ± 200 nm; (c; d) Φ2 =

1000 ± 100 nm; (e; f) Φ3 = 3000 ± 1000 nm. Scale bar: (a), (c),

and (e) 5 µm; (b), (d), and (f) 2 µm. . . . . . . . . . . . . . . . . . 141


d; f) images of S. aureus cells colonising PS electrospun meshes with

fibre diameter ranges: (a; b) Φ1 = 500 ± 200 nm; (c; d) Φ2 = 1000

± 100 nm; (e; f) Φ3 = 3000 ± 1000 nm. Scale bar: (a), (c), and (e)

5 µm; (b), (d), and (f) 2 µm. . . . . . . . . . . . . . . . . . . . . . 143

5.10 Photograph of the plasma coated and silver releasing meshes ex-

posed to E. coli layer. . . . . . . . . . . . . . . . . . . . . . . . . . 147

5.11 Confocal images of LIVE/DEAD stained E.coli cells onto (a) un-

treated PS mesh; (b) ppAAc; (c) ppAAm; (d) ppOct; and (e) ppCo

meshes after removal from the E.coli agar culture. Scale bar 5 µm. 148

5.12 SEM images of (a) untreated PS; (b) ppAAc; (c) ppAAm; (d)

ppOct; and (e) ppCo coated meshes after removal from the E.coli

agar culture. E.coli cells were false coloured in red. Scale bar 2 µm. 149

5.13 Quantification of E.coli cells that transferred onto the ppAAm coated

meshes from the agar plates at different culturing conditions. . . . 155

6.1 Bright-field optical microscopy images images of 70-80% confluent

cultures of (a) human dermal fibroblasts; (b) human dermal ker-

atinocytes. Scale bar 100 µm. . . . . . . . . . . . . . . . . . . . . . 161

xviii / 223

LIST OF FIGURES

6.2 Photographs of (a) electrospun meshes; (b) 6 well plate after fibrob-

last transfer experiment. The viable fibroblast cells that transferred

on the meshes or remained on the plates are stained purple. . . . . 162

6.3 Absorbance at 570 nm of the MTT dye dissolved from the fibroblast

cultured well plates and dressings. . . . . . . . . . . . . . . . . . . 163

6.4 Photographs of (a) electrospun meshes; (b) 12 well plate after ker-

atinocyte transfer experiment. The viable keratinocyte cells that

transferred on the meshes or remained on the plates are stained

purple. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 164

6.5 Absorbance at 570 nm of the MTT dye dissolved from the HDK

cultured well plates and dressings. . . . . . . . . . . . . . . . . . . 165

6.6 H & E histology images of the skin specimens (a) before and (b) after

the de-cellularization and de-epidermization of the split thickness

skin grafts. Scale bar 20 µm . . . . . . . . . . . . . . . . . . . . . . 170

6.7 H & E histology image of the superficial partially de-epidermised

wound model. Scale bar 20 µm . . . . . . . . . . . . . . . . . . . . 171

6.8 Photographs of (a) skin specimens; (b) electrospun meshes after

MTT assay. No transfer of viable cells occurred from the skin spec-

imens onto the meshes. The wound model was developed to mimic a

superficial partially de-epidermized wound, with keratinocytes dif-

ferentiating above the BM and fibroblasts spread through the dermis.172

6.9 Photographs of the skin specimen onto which the ppAAc mesh was

placed. (a) The purple ring corresponding to an area of viable cells

can be visualised; (b) the edges of the skin specimen and of the fibre

layers that remained adhered onto the skin after mesh removal are

underlined in blue and red respectively. . . . . . . . . . . . . . . . 172

6.10 H & E histology image of the superficial de-epidermised wound

wound model. Scale bar 20 µm . . . . . . . . . . . . . . . . . . . . 174

xix / 223

LIST OF FIGURES


MTT assay. No transfer of fibroblast cells occurred from the skin

specimens onto the meshes. The wound model was developed to

mimic a superficial de-epidermized wound, with fibroblasts spread

through the dermis, underneath the BM. . . . . . . . . . . . . . . 175

6.12 H & E histology image of the deep wound wound model. Scale bar

20 µm . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 176


MTT assay. Transfer of fibroblast cells from the skin specimens

onto the meshes occurred. The wound model was developed to

mimic a deep wound, with loss of BM. . . . . . . . . . . . . . . . . 177

6.14 Histology images of the 3-dimensional wound model: (a) H & E

stained section of the skin composite, before generating the burn.

Scale bar 50 µm; (b-e) Gram stained tissue sections, showing the

progressive development of the model, from (b) epidermis formation;

(c) thermal burn; (d) P.aeruginosa biofilm formation. Scale bar 100

µm; (e) magnification over the bacteria cells forming the biofilm.

Scale bar 20 µm. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 179

6.15 PI stained (a) fibroblast and (b) P.aeruginosa cells attached onto

PS meshes. Scale bar 20 µm . . . . . . . . . . . . . . . . . . . . . . 181

6.16 Graph showing the number of P.aeruginosa cells counted on the

meshes on each of the three experiments. . . . . . . . . . . . . . . 182

6.17 Graph showing the quantification of P.aeruginosa cells remained

on the 3D wound model specimens after the removal of the plasma

coated meshes and the PS control mesh. . . . . . . . . . . . . . . . 183

xx / 223

LIST OF FIGURES

7.1 Example of the imageJ code that was developed for an automatic se-

lection and counting of the particles present on the confocal images

of the meshes tested on the infected wound model. Each parti-

cle counted by the software corresponds to a PI stained bacterial

cell. The code implements the z-projection of each image, threshold

adjustment and particle counting. The comments describing each

function of the code are shown in green. . . . . . . . . . . . . . . . 217

xxi / 223

LIST OF FIGURES

xxii / 223

List of Tables

1.2 Summary of the phases constituting the physiological healing pro-

cess of a wound [5]. . . . . . . . . . . . . . . . . . . . . . . . . . . . 13

1.3 Classification of commercially available wound dressings . . . . . . 21

1.4 Ideal properties of nanofibrous meshes for wound healing applica-

tions [1, 6–8]. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23

1.6 Selection of the most frequently used synthetic and natural polymers

for fabrication of nanofibrous wound dressing by the electrospinning

technique and their advantages and disadvantages [9–12]. . . . . . 31

1.7 Surface modification techniques of electrospun nanofibres . . . . . 44

2.1 Molecular formula and structure of the monomers used for the

plasma polymerisation of the PS meshes. . . . . . . . . . . . . . . . 61

3.1 Process parameters tested for the electrospinning of PS dissolved in

chloroform and DMF at different concentrations. . . . . . . . . . . 89

3.2 Solution parameters and average fibre diameter of electrospun polystyrene

solutions containing hexadecyltrimethylammonium bromide. . . . . 101

3.3 Solution conductivity and average fibre diameter obtained after the

addition of CTAB and SDS surfactants to 15% and 20% w/v PS

solutions in DMF. . . . . . . . . . . . . . . . . . . . . . . . . . . . 102

xxiii

LIST OF TABLES

4.1 XPS theoretical and measured atomic composition and atomic ra-

tios relative to the total concentration of carbon (O/C and N/C)

of the uncoated and plasma coated meshes. The measured values

are the mean values ± deviation based on +3 analyses performed

on each sample. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118

xxiv / 223

Introduction

Chronic non-healing wounds, such as diabetic foot ulcers, pressure ulcers

and venous leg ulcers exhibit a pathologically delayed healing process, re-

maining open or partially healed for several weeks or months [5]. These

wounds are characterized by the presence of persistent inflammatory stimuli

which interrupt the physiological healing mechanisms and they can become

contaminated by a complex population of many different bacteria. These

pathogens lead to the development of infection through the formation of a

biofilm, that isolates bacteria from the immune system and can develop high

resistance against antibacterial agents, which in turn leads to higher risk for

systemic infection [13].

Currently, chronic wounds are treated with a broad variety of dressings tai-

lored to the requirements of the wound (dry or exuding, clean or infected, su-

perficial or deep) [14, 15]. Although wound management has significantly im-

proved in the last decades, a treatment capable of effectively healing chronic

wounds while tackling infection does not exist.

The manufacture of polymeric micro/nanofibrous meshes is central to the de-

velopment of many wound dressings. These structures are made of ultra-fine

fibres with diameters ranging from several micrometers down to few nanome-

ters. Several intrinsic properties of micro/nanofibrous meshes, such as high

surface area and microporosity, make these structures particularly interest-

1

Introduction

ing for wound healing applications [10]. Various techniques, including phase

separation or self-assembly, can be used for the fabrication of the meshes, but

electrospinning is most frequently chosen because it is a simple, cost-effective

and versatile process [16].

Electrospun scaffolds in the form of two dimensional non-woven meshes have

been shown to be promising candidates as wound dressings because they

promote haemostasis, fluid absorption, cell respiration and gas permeation

when implanted onto open wounds [8]. Ideally, the meshes should be able to

actively initiate the healing processes, while reducing the bacterial contami-

nation and treating infection only if necessary. Since the dressing is designed

to be removed once the wound has healed, the mesh should promote cell

migration and proliferation within the wound bed while preventing tissue

ingrowth within the fibrous structure.

Different strategies are currently used by researchers to create electrospun

meshes with the ability to assist the healing processes while preventing

wound infection. Various synthetic and natural polymers can be combined

to develop materials that actively support and supplement the deposition

of healthy tissue. The literature also significantly focuses on incorporating

drugs, silver nanoparticles and plant-derived compounds, including essential

oils and honey, which exhibit antimicrobial properties [17].

Although these strategies are very promising for achieving a multifunctional

effective device, still multiple challenges need to be overcome. To our knowl-

edge, despite bacterial infection representing a major challenge in chronic

wound care, studies on the mechanisms of adhesion, spreading and colonisa-

tion of electrospun meshes by bacteria do not exist.

Bacterial attachment on flat substrates and the factors that influence this

process have been widely investigated and it is currently recognised that

2 / 223

Introduction

apart from cell surface characteristics, bacterial attachment mechanisms are

also regulated and influenced by a wide range of substratum properties, such

as morphology, surface chemistry and roughness [18]. The influence of fi-

bre morphology and surface chemistry on microbial behaviour needs to be

investigated as this knowledge could lead to tailoring either physically or

chemically the properties of fibre surfaces to specifically address microbial

behaviour.

Another important aspect, which has not been deeply explored yet is the ef-

fects induced by electrospun meshes on skin cells and bacteria in co-cultures.

The most complete studies on electrospun wound dressings involve separate

in vitro experiments on skin cells and bacteria. These tests constitute valu-

able tools for studying the cytotoxicity as well as antimicrobial activity of

electrospun meshes. However, they are performed in highly defined and con-

trolled culture conditions, which do not reproduce the real environment of

chronic wounds [19]. Strategies for establishing in vitro chronic wound mod-

els by co-culturing various types of skin cells and bacteria can be found in the

literature, but those models have not yet been used for testing electrospun

dressings.

In this thesis, the responses of wound bacteria and skin cells to electrospun

polystyrene (PS) meshes with different morphological and surface chemistry

properties were investigated. PS was chosen as a model system being the

standard material used to fabricate tissue culture plates for in vitro cell and

bacterial culture; moreover PS is a non degradable synthetic polymer which

allowed to study the role of fibre morphology preventing additional uncer-

tainties, such as polymer degradation that occurs in presence of materials

most frequently chosen for wound healing meshes, such as poly(lactic acid),

poly(glycolic acid) and copolymers [20].

3 / 223

Introduction

The influence of fibre size on initial attachment and spreading of three bac-

terial species (E. coli, P. aeruginosa and S. aureus) was investigated. Fibre

diameter was controlled and tuned by adjusting the electrospinning param-

eters, including solution concentration and conductivity.

To find the best approach to mimic the wound environment, two methods

of bacterial culturing (solution and agar cultures) were performed. The at-

tachment and growth of bacteria in and on the meshes was assessed using a

combination of scanning electron microscopy (SEM) and confocal laser scan-

ning microscopy after cell viability staining.

The same approach was used to investigate the influence of fibre surface

chemistry on the capacity of bacteria to transfer and attach on the elec-

trospun fibres. Among various methods for surface modification, plasma

polymerisation was chosen as it generates polymeric films with controllable

thickness and chemistry, conformable to the substrate surface features [21].

The polymer monomers (acrylic acid (ppAAc); 1,7-octadiene (ppOct); and

allylamine (ppAAm)) were chosen to investigate bacterial response to differ-

ent chemical functionalities with various degrees of wettability and surface

charge on the fibre surface. In addition, 1,8-cineole (ppCo), which is a com-

ponent of tea tree oils, was also included in the study as a material with

potential antibacterial activity [22].

To simulate the microbiological environment that wound dressings have to

face once implanted on patients, four in vitro tissue engineered skin models

of wounds at different depths and severity were developed.

Three models of non-infected wounds were used to investigate the mecha-

nisms of transfer of fibroblasts and keratinocytes onto the plasma modified

electrospun materials and to design a surface chemistry capable of preventing

cell ingrowth. In fact, for wound healing applications, the study of the mech-

4 / 223

Introduction

anisms of transfer and ingrowth of cells onto and within wound dressings is

essential for preventing wound reopening upon dressing removal.

In addition, to investigate the complex interactions occurring between the

electrospun fibres and the co-cultures of skin cells and bacteria, P.aeruginosa

cells were co-cultured with fibroblasts and keratinocytes in skin grafts to cre-

ate the model of a infected wounds.

5 / 223

Introduction

6 / 223

Chapter 1

Literature review

Contents1.1 The structure and function of human skin . . . 8

1.1.1 Physiological wound healing . . . . . . . . . . . . . 9

1.1.2 Chronic wounds . . . . . . . . . . . . . . . . . . . 11

1.2 Wound dressings . . . . . . . . . . . . . . . . . . . 14

1.3 Nanofibrous meshes . . . . . . . . . . . . . . . . . 22

1.3.1 The electrospinning techniques . . . . . . . . . . . 23

1.3.2 Control over the morphology of electrospun fibres . 25

1.4 Electrospun meshes as wound dressings . . . . . 29

1.5 Controlling biological interaction with electro-spun meshes . . . . . . . . . . . . . . . . . . . . . 38

1.5.1 How Do Bacteria Respond to Nanofibrous Meshes? 38

1.5.2 Role of fibre size and surface chemistry . . . . . . 40

1.6 Surface modification strategies . . . . . . . . . . 41

1.7 Biological responses to plasma polymerised sur-faces . . . . . . . . . . . . . . . . . . . . . . . . . . 45

1.7.1 Bacterial interactions with plasma polymerised sur-faces . . . . . . . . . . . . . . . . . . . . . . . . . . 45

1.7.2 Skin cell interactions with plasma polymerised sur-faces . . . . . . . . . . . . . . . . . . . . . . . . . . 46

1.8 In vitro Wound Models . . . . . . . . . . . . . . . 48

7

1. Literature review

1.9 Aims & objectives . . . . . . . . . . . . . . . . . . 51

1.1 The structure and function of human skin

The skin is the largest organ of the body and it is composed of different types

of tissue (connective, nervous, muscular, epidermal) [23, 24]. These tissues

constitute a multifunctional organ responsible for providing sensation, ther-

moregulation, biochemical, metabolic and immune functions and physical

protection [25, 26].

Skin is composed of two primary layers (Figure 1.1): the epidermis, which

acts as a barrier to infection; and the dermis, which serves as a location for

the appendages of skin.

Figure 1.1: Schematic representation of the layered structure of the human skin.

The dermis is composed primarily of collagen I, with dermal inclusions of hair

shafts, blood vessels, and sweat glands; its thickness varies depending on the

site in the body. The dermis is structurally divided into a superficial area

adjacent to the epidermis, called the papillary region, and a deep thicker area

known as the reticular region. The epidermal barrier layer is relatively thin

8 / 223


(0.1-0.2 mm in depth) and securely attached to the underlying dermis by a

specialized basement membrane zone. This consists of a fibrous, non-cellular

region of tissue formed by different types of collagen fibre, which attach cells

securely to the underlying dermis. Human skin comprises several different

cell types. Keratinocytes are the most common cell type in the epidermis,

while melanocytes are found in the lower layer of the epidermis and provide

skin colour. Fibroblasts form the lower dermal layer and provide strength

and resilience. Keratinocyte cells progressively differentiate from the cells in

the basal layer, which is located on the basement membrane, forming daugh-

ter keratinocytes, which are pushed upwards. These stratify, lose their nuclei

and eventually become an integrated sheet of keratin, which is later shed.

The upper keratinised epidermal layers (stratum corneum) provide the bar-

rier layer, which resists bacterial entry and prevents fluid and electrolyte loss

[23, 27].

The dermis is separated from the epidermal layers by the basement mem-

brane, thin, formed by various proteins, including collagen IV and proteo-

glycans.

1.1.1 Physiological wound healing

Human skin has inherent properties for promoting wound healing and pre-

venting infections of the wound bed, such as low moisture content, acidic pH,

high salt and lipid content and the presence of over 20 antimicrobial peptides.

In addition, the skin is colonized by different types of bacteria which form a

protective barrier against the adhesion and proliferation of other pathogens.

However the beneficial bacterial barrier protecting the skin surface is also

considered a potential source of infection when a disruption of the skins nor-

mal microbiological balance occurs [23, 28, 29].

9 / 223


A wound causes the loss of skin integrity and consequently the exposure of

subcutaneous tissues that can provide a moist, warm and nutritious environ-

ment particularly favourable for microbial colonisation and proliferation.

The physiological healing process of a wound consists in a cascade of se-

quential events that are perfectly coordinated and can be divided into four

successive phases: haemostasis, inflammation, proliferation and repair, re-

modelling [5]. The mechanisms involved in these four steps are summarised

in Table 1.2 (page 13).

During the last two phases of the healing process, the extracellular matrix

(ECM) plays an important role because it provides a frame which supports

and encourages epithelial cell proliferation [6]. The ECM is the non-cellular

component present within all tissues in the human body. It is a dynamic

and hierachically organised structure mainly composed of water, polysaccha-

rides and fibrous proteins (such as collagen, elastin, laminin, fibronectin, and

elastin). The fibrils forming the ECM have diameters ranging from 50 to 500

nm and form an interconnected fibrous network displaying specific ligands

that can bind to cell membrane receptors such as integrins [6, 30]. The ECM

acts as a scaffold by physically supporting cells and providing conditions for

cell adhesion and growth [30]. For this reason, one of the main goals for

effective wound care lies in reproducing the natural ECM-like environment

that is able to enhance and drive the healing process.

If the wound heals within a predictable time frame (8-12 weeks) and all the

described phases occur sequentially, the injury is classified as acute wound.

Chronic wounds fail to heal through these natural physiological processes.

Chronic wounds are classified in a number of ways: those which have not

healed after a fixed period of time (anywhere between four and six weeks to

up to three months); and those that do not show a 20-40% reduction in area

10 / 223


after two to four weeks of treatment. The most prevalent chronic wounds

are various forms of leg and foot ulcer. In most patients, the origins of de-

layed healing include dysfunction in the diabetic fibroblasts, immunological

defects due to genetic defects or cancer, malnutrition, obesity, drug abuse,

alcoholism, and smoking [31].

1.1.2 Chronic wounds

Several differences in the molecular environments of chronic and acute wounds

have been shown to be involved in the pathophysiology of chronic wounds. In

particular, chronic wounds exhibit higher protease activity, reduced growth

factor activity, and elevated levels of pro-inflammatory cytokines, if com-

pared to acute wounds [32]. Mast et al. provided a detailed description

of the pathophysiology underlying impaired healing in chronic wounds [32].

Although different wound types have different origins or causes, all chronic

wounds seem to be characterized by one or more persistent inflammatory

stimuli (repeated trauma, ischemia, or low-grade bacterial contamination),

which impair the physiological progression toward healing. When the skin

barrier is disrupted and bacterial colonisation occurs, endotoxins from bacte-

ria, platelet products, and fragments of extracellular matrix (ECM) attract

neutrophils and macrophages to the wound. These inflammatory cells are

responsible for the secretion of inflammatory cytokines, which increase the

production of metalloproteinases (MMPs) while reducing the production of

tissue inhibitors of metalloproteinase (TIMPs). The uncontrolled activity

of MMPs degrade the ECM, reducing cell migration and new connective

tissue deposition; moreover MMPs degrade growth factors, which are essen-

tial mediators within the cascade of mechanisms constituting the healing

process. Chronic wounds often fail to heal because tissue inflammation is

11 / 223


continuously stimulated and never overcome, and consequently the repair

stage of the healing process is impaired [32]. Chronic wounds are highly ex-

posed to the risk of bacterial infection because the longer the wound remains

opened and unhealed, the more likely it will be colonised by microorganisms

coming from different sources (external environment, surrounding skin, and

endogenous sources) [33]. Moreover, the devitalized tissue often found in

non-healing wounds facilitates the colonisation and proliferation of a wide

range of pathogens. Chronic wounds are contaminated by a polymicrobial

population of aerobic and anaerobic bacteria. Common aerobic or facultative

pathogens are Staphylococcus aureus, Methicillin-resistant Staphylococcus au-

reus (MRSA), Pseudomonas aeruginosa and Streptococci. Anaerobic bacte-

ria (Bacteroides, Prevotella, Porphyromonas, Peptostreptococcus) constitute

on average 38% of the total number of microorganisms found in chronic

wounds [28]. Their proliferation is encouraged by the low tissue oxygen level

often observed in chronic wounds. Due to their nature, anaerobes are hard

to be recovered and isolated form contaminated wounds with the traditional

clinical methods, thus further increasing the risk of infection [34].

12 / 223

1. Literature reviewP

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13 / 223


Wound infection develops through a process that results in the forma-

tion of a biofilm in the wound bed. Bacterial biofilms consist of a complex

microenvironment formed by single or mixed species of bacteria attached

to each other and encased in an extracellular polymeric matrix that bac-

teria themselves produce. Through the biofilm, bacteria can develop high

resistance against the immune system and antimicrobial agents, thus leading

to a quick proliferation [35, 36]. The biofilm protects microorganisms from

outer perturbations, allowing for microbial communication, enhanced viru-

lence and breakdown of nutrients. Studies have shown that the majority of

chronic wounds (60%) have a biofilm presence, compared with only 6% of

acute wounds [28].

1.2 Wound dressings

The ideal wound dressing accelerates the healing process, prevents infection

and restores the structure and function of the skin. Historically the first

documentation of wound care can be found in the ancient Sumerians who

used to apply poultices of mud, milk and plants to wounds. The Egyptians

prepared plasters of honey, plant fibres and animal fats as bandages for the

wounds. The most important advances in the field came with the develop-

ment of microbiology and cellular pathology during the 19th century [37].

One of the main contributions was the discovery in the 1960s that keeping

a wound moist accelerates the healing process. This became a key parame-

ter in the design and development of wound dressings [38]. However wound

dressings should satisfy other essential requirements for encouraging healing,

including: 1) absorbing excessive exudates from the wound bed, 2) provid-

ing thermal insulation, protecting the wound bed from mechanical trauma

14 / 223


and bacterial infiltration, 3) allowing gaseous and fluid exchanges, 4) being

removable without trauma, and 5) being nontoxic and nonallergenic [37].

Currently available wound dressings can be divided into four main cate-

gories according to the provided treatment: passive, interactive, advanced

and bioactive wound dressings (Table 1.3). Passive wound dressings provide

protection of the wound bed from mechanical trauma and bacterial infiltra-

tion. They are dry and do not control moisture levels in the wound, thus

they can adhere to wound bed causing pain and mechanical trauma when re-

moved [37]. Interactive dressings are fabricated with polymeric films and/or

foams, which are transparent and permeable to water vapor and oxygen;

they provide an effective barrier against permeation of bacteria or other mi-

croorganisms from the external environment. Advanced dressings such as

hydrocolloids and alginates are capable of providing or maintaining a moist

environment around the wound, thus facilitating the healing process [39].

The fourth category of bioactive dressings include those incorporating drug

delivery systems, skin substitutes and biological dressings which play an ac-

tive role in the healing process, by activating or driving cellular responses

[40]. Bioactive dressings constitute an important step forward towards the

development of effective systems capable of healing chronic wounds. How-

ever, research is still very intensive since these systems are only suitable for

specific types of wounds; moreover, costs and fabrication techniques can be

excessive, and a better control over drug release profiles and rates is an impor-

tant parameter that to date has not been optimized. A detailed description

of the wound dressings belonging to the described categories is provided in

Table 1.3.

A multifunctional device, able to treat different types of chronic wounds

while minimizing the risk of infection and wound recurrence is currently not

15 / 223


available to patients. Research in this field currently focuses on the develop-

ment of dressings able to combine three essential properties: 1) controlling

the physiological mechanisms on which the healing process is based; 2) moni-

toring markers of the healing and infection processes, including temperature,

pH and presence of bacteria; and 3) controlled release of drugs in response

to wound infection. A wound dressing capable of delivering all three of these

requirements would both stimulate the healing process while preventing in-

fections.

16 / 223

1. Literature reviewD

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diffi

cult

anat

omic

alsi

tes

(ove

rjo

ints

)

Con

tinued

onnextpage

17 / 223


Tab

le1.

3–Con

tinued

from

previouspage

Dre

ssin

gca

tego

ryP

roduct

Des

crip

tion

Applica

tion

s

Inte

ract

ive

Sem

i-p

erm

eable

foam

s

Mad

eof

pol

yure

than

eor

silico

ne

foam

.V

apor

and

oxyge

nex

chan

gean

dth

erm

alin

sula

tion

pro

vid

ed;

hig

hly

abso

rben

t,cu

shio

nin

gan

dpro

tect

ive.

Gen

eral

lynon

adhes

ive,

thus

requir

ing

seco

ndar

ydre

ssin

gs

Fla

t,sh

allo

w,

moder

ate

tohea

vily

exudin

gw

ounds;

not

for

ligh

tex

udin

gw

ounds

Am

orphou

shydro

gels

Am

orphou

sge

lor

elas

tic,

solid

shee

tor

film

.M

oist

ure

ism

ainta

ined

,va

por

and

oxyge

nex

chan

geal

low

ed;

wou

nd

deb

ridem

ent

pro

mot

ed.

Flu

idac

cum

ula

tion

wit

hin

the

dre

ssin

gca

nca

use

skin

mac

erat

ion

and

bac

teri

alpro

life

rati

on

Dry

,sl

ough

ing

ornec

roti

cw

ounds.

Not

for

moder

ate

tohea

vily

exudin

gw

ounds

Hydro

-co

lloi

ds

Mad

efr

omco

lloi

dal

mat

eria

ls,

com

bin

edw

ith

elas

tom

ers

orad

hes

ive

mat

eria

ls.

Thin

film

san

dsh

eets

oras

com

pos

ite

dre

ssin

gs.

Moi

sture

mai

nta

ined

orpro

vid

ed;

gas

and

fluid

sex

chan

geal

low

ed;

pH

ofw

ound

bed

reduce

dfo

rlim

itin

gbac

teri

algr

owth

Lig

ht

tom

oder

ate

exudin

gw

ounds.

Not

for

infe

cted

,nec

roti

cor

hea

vily

exudin

gw

ounds

Con

tinued

onnextpage

18 / 223

1. Literature reviewT

able

1.3

–Con

tinued

from

previouspage

Dre

ssin

gca

tego

ryP

roduct

Des

crip

tion

Applica

tion

s

Advance

d

Alg

inat

esM

ade

from

the

calc

ium

and

sodiu

msa

lts

ofal

ginic

acid

.F

reez

e-dri

edp

orou

ssh

eets

(foa

ms)

orflex

ible

fibre

s.H

ighly

abso

rben

t.O

pti

mal

moi

sture

leve

lan

dte

mp

erat

ure

mai

nta

ined

.C

lott

ing

mec

han

ism

sen

coura

ged

Moder

ate

tohea

vily

exudin

gw

ounds.

Not

for

dry

ornec

roti

cw

ounds

Hydro

fibre

sM

ade

from

sodiu

mca

rbox

ym

ethyl

cellulo

sefibre

s.W

ound

exudat

esab

sorb

edan

dm

oist

ure

pro

vid

ed.

pH

ofw

ound

bed

reduce

dfo

rlim

itin

gbac

teri

algr

owth

Infe

cted

,m

ediu

mto

hea

vily

exudin

gw

ounds.

Not

for

dry

orligh

tex

udin

gw

ounds

Dex

tra-

nom

ers

Hydro

philic

pol

ysa

cchar

ide

gran

ule

sav

aila

ble

inp

owder

orpas

tefo

rm.

Hig

hly

abso

rben

t;op

tim

alm

oist

ure

leve

lpro

vid

ed

Med

ium

tohea

vily

exudin

g,in

fect

edw

ounds

Con

tinued

onnextpage

19 / 223


Tab

le1.

3–Con

tinued

from

previouspage

Dre

ssin

gca

tego

ryP

roduct

Des

crip

tion

Applica

tion

s

Dru

gdel

iver

yC

onti

nuou

sre

leas

ein

the

wou

nd

ofan

tim

icro

bia

lag

ents

(hon

ey,

iodin

e,si

lver

,p

olyhex

amet

hyl

big

uan

ide,

chlo

rhex

idin

egl

uco

nat

e)lo

aded

into

inte

ract

ive

orbio

acti

vedre

ssin

gs.

Age

nts

rele

ased

also

ifw

ound

isnot

infe

cted

.R

elea

sepro

file

san

dra

tes

tob

eop

tim

ized

Infe

cted

orhig

hly

conta

min

ated

wou

nds

Bio

act

ive

Bio

logi

cal

dre

ssin

gsM

ade

from

nat

ura

lor

bio

logi

cal

syst

ems

(mic

roor

ganis

ms,

pla

nts

,an

imal

s)or

chem

ical

lysy

nth

etiz

edfr

ombio

logi

cal

star

ting

mat

eria

ls(s

tarc

h,

nat

ura

lfa

ts,

oils

,su

gars

).C

olla

gen,

Gel

atin

,C

hit

osan

,H

yalu

ronic

acid

bas

eddre

ssin

gsen

coura

gefibro

bla

stac

tivit

yan

den

dot

hel

ial

cells

mig

rati

on.

Imm

unog

enic

resp

onse

can

be

induce

d

Cle

an,

non

-infe

cted

and

non

-nec

roti

cw

ounds

Con

tinued

onnextpage

20 / 223

1. Literature reviewT

able

1.3

–Con

tinued

from

previouspage

Dre

ssin

gca

tego

ryP

roduct

Des

crip

tion

Applica

tion

s

Skin

subst

itute

sT

issu

ecu

lture

(allog

enic

orau

tolo

gous

sect

ion

ofsk

inhar

vest

edan

dcu

lture

din

lab

orat

ory

tofo

rmsh

eets

ofce

lls

tob

eim

pla

nte

d)

orT

issu

een

ginee

ring

(nat

ura

lor

synth

etic

pol

ym

ers

are

use

das

mat

rice

sto

cult

ure

cells)

Sev

ere

burn

sor

chro

nic

wou

nds

wit

hlo

ssof

imp

orta

nt

por

tion

ofth

esk

in.

Cle

an,

non

-infe

cted

and

non

-nec

roti

cw

ounds

Tab

le1.3

:C

lass

ifica

tion

of

com

mer

ciall

yava

ilabl

ew

ou

nd

dre

ssin

g[1

0,

14,

27,

37,

40,

41].

21 / 223


1.3 Nanofibrous meshes

One of the principal research drivers in the field of wound care development

focuses on the manufacture of wound dressings in the form of nanofibrous

meshes [8]. These structures are made of non-woven, ultra-fine polymeric fi-

bres with diameters ranging from several micrometers down to a few nanome-

ters. Nanofibrous meshes have several intrinsic properties, which make them

particularly interesting for wound healing applications. First of all, the ideal

wound dressing should be able to mimic the structure and the functional

biology of the extracellular matrix (ECM) in order to encourage the prolif-

eration of epithelial cells and the formation of new tissue [7]. During the

healing process the ECM acts as a scaffold for physically supporting cells

and providing conditions for cell attachment, proliferation, migration and

differentiation [30].

Nanofibrous meshes offer a good starting point towards the development of

a synthetic scaffold able to reproduce the structure of the natural ECM.

In fact, due to their nanometer diameter and random alignment within the

mesh, fibres tend to imitate the fibrous architecture of the natural ECM. In

addition, nanofibrous meshes have been shown to promote the hemostasis

of injured tissues thanks to the presence of small interstices and the high

surface area of the fibres. The high surface area is also essential for fluid

absorption, enhanced dermal drug and antimicrobial delivery and it provides

the opportunity to modify the surface of the fibres with specific chemical

functionalities [8]. Nanofibrous meshes show high interconnected porosity

(60-90%), allowing cell respiration and high gas permeation and prevention

of wound desiccation and dehydration [6].

To prevent the infiltration of microorganisms from the external environment

and discourage cell/tissue ingrowth, the ideal fibrous mesh for wound healing

22 / 223


Properties Advantages

Fibre diameter 50-500 nm Mimic of the physical structure of thenatural ECM

High surface area to volumeratio

Hemostasis promotion; surfacefunctionalisation

High porosity (60-90%) Cell respiration; gas permeation; wounddehydration prevention

Interconnectednano-porosity

Prevention from microbial infiltration andcell ingrowth

Mechanical strength Similar to natural skin

Table 1.4: Ideal properties of nanofibrous meshes for wound healing applications[1, 6–8].

should have pores with nanometer dimensions [8]. A list of the key properties

that an effective wound dressing should possess is provided in Table 1.4.

Various techniques are available for the fabrication of nanofibrous meshes

and they have been reviewed in detail by Yanzhong et al. [8]. Currently,

electrospinning is the preferred technique of the majority of researchers for

the range of advantages outlined in the following sections.

1.3.1 The electrospinning techniques

Compared with other polymeric materials fabrication techniques (i.e. phase

separation or self-assembly), electrospinning provides a simple and cost-

effective way to produce fibrous meshes with an inter-connected pore struc-

ture and fibre diameters in the sub-micron range. It allows the fabrication

of fibres with high surface area due to their diameters being scalable down

to a few nanometres. Electrospun meshes can be surface functionalized to

23 / 223


tune the physical and chemical properties of the fibre surface while the fibre

structure, morphology and spatial distribution can be controlled to achieve

specific mechanical properties. In addition, electrospinning allows for the

combination of different synthetic and natural polymers to be used to make

nanofibres. The possibility of large scale production combined with simplicity

and versatility makes the electrospinning process very attractive for a broad

variety of applications which have been reviewed by Jian et al. [42] and by

Huang et al. [1] (Figure 1.2). The use of electrospun 2- or 3-dimensional

scaffolds for biomedical applications including drug delivery, vascular, bone

and heart tissue engineering has been reviewed by various authors [9, 43, 44].

A typical electrospinning setup (Figure 1.3) consists of a syringe and capillary

needle through which a polymer solution or melt is passed (the spinneret);

a high voltage power supply and a grounded collector [45]. Bhardwaj et al.

provided a detailed description of the electrospinning technique as well as

the parameters affecting the process [45]. Briefly, a high voltage up to 30 kV

is applied at the tip of the capillary needle, where a pendent droplet of the

polymer solution or melt gets electrified, inducing charge accumulation on

the droplet surface. The charge causes the deformation of the droplet into

a cone, called the Taylor’s cone, from which a fine charged polymer jet is

ejected. The jet moves towards the collector while the solvent evaporates,

thus obtaining ultrafine dry fibres that can be collected on the grounded

electrode in form of a fibrous mesh. The basic configuration shown in Figure

1.3 is used for the fabrication of non-woven meshes composed of randomly

aligned fibres [45]. More complex setups are available and have been re-

viewed by Sahay et al. [46] and by Migliaresi et al. [47]. Various types of

collector, including rotating mandrel, rotating wheel, parallel electrodes and

rings and patterned electrodes, enable fibre alignment along a specific direc-

24 / 223


tion with uniform fibre distributions within the mesh (Figure 1.4). Two or

more extruding capillaries can be used simultaneously for fabricating fibres

in different polymers within the same mesh [46, 47]. Multi-needle, needless,

coaxial electrospinning are advanced setups that create the opportunity to

combine materials and compounds that normally do not tend to mix ho-

mogeneously, but when added together in a fibre structure add significant

functionality to the final material [47].

1.3.2 Control over the morphology of electrospun

fibres

The control over the diameter and surface morphology of fibres fabricated

through the electrospinning process can be challenging given the range of

process and solution parameters involved. Since the fibres are formed by the

evaporation or solidification of polymer fluid jets, fibre diameters will depend

primarily on the jet sizes and on the polymer content in the jets. Polymer

concentration is one of the principal parameters affecting the diameter of

the resultant fibres. The higher the polymer concentration the larger the

resulting nanofibre diameters will be [1]. However, to fabricate fibres in the

nanoscale, reducing the polymer concentration might not be the only ap-

proach. In fact, if the concentration is too low the solution will form droplets

before reaching the collector; moreover at low concentrations defects in form

of beads and polymer agglomerates tend to form along the fibres resulting in

non-uniform discontinuous meshes [1].

The conductivity of the polymer solution can be tuned to achieve a better

control over fibre morphological properties. Increasing the solution conduc-

tivity results in a greater tensile force acting on the polymer droplet when

the electric field is applied, in turn resulting in a reduction of fibre size [48].

25 / 223


Figure 1.2: (a) Application fields of electrospun nanofibres targeted by USpatents and (b) potential application of electrospun polymeric nanofibres [1].

26 / 223


Figure 1.3: Schematic representation of electrospinning basic set up. (a) Imageof Taylor cone forming at the spinneret during the electrospinning process [2]; (b)Image of polymeric filament forming from Taylor cone and moving toward the

collector [3].

27 / 223


Figure 1.4: Schematic representation of electrospinning collectors: (a) planarcollector for non-woven meshes; (b) square frame for unidirectional oriented

fibres; (c) cylindrical collector for tubular oriented fibres; (d) Non-woven fibresfabricated using collector (a); (e) aligned fibres that can be fabricated using

collectors (b) or (c).

Solution conductivity can be tuned by choosing an appropriate solvent or by

adding surfactants to the polymer solutions. However, this approach might

result in the unprocessability of the solution if the conductivity is too high

[45].

Besides solution parameters, several process parameters can also affect fibre

diameter, including voltage, solution feed rate, spinning distance, tempera-

ture, and humidity. Voltage is an important parameter in electrospinning

because the charges on the polymer molecules that form the fibre jet orig-

inate from the applied voltage. At high voltage, the solution jet will be

drawn at a faster rate and will experience a greater acceleration. Moreover,

the greater acceleration causes the solvent in the solution to evaporate faster,

reducing the fibre jet volume. These factors are in favor of reducing fibre di-

ameter. However, increased solution jet acceleration also reduces the flight

time, which means that the jet will reach the collector quicker. A reduced

28 / 223


flight time reduces the time available for the fibres to stretch, leading to a

greater fibre diameter. This ultimately implies that as the voltage increases,

fibre diameter will decrease, but when the voltage increases past an optimum,

fibre diameter may increase [49].

The variety of studies and mathematical models on the electrospinning pro-

cess available in the literature highlight the complexity of the mechanisms

involved in the formation of the fibres and in the control of their morphologi-

cal features. Control over fibre size can be achieved only through a systematic

empirical approach that will result in the best combination of process and

solution parameters for the chosen polymer/solvent system.

1.4 Electrospun meshes as wound dressings

Electrospun polymer nanofibres for wound healing applications can be broadly

classified as synthetically or naturally derived. The most frequently selected

polymers have been reviewed by Zahedi et al. [10] and the key materials are

summarized in Table 1.6.

Traditionally, electrospun meshes for biomedical applications have been fab-

ricated from single solutions of polymers. Considering the advantages and

disadvantages of both synthetic and naturally derived materials (Table 1.6),

mixtures of different polymers are becoming widespread. The so-called ”poly-

blended” nanofibres are obtained by electrospinning premixed or multiple

polymer solutions. Synthetic polymers ensure easy processability and good

mechanical properties of the resulting mesh, while natural polymers increase

the capability of the fibres to actively interact with biomolecules involved

in the healing process [12]. Electrospun meshes fabricated during the past

decade for wound healing applications can be classified according to the same

29 / 223


system previously adopted for existing commercial dressings: passive; inter-

active; advanced, and bioactive. This classification reflects the evolution of

electrospun wound dressings in terms of selected materials and technologies

for both fabrication and functionalisation of the fibres.

30 / 223


Ad

vanta

ges

Dis

adva

nta

ges

Pol

ym

ers

Synth

etic

sE

asi

lyta

ilor

edto

pro

vid

ea

wid

era

nge

of

funct

ional

pro

per

ties

.S

tron

g,ch

eap

and

reli

able

,ea

sily

pro

cess

able

,su

rfac

em

od

ifiab

lean

dst

eril

isab

le

Som

em

ater

ials

can

rele

ase

toxic

deg

rad

atio

np

rod

uct

s.S

mal

lp

arti

cles

can

be

rele

ased

du

rin

gd

egra

dat

ion

cau

sin

gin

flam

mat

ory

resp

onse

.S

yst

emic

orlo

cal

reac

tion

sca

nb

ein

du

ced

.L

oss

ofm

ech

anic

alp

rop

erti

esca

nocc

ur

very

earl

yd

uri

ng

deg

rad

atio

n

Pol

yla

ctic

acid

(PL

A);

Pol

ygl

yco

lic

acid

(PG

A);

Pol

yla

ctic

-co-

glyco

lic

acid

(PL

GA

);P

olyca

pro

lact

one

(PC

L);

Poly

vin

yl

alco

hol

(PV

A);

Pol

yu

reth

ane

(PU

);P

olyst

yre

ne

(PS

)

Nat

ura

lsM

any

mat

eria

lsp

rese

nt

nat

ive

bio

mol

ecu

lar

sign

als

ass

oci

ate

dw

ith

cell

bin

din

g/p

roli

fera

tion

/mig

rati

onan

dim

mun

ere

spon

ses

Pro

cess

ing

can

ind

uce

den

atu

rati

on.

Har

vest

ing

and

pro

cess

ing

can

be

com

ple

x.

Sou

rces

ofb

iop

olym

ers,

pu

rity

and

mol

ecu

lar

wei

ght

dis

trib

uti

onca

nin

flu

ence

pro

cess

pro

per

ties

and

resu

ltin

gar

chit

ectu

reof

mes

hes

.R

isk

ofd

isea

setr

ansm

issi

onan

dp

ossi

ble

anti

gen

icit

y.

Col

lage

n;

Gel

atin

;C

hit

osan

;C

hit

in;

Fib

rin

ogen

;H

yal

uro

nic

acid

;C

ellu

lose

Tab

le1.6

:S

elec

tion

of

the

most

freq

uen

tly

use

dsy

nth

etic

an

dn

atu

ral

poly

mer

sfo

rfa

bric

ati

on

of

nan

ofi

brou

sw

ou

nd

dre

ssin

gby

the

elec

trosp

inn

ing

tech

niq

ue

an

dth

eir

adva

nta

ges

an

ddis

adva

nta

ges

[9–12].

31 / 223


Passive Electrospun Meshes Meshes that provide the physical (i.e. wa-

ter and gas permeability) and morphological (i.e. adequate porosity and

nanometer scale) properties of wound dressings are classified as passive.

These systems are able to maintain suitable levels of moisture in the wound

bed and protect tissues from mechanical trauma. Passive electrospun meshes

are fabricated with both natural and synthetic polymers and are designed for

those wounds that require ideal moisture levels and protection from external

pathogens to achieve complete healing. Khil et al. produced electrospun

poly(urethane) membranes and both morphological characterization and in

vivo experiments indicated that the membranes could be employed as wound

dressings [50]. Phachamud et al. optimized the electrospinning parameters

for the fabrication of poly(vinyl alcohol) (PVA) fibres. Due to the homo-

geneous fibre distribution, high swelling and weight loss of the meshes, the

authors suggested their potential use for wound healing applications [51]. Up-

pal et al. fabricated nanofibrous meshes by the electrospinning of hyaluronic

acid (HA). In vivo studies showed that meshes offered the best treatment of

full-thickness wounds when compared with other four commercial dressings

(adhesive bandage, a sterilized HA film, gauze with Vaseline and an antibi-

otic dressing) [52].

This initial phase of research into fabrication of nanofibrous meshes as wound

dressings focused on the optimization of the electrospinning process of var-

ious natural and synthetic polymers for achieving suitable morphological,

physico-chemical and mechanical properties. The more recent developments

aim to create active devices able to drive the healing process and prevent or

treat infection.

32 / 223


Interactive Electrospun Meshes Electrospun meshes that combine the

necessary morphological and physical requirements for wound healing with

the value-added capability to address optimal cell responses and limit bacte-

rial proliferation in the wound bed are classified as interactive.

The main strategy which has been used to develop interactive systems con-

sists of the combination of synthetic polymers and biopolymers, which exhibit

antibacterial properties and affinity towards ECM components. Multicompo-

nent systems more closely mimic the ECM. In fact, the ECM is composed of

an interconnected structure of proteins (i.e. collagens, laminin, fibronectin,

elastin), proteoglycans (i.e. heparan sulfate, chondroitin sulfate, keratan sul-

fate), and glycoaminoglycans (i.e. hyaluronic acid) that can be included in

the polymer formulation to be electrospun.

Two main approaches for combining natural and synthetic polymers within

the same electrospun mesh can be identified: different polymers can be

blended to form a single solution to be electrospun; synthetic polymers are

electrospun and subsequently the mesh is coated with the selected natural

polymer. The second strategy aims to exploit the higher mechanical prop-

erties and easier spinnability of synthetic polymers for the fabrication of the

mesh.

A broad variety of reports based on the combination of natural and synthetic

polymers is available in literature. Yuan et al. fabricated meshes for wound

healing by electrospinning a blend of modified keratin and poly(hydroxybutylate-

co-hydroxyvalerate) (PHBV). Keratin is a family of fibrous proteins that are

present in a wide range of biological tissues, performing a structural role in

skin and hair. From wound healing and histological tests the authors showed

that the composite meshes accelerated wound recovery [53]. Kim et al. fab-

ricated electrospun meshes by blending polyurethane (PU) and gelatin and

33 / 223


showed the potential application in wound healing. Gelatin is a natural poly-

mer derived from collagen often chosen for biomedical applications since it

is biodegradable, non-toxic and easily available at low cost [54]. Chen et al.

fabricated composite nanofibrous meshes by blending type I collagen, chi-

tosan and poly(ethylene oxide) (PEO), that showed better performance in

wound healing rates in rat models than traditional dressings [55]. Chitosan

is frequently chosen for fabricating composite electrospun meshes because

it can function as a proliferation promoter, antibacterial agent and wound

healing accelerator [55–59]. Spasova et al. [60] coated electrospun poly(L-

lactide) (PLLA) and bicomponent PLLA/poly(ethylene glycol) meshes with

chitosan. Hemostatic and antibacterial activity against S.aureus of the coat-

ing was demonstrated, thus presenting the meshes as possible candidates for

wound dressings. Ignatova et al. [61] overviewed the most recent studies on

electrospun chitosan-based meshes for biomedical applications.

There is also significant focus on alternative plant-derived compounds, in-

cluding essential oils and honey [62]. Natural substances cannot generally

be electrospun into fibres unless they are blended with synthetic polymers

as they lack mechanical and structural stability upon hydration. Normally

pure solutions of natural materials are not electrospun because the process

would result in electrospray at low concentrations or complete occlusion of

the spinneret at higher viscosities [63]. Among natural compounds, honey

is a very attractive material due to its anti-inflammatory and antimicrobial

properties [64, 65]. Due to its low pH (3.5-4), honey is theorised to mod-

ify the alkaline environment characteristic of chronic wounds towards more

acidic conditions favorable for wound healing [66].

Although reducing the pH in the wound bed has been shown as a possible

strategy for controlling the bacterial load, very few reports exist trying to

34 / 223


fabricate nanofibre meshes capable of providing this capability. Opportu-

nities to develop interactive dressings able to provide healing conditions to

the wound by reducing the pH to acidic values exist and should be further

explored.

Although interactive electrospun meshes have been shown to encourage wound

healing, they are not yet available on the market. This is due to the difficul-

ties associated in electrospinning naturally derived polymers in a reproducible

manner and in large scale. The complexity associated with processing nat-

ural polymers due to issues with impurities and the possibility of inducing

immunogenic reactions upon implantation are currently limiting the trans-

ference of these meshes into devices that can be used in reality.

Advanced Interactive Electrospun Meshes In order to manufacture

interactive dressings capable of treating bacterial infection, many researchers

are currently developing drug loaded nanofibrous meshes. Meinel et al.

overviewed drug loaded electrospun nanofibres, providing the most frequently

selected drugs for wound healing applications, which include antibiotics, an-

tiseptics and antibacterials, and anesthetics [67].

The most common technique for loading compounds into the nanofibres is

known as coaxial electrospinning, which allows the compound to be retained

in the fluid environment after being loaded into the fibre. The resultant

nanofibres present a core/shell structure where the shell is normally made of

a synthetic polymer for structural integrity while the bioactive compounds

in their original liquid state or encapsulated in a second polymer remain in

the core of the fibre. A slightly different setup than the traditional elec-

trospinning is used at this purpose. Two syringes are used to transfer the

polymer solution and the compound to the spinneret, which is constructed

35 / 223


of a single capillary with an inner and an outer channel. The polymer solu-

tion normally feeds the outer channel while the compound, which can be a

liquid or a polymer composite, is extruded through the inner one [68]. This

process is used to load a variety of compounds into nanofibres that could

potentially lose their functionality unless they are in fluid or non-denaturing

environment and include drugs, growth factors and vitamins.

The possibility of loading antibiotics or antimicrobials into electrospun fibres

represents a great advantage in the development of systems able to treat in-

fections in the wound bed. However, these systems have not been translated

into an effective therapy for various reasons [67, 69]. Leung et al. highlighted

that, depending on the type of wound optimal drug release profiles and rates

of release are required [70]. Furthermore, drug release is always associated

with an initial burst effect, which can cause the local drug concentration to

be toxic towards tissue cells [9].

The ideal wound dressing should be a smart device adaptable to treat every

different kind of wound. It should be able to monitor the conditions of the

tissues in the wound and subsequently trigger the release of drugs with the

optimal delivery profile only when needed.

Some papers report on nanofibres functionalized with growth factors to stim-

ulate cell growth, and encourage and accelerate wound healing. Although the

efficacy of these meshes in terms of encouraging proliferation and differenti-

ation of fibroblasts and keratinocytes have been demonstrated [71, 72], the

high cost of fabrication as well as the difficulties associated with processing

are limiting the popularity of these systems. Moreover, as previously stated,

the ideal wound dressing should be able to promote skin cell migration and

proliferation within the wound bed, while preventing tissue ingrowth within

the fibrous structure, in order to avoid tissue damage after dressing removal.

36 / 223


The capability of electrospun meshes to prevent fibroblast and keratinocyte

ingrowth has not been demonstrated yet.

Bioactive Electrospun Meshes Bioactive electrospun meshes aim to be

multifunctional systems, combining a range of properties capable of treating

all aspects of the wound. Adequate mechanical and physico-chemical prop-

erties provide wound protection, the healing process is stimulated and the

bacterial load in the wound bed controlled.

In many studies, work is moving to include wound status monitoring as an

indicator of the progression of healing and/or the bacterial load. This can

be achieved by integrating a sensor within the electrospun mesh thus allow-

ing the real-time detection of specific parameters from the wound bed. The

sensor generates a visible output for the patient or the doctor providing con-

tinuous monitoring of the wound status.

Dargaville et al. reviewed the state-of-the art in the fabrication of sensors for

monitoring the healing process of wounds [73]. There are a range of poten-

tial markers and parameters associated with wound healing and infections

that can be detected, including pH and temperature. A number of groups

have developed pH sensitive dyes and immobilized them onto films and into

fibres [74–76]. A few reports exploit the capability of hydrogels to swell in

response to pH, temperature or analyte concentration for sensing the status

of the wound [77, 78]. Van der Werff et al. developed a bandage that changes

colour according to the temperature of underling tissues in order to monitor

the healing processes of wounds [79].

Although the literature suggests that pH and temperature sensors for wound

monitoring are possible, few studies have actually integrated these systems

into wound dressings or electrospun meshes for testing.

37 / 223


Biofouling represents a significant obstacle to the inclusion of sensors within

wound dressings. As with any sensor, the uncontrolled adsorption of biomolecules

(peptides, proteins and subsequent attachment of cells) will impeded analyte

detection and can cause the failure of the device [80]. Moreover, when de-

signing a sensor to be integrated into a wound dressing, an essential criterion

needs to be addressed: the sensor outputs should never be used in isolation.

Multiple signals and parameters should be simultaneously detected in the

wound environment and combined for determining the final output in terms

of current status of the wound [73]. Finally, cost-effectiveness of the fabri-

cation and engineered processes cant be ignored and given the low per-item

cost of many wound dressings, this is a critical parameter.

1.5 Controlling biological interaction with elec-

trospun meshes

1.5.1 How Do Bacteria Respond to Nanofibrous

Meshes?

There is significant data available from the tissue engineering literature on

how skin cells, in particular fibroblasts and keratinocytes interact with elec-

trospun meshes with detailed in vitro studies used to assess cell viability

and growth [45, 81, 82]. Sun et al. studied the influence of fibre diameter,

inter-fibre distance and fibre alignment on the behaviour of human dermal

fibroblasts. They identified minimum values of fibre diameter and inter-fibre

space necessary for cell adhesion and migration and for cell aggregate forma-

tion [82]. Fibre alignment was shown to affect cell behaviour by inducing cell

guidance [30, 82]. Nisbet et al. provided a detailed review of how different

38 / 223


cell types respond to electrospun nanofibres [83].

Although this knowledge is very useful for developing effective scaffolds capa-

ble of actively drive cell behaviour, the study of cell responses is not the only

aspect that must be considered in designing a dressing able to control and

address the healing process. In fact, during the healing process, nanofibrous

meshes are inevitably involved in dynamic interactions with the wound envi-

ronment, which includes coexistence with bacteria [84]. Since all wounds are

contaminated by bacteria, an understanding of how bacteria interact with

the electrospun meshes is essential to develop devices able to not only com-

municate with cells but also minimise the microbial load in the wound bed

and reduce the risk of infection.

Theoretical approaches, thermodynamic theories, and cell studies have pro-

vided important insights on the mechanisms that control bacterial adhesion

and on the role played by cell surface properties. It is currently recognized

that, apart from cell surface characteristics, bacterial attachment mechanisms

are also regulated and influenced by a wide range of substratum properties,

such as morphology, surface chemistry, and roughness [18]. The mechanisms

that bacteria use to adhere to flat surfaces with different chemistries and sub-

sequently develop into biofilms has been reviewed in some detail by Mitik-

Dineva et al. [85]. In other work, Mitik-Dineva et al. and Anselme et al.

studied cellular and bacterial interactions with nano-structured flat surfaces

and showed that the nanoscale topography of surfaces could be exploited to

limit bacterial proliferation [86, 87].

To the best of our knowledge, despite bacterial infection representing a ma-

jor challenge in chronic wound care, few studies focus on microbial adhesion

and growth and biofilm formation on the surface of electrospun fibres and

those that exist tend to focus on membrane fouling in water environments or

39 / 223


on bactericidal effects of drug loaded meshes [88]. The mechanisms of inter-

actions between individual bacteria and fibres with different morphological

and chemical properties need to be explored. The knowledge of these mecha-

nisms could allow to tune the morphological and surface chemistry properties

of electrospun fibres to control and block microbial growth, without the need

to incorporate drugs.

1.5.2 Role of fibre size and surface chemistry

As the interaction of mammalian cells with electrospun substrates have been

proven to strictly depend on the size of the fibres, the need exists to explore

the response of bacterial cells to meshes with different average fibre diameters

[82]. The work that most closely approaches this problem in the literature

was provided by Kargar et al., who investigated the state of adhesion of

P.aeruginosa bacteria to PS flat surfaces texturized with aligned PS fibres

with different diameters and spacing. The minimum value of bacterial ad-

hesion density was found to occur for fibres with diameter close to bacterial

diameter at a spacing less than bacterial diameter; the highest density was

measured when the spacing between fibres and fibre diameter were bigger

than bacterial size [89]. These results were obtained by studying the single-

cell level, thus bringing up the question about the influence of fibre diameter

on the capacity of bacteria to spread and colonise the meshes.

The literature provides a variety of studies on the response of bacteria to flat

substrates with different surface chemistries. Mitik-Dineva et al. reviewed

the general principles that apply to mechanisms of bacterial attachment onto

flat surfaces, highlighting that the surface characteristics of both the sub-

strate and the bacteria play a role in the attachment process [85].

Physico-chemical properties of the substrate including surface roughness,

40 / 223


wettability and charge density were correlated to the ability of the bacte-

ria to attach and proliferate [85]. Whitesides et al. investigated the adhesion

of bacteria adsorbed to arrays of self-assembled monolayers (SAMs) on gold

displaying different functional groups. Authors demonstrated that surface

chemistry, particularly specific functional groups on the material, influence

the attachment of the bacteria [90, 91]. Cunliffe et al. carried out a system-

atic investigation into the effect of surface chemistry on bacterial adhesion

involving the grafting of groups varying in hydrophilicity, hydrophobicity,

chain length, and chemical functionality onto glass substrates. Surface charge

and wettability were proven to affect the adhesion process depending on the

bacterial species tested [92].

The numerous studies available in the literature agree that the mechanisms

of microbial attachment onto flat substrates are controlled by the wettability,

surface charge and functional groups of the substrate chemistry. However,

a universal general trend cannot be identified as the obtained results are

simultaneously dependent on other factors, including bacterial species and

topography and morphology of the substrates.

The need now exists to translate these studies onto more complex multidi-

mentional substrata designed to interact with a variety of bacterial species.

The open question refers to the influence of the surface chemistry of electro-

spun fibres on bacterial behaviour to understand if changing the overall mesh

properties can specifically address bacterial attachment and proliferation.

1.6 Surface modification strategies

Yoo et al. reviewed the surface modification and functionalisation strategies

that are available for electrospun nanofibres for tissue engineering and drug

41 / 223


delivery applications [4]. Surface modification of electrospun fibres includes

plasma treatment, wet chemical method, surface graft polymerisation, and

co-electrospinning. For chronic wound applications most of these strategies

are used to generate reactive fibre surfaces onto which bioactive molecules,

including antibiotics, vitamins, and growth factors, can be entrapped or im-

mobilised for controlled drug delivery. In addition, fibre surface modification

has been carried out to immobilise a wide variety of natural polymers having

unique biological functions onto the nanofibrous surface of synthetic poly-

mers without compromising the bulk properties [4].

A description of the surface modification techniques for electrospun meshes

is provided in in Figure 1.5. Table 1.7 summaries the mechanisms involved

and the advantages and limitations of each technique.

In the next sections significant attention will be given to plasma polymeri-

sation as this technique was chosen for the surface modification of the elec-

trospun fibres. Plasma polymerisation was selected among the other surface

modification techniques because it is a dry single-step process that can po-

tentially be applied to any type of substrate without compromising the bulk

mechanical properties; the thickness of the plasma polymer films can be

tightly controlled from 5-10 nanometres up to micrometres. Plasma poly-

mers successfully adhere to a range of substrates and, due to the gaseous

nature of the monomers, films can coat complex structures and conform to

them [93].

42 / 223


Figure 1.5: Approaches for surface modification of electrospun fibres: (a)plasma polymerisation; (b) wet chemical method; (c) surface graft polymerisation;

and (d) co-electrospinning [4].

43 / 223


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44 / 223


1.7 Biological responses to plasma polymerised

surfaces

1.7.1 Bacterial interactions with plasma polymerised

surfaces

Plasma polymerisation and plasma treatment have been widely used for de-

veloping coatings to prevent the attachment of bacteria and the subsequent

biofilm formation onto a variety of device surfaces [95].

One approach consists in developing anti-fouling surfaces that can resist not

only protein adsorption and mammalian cell attachment but also bacterial

attachment. Fully attachment-resistant surfaces have been obtained by the

grafting of hydrogel polymer layers with poly(ethylene glycol) (PEG). Plasma

polymer coatings including n-heptylamine and allylamine were used as inter-

layers for the covalent grafting of fouling-resistant PEG hydrogel layers and

some resistance to bacterial colonisation was shown [96–98] .

Another medical use of plasma polymers consists in carrier matrices for

nanoparticles, metal ions, and drugs. Compared to other release matrices,

such as hydrogels or osmotic systems, plasma polymers have the key advan-

tage of ensuring excellent adhesion to the substrate surface without affecting

the bulk properties. In addition, the release profile of the embedded com-

pounds can be controlled via the plasma polymer crosslink density or via

the deposition of a second plasma polymer [99]. Silver nanoparticles or ions

have become one of the most popular antimicrobial agents incorporated and

released by plasma coatings, including polytetrafluoroethylene, allylamine,

and siloxane plasma polymers deposited onto various substrates [99–101].

Another common strategy to incorporate antibiotic molecules in plasma poly-

45 / 223


mer carriers consists in spreading nanocrystals of the drug onto the substrate

surface and consequently depositing the plasma polymer layer to ensure

that the nanocrystals are held in place and the release profile is controlled

[102, 103].

Antimicrobial compounds, such as quaternary ammonium compounds or an-

timicrobial peptides, have also been covalently grafted onto plasma polymers

(allyl alcohol, n-butyl methacrylate, and allylamine) for developing sur-

faces with longer-lasting antibacterial activity than is possible via release

approaches [95].

Clearly, most of the literature on plasma polymers for medical devices focuses

on the development of drug release surfaces. However, plasma polymerisa-

tion also offers the possibility of controlled surface chemistry modification

and can be used to generate surfaces with different functional groups, wetta-

bility and surface charges on multidimensional substrata. The mechanisms

of interactions occurring at the interface between plasma polymerised sur-

faces and mammalian cells have been widely explored, but surprisingly no

systematic study involving bacteria is available to the best of our knowledge.

1.7.2 Skin cell interactions with plasma polymerised

surfaces

In the tissue engineering literature there is an increasing interest towards the

surface modification through plasma polymerisation of porous 3-dimensional

scaffolds to control and address cell attachment and proliferation [104, 105].

Significant efforts were run to individuate the effect of different functional

groups introduced by plasma polymerisation (hydroxyl, carbonyl, carboxyl,

etc.) on cell adhesion and growth. Significant contribution to chronic wound

and burn management was provided by MacNeil et al. who developed plasma

46 / 223


polymerised surfaces to induce the attachment and culture of human ker-

atinocytes and their subsequent transfer to a wound bed [106]. The culture

of human keratinocytes was successfully achieved on acrylic acid plasma poly-

merised flat surfaces, with the number of cells attached comparable to the

performance of cells on collagen I, which is a preferred substrate for ker-

atinocyte culture. Different concentrations of carboxylic acid groups were

tested and cell attachment was enhanced on surfaces with low amounts of

acid functionality due to the higher stability of the coating [106].

Nitrogen containing plasma polymers obtained from the polymerisation of al-

lylamine were also shown to promote the attachment of human keratinocytes,

although attachment did not attain the level achieved on the acid function-

alised surfaces [107]. On the contrary, hydrocarbon plasma polymers ob-

tained from the polymerisation of 1,7-octadiene did not promote keratinocyte

attachment, possibly due to the hydrophobicity of the coatings and/or the

hydrocarobon groups introduced to the surface [107]. The preference ex-

hibited by keratinocytes towards carboxylic acid and nitrogen containing

plasma polymers compared to hydrocarbon surfaces confirms other findings

obtainied with fibroblast cells [108, 109].

Compared to other tissue engineering applications, the design of effective

scaffolds for chronic wound applications introduces an additional significant

challenge that has not yet been addressed. The transfer and ingrowth of

fibroblasts and keratinocytes from the wound bed onto and within the dress-

ing has to be prevented to avoid wound re-opening upon dressing removal.

While the dressing has to enhance cell proliferation within the wound bed,

a strategic surface chemistry needs to be designed to impair cell attachment

and growth onto and within the fibres.

The majority of the studies available in the literature on the interaction of

47 / 223


skin cells with plasma polymerised substrates were performed by seeding a

solution culture of the cells onto the modified surface. The study of the mech-

anisms of transfer and ingrowth of cells onto and within fibrous substrates

with different surface chemistries requires a different experimental approach

that involves the exposure of the mesh to an underlying cell culture that

mimics a wound bed. This can be achieved through the development of in

vitro wound models, as described in the following section.

1.8 In vitro Wound Models

The use of experimental wound models has become inevitable to investigate

the mechanisms involved in the healing process as well as for testing product

safety and efficacy during the development of therapeutics for clinical use

[110]. Gottrup et al. reviewed the most widely used strategies to produce

wound models, including in vitro and in vivo approaches and artificial models

[110]. In vitro models are most frequently chosen by researchers as they are

generally rapid, simple, relatively inexpensive and involve minimal ethical

considerations [110]. Several authors have developed in vitro wound models

by culturing cells (keratinocytes or fibroblasts) in monolayer directly on the

plastic surface of culture dishes; wound closure is simulated by creating de-

fects in the monolayers using sterile pipette tips [111–113], razor blades [114],

needles [115] or also electrically [116]. Although the monolayer culture model

is important in basic research to assess cell migration and proliferation, such

studies do not accurately reflect the behavior, pathophysiology, or microenvi-

ronment of wounded skin [117]. The throughput and reproducibility of these

wound healing assays can often be limited as the scraping speed and the

geometry of the wounded region can vary among different experiments [118].

48 / 223


Cells in monolayer culture are in isolation, and as a result these studies do not

encompass the intricate interactions that occur in vivo, where different cell

types interact within the extracellular matrix in a complex three-dimensional

structure [117].

These limitations have been partially circumvented by in vivo models of

wounds, developed using animals including mice [119–122], pigs [123], and

sheep [124]. However, there are significant differences between the mech-

anisms of healing of humans and animals, and many molecules that play

important roles in the innate response in humans are absent in animal mod-

els [117]. In addition, there are ethical issues associated to the use of animals

as experimental models.

Advances in tissue engineering over the last 20 years have led to the develop-

ment of tissue-engineered skin models that closely resemble normal human

skin [117]. Zhang et al. reviewed various aspects of the state of the art

of human skin equivalents [125] and MacNeil discussed the advances and

the opportunities for tissue engineered skin [27]. The overriding function of

tissue-engineered skin is to restore the barrier function of the natural skin

when this has been severely compromised [27]. Most tissue-engineered skin

is produced by expanding fibroblasts and/or keratinocytes in the laboratory

and using them for clinical applications, such as treating burns or initiating

healing in chronic wounds [27, 126, 127]. In addition to the clinical uses,

there are many non-clinical research applications for reconstructed skin tis-

sues. These models have been used to study the mechanisms of various

cutaneous conditions, including skin graft contraction [128], melanoma [129],

chemical irritation [130], and skin pigmentation [131].

Although the literature offers a variety of established protocols for develop-

ing tissue-engineered skin for laboratory and clinical uses, the need exists to

49 / 223


optimize and adapt such methodologies for creating tissue engineered models

of dermal wounds. To the best of our knowledge tissue engineered models

that mimic non-infected dermal wounds at different depths and degrees of

severity are not yet available.

Although such models cannot completely remove the need for animal ex-

perimentation (because of the lack of immune and circulatory responses),

they would constitute a valuable platform for laboratory research that can

reduce animal testing while overcoming the limitations of traditional in vitro

models. These models could be used for a variety of research applications,

including the study on the mechanisms of skin cell transfer and ingrowth

onto an within electrospun meshes.

Another important aspect, which has not been deeply explored yet is the ef-

fects induced by electrospun meshes on skin cells and bacteria in co-cultures.

The most complete studies on electrospun wound dressings involve separate

in vitro experiments on skin cells and bacteria. These tests constitute valu-

able tools for studying the cytotoxicity as well as antimicrobial activity of

electrospun meshes. However, they are performed in highly defined and con-

trolled culture conditions, which do not reproduce the real environment of

chronic wounds [19].

Strategies for establishing an in vitro chronic wound model by co-culturing

various types of skin cells and bacteria can be found in the literature, but

those models have not yet been used for testing electrospun dressings. Duell

et al. provided an overview on epithelial cells co-culture models for study-

ing infectious diseases in vitro [132]. Wiegand et al. established an in vitro

infected chronic wound model by co-culturing human keratinocytes and S.

aureus [19]. Hill et al. developed an in vitro model of a chronic wound

biofilm by co-culturing various species of bacteria [36]. Shepherd et al. devel-

50 / 223


oped three-dimensional tissue-engineered models of bacterial infected dermal

wounds by seeding keratinocytes and fibroblasts in human skin and infecting

the composites with two bacterial species. The model was developed to study

any cutaneous invasive bacterial or fungal infection [117, 133].

Studies of the responses of nanofibrous meshes in co-cultured experiments

between skin cells and bacteria could bring new insights that cannot be de-

termined through traditional single-culture methods. This could allow fur-

ther improvements to the design of a new generation wound care dressings

considering the fact that cells and bacteria have different dimensions and

most likely will respond differently to fibers of nanoscale dimensions [17].

1.9 Aims & objectives

The overall aim of this thesis lies in the design, fabrication and testing of

electrospun wound dressings with adequate mechanical and physico-chemical

properties to induce healing and with the additional capability of reducing

the bacterial load in the wound bed. In order to achieve such result, five

main objectives were defined and constitute the general guideline behind the

experimental works:

1. Electrospinning of fibrous meshes with controlled structure and mor-

phology. The electrospinning setup was designed and installed. Dif-

ferent process variables were optimised in order to reproducibly create

both micro and nano scale fibre meshes. This involved varying a range

of parameters, including solution properties, applied voltage and solu-

tion flow rate.

2. How do bacteria responsible for wound infections interact with elec-

trospun meshes? Bacterial behaviour might be affected by fibre mor-

51 / 223


phology and surface chemistry, as these properties have been proven to

affect skin cell adhesion and proliferation.

Various microbiological assays were optimised for understanding the

mechanisms of adhesion, spreading and proliferation of bacteria inter-

acting with micro/nanofibrous electrospun materials.

Surface modification processes were performed to provide specific chem-

ical functionalities to the surface of the meshes and the response of

bacterial cells to the surface modified substrates was investigated.

3. Cells interaction with electrospun meshes. Unlike tissue engineering ap-

plications, cell migration and ingrowth within the fibrous structure has

to be prevented when the mesh is in contact with the wound bed to pre-

vent wound re-opening upon dressing removal. The transfer of skin cells

onto and within electrospun meshes with various surface chemistries

was investigated to understand the mechanisms regulating this phe-

nomenon and to design a device capable of preventing cell ingrowth.

4. Four in vitro chronic wound models were developed by co-culturing

various types of skin cells and bacteria in order to reproduce wounds

at different depth and severity. Electrospun meshes with different sur-

face chemistries were tested on each model to investigate skin cell and

bacterial transfer mechanisms.

1

1

The work presented in this chapter has been partially published in the review article: M.Abrigo, S. L. McArthur, P. Kingshott, ”Electrospun Nanofibers as Dressings for ChronicWound Care: Advances, Challenges, and Future Prospects,” Macromol. Biosci., Vol. 14,pp. 772792, 2014

52 / 223

http://pubs.acs.org/doi/abs/10.1021/acsami.5b00453




Chapter 2

Experimental methods and

techniques

Contents2.1 Electrospinning . . . . . . . . . . . . . . . . . . . . 54

2.1.1 Electrospinning apparatus . . . . . . . . . . . . . . 54

2.1.2 Fibre fabrication . . . . . . . . . . . . . . . . . . . 58

2.2 Plasma polymerisation . . . . . . . . . . . . . . . 61

2.3 Bacterial culture techniques . . . . . . . . . . . . 64

2.4 Cell culture techniques . . . . . . . . . . . . . . . 66

2.5 Wound models . . . . . . . . . . . . . . . . . . . . 69

2.5.1 De-epidermisation of STS . . . . . . . . . . . . . . 70

2.5.2 Decellularisation of STS . . . . . . . . . . . . . . . 71

2.5.3 Model of superficial partially de-epidermised wounds 71

2.5.4 Model of superficial de-epidermised wounds . . . . 72

2.5.5 Model of deep wounds . . . . . . . . . . . . . . . . 73

2.5.6 3-Dimensional deep infected wound . . . . . . . . . 74

2.6 Characterisation . . . . . . . . . . . . . . . . . . . 76

2.6.1 Physico-chemical characterisation . . . . . . . . . . 77

2.6.2 Biological characterisation . . . . . . . . . . . . . . 79

53

2. Experimental methods and techniques

This chapter presents methodologies and experimental techniques used for

the research work presented in this thesis. It describes materials, methods

and techniques used for the installation of the electrospinning set up; the

fabrication, surface modification and characterisation of micro/nanofibrous

meshes; and the biological experiments conducted using bacteria (E. coli, P.

aeruginosa and S. aureus) and fibroblasts and keratinocytes.

2.1 Electrospinning

2.1.1 Electrospinning apparatus

The electrospinning technique is based on the application of a high voltage

(HV) (typically 15-30 kV) to a solution of the selected polymer. The solution

is extruded by a syringe, which is positioned at a selected distance (10-50

cm) from a target, acting as a collector of the fibres. The applied high

voltage generates a static electric field between the polymer solution and the

collector, inducing the formation of a fine charged polymer jet which moves

towards the collector while the solvent evaporates. Ultrafine dry fibres are

thus obtained in form of a fibrous mesh [134]. The electrospinning set up

used in this study was designed to achieve a versatile system, able to adapt to

both horizontal and vertical collection configurations and to different types of

collectors. The electrospinning set up is composed of three main components:

1. Spinneret, which is the unit through which the polymer solution is

extruded. As shown in Figure 2.1 the spinneret was obtained by con-

necting a syringe pump (model KDS Legato 111) to a 5 ml disposable

plastic syringe. A 8 cm long Tygon R FEP-lined tubing (1/8” inner

diameter, 1/4” outer diameter purchased from John Morris Scientific,

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NSW, Australia) was used to connect the syringe to a blunt needle

which was inserted in a poly(methyl methacrylate) (perspex) support

that can be horizontally and vertically moved to place the needle in the

desired position. Male and female polyvinylidene fluoride (PVDF) Luer

fittings (John Morris Scientific, NSW, Australia) were used to securely

connect the tubing to the syringe and to the needle respectively. Three

types of blunt needles were used: stainless steel 22 ga (51 mm long)

and nickel 24 ga (25.5 mm long) needles were purchased from Sigma-

Aldrich (NSW, Australia); stainless steel 18 ga (38 mm long) needles

were obtained from Livingstone International (NSW, Australia).

2. HV power supply. This unit is responsible for the high voltage that

has to be applied at the tip of the needle. The power supply (model

CZE2000) was purchased by Spellman, Hauppauge, NY; output voltage

and current were 0-30 kV and 0-300 µA respectively; polarity could

be manually reversed. The power supply was connected to the tip

of the needle through a crocodile clip soldered to a 20 cm long extra

high tension cable (model Belden Wire & Cable, Mouser electronics,

Austrlaia). The cable is represented as a red line in Figure 2.1.

3. Collector, which is the grounded unit onto which fibres are deposited.

Different collectors can be used depending on the desired alignment of

the fibres. For the electrospinning of non-woven meshes, a stainless steel

square plate collector (20 x 20 x 0.2 cm3) was fabricated, as shown in

Figure 2.1. For the spinning of fibres aligned along a specific direction,

a rotating mandrel was designed and fabricated. The rotating mandrel

(Figure 2.2) was fabricated from an aluminium hollow tube (70 mm

diameter x 150 mm long). Two aluminium plates were used to seal the

55 / 223


Figure 2.1: Schematic representation of the assembly of the componentsconstituting the electrospinning set up.

edges of the tube and to insert an aluminium bar (20 mm diameter x

500 mm long). Grub screws were used to connect the bar to the tube

preventing reciprocal movements. The system was then assembled into

a Polytetrafluoroethylene (PTFE) support that was securely screwed at

the bottom of the electrospinning unit to prevent undesired vibration

movements. To activate the rotation of the mandrel the bar was con-

nected to a motor (micro motor used in printer, copy machine) through

a motor belt. The motor was powered by a power supply (model PS-

3010D) purchased from Wavecom Instruments. Syringe pump, HV

power supply and collector were connected to the ground.

The three components were assembled inside a home-made safety box built

to protect the user from the high voltage. The front door of the box was fab-

ricated using a polycarbonate sheet, while the rest of the box was assembled

using perspex sheets. The polycarbonate material was chosen for the front

door due to its high mechanical resistance to ensure user’s protection. To

prevent user’s direct exposure to high voltage, the front door of the box and

56 / 223


Figure 2.2: Photograph of the rotating mandrel, used to collect fibres alignedalong one direction. The black and red cables connect the motor to a power

supply, while the green cable ensures the connection of the collector to the ground.

57 / 223


the power supply were connected to a home-built interlock safety system that

was placed at the edge of the safety box. The door of the box was designed

and built to slide within a rail along the box; when in place and securely

closed, the door triggers the interlock, allowing the use of the high voltage.

If the door is open, the interlock resets the voltage, thus ensuring the user’s

safety. The electrospinning apparatus was placed under a chemical safety

hood. Figure 2.3 is a photograph of the system after installation that shows

the various components of the electrospinning rig.

Figure 2.3: Photograph of the electrospinning set up after installation. The redcable terminating with a crocodile clip connects the needle to the HV power

supply, positioned on the top of the safety box. The needle is held in place by aperspex support connected to a wood system that allows to move the needle

horizontally and vertically. The teflon tube connecting the syringe to the needlepasses through a hole drilled in the safety box.

2.1.2 Fibre fabrication

Chemicals Polystyrene PS (Molecular weight (MW) = 250,000) was pur-

chased from Acros Organics (VIC, Australia); ethanol, chloroform and dimethyl-

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formamide (DMF), AR grade, 100% purity, were supplied from Science Sup-

ply Australia (VIC, Australia). Sodium dodecyl sulfate (SDS) was obtained

from Chem-Supply (SA, Australia); cetyltrimethylammonium bromide (CTAB)

and hexadecyltrimethylammonium bromide (HTAB) were purchased from

Sigma-Aldrich (NSW, Australia).

Electrospinning of polystyrene The optimisation of electrospinning pro-

cess parameters and polymer solution properties was undertaken by the elec-

trospinning of PS solutions in either chloroform at concentrations 20, 30, and

35% w/v or in DMF at concentrations 10, 15, 20, 30 and 35% w/v.

Various process parameters (voltage, flow rate, needle-collector distance (N-

C)) were tested to find the best combination for each polymer solution. The

electrospinning apparatus was established in a horizontal configuration and

blunt needles with diameters 18, 22, and 24 ga were used. Both the square

plate and the rotating mandrel were used to collect the meshes; alluminium

foil was used to cover the surface of the collectors to facilitate the removal

of the deposited meshes. The electrospinning was performed for 5 minutes

and subsequently the aluminium foil was removed from the collector and cut

into 0.5 x 0.5 cm2 square pieces that were used for the characterisation of

the fibres. During the electrospinning, the temperature in the laboratory was

measured through a digital thermometer and maintained constant through

the heating / air conditioning system. The humidity was also measured using

a digital hygrometer.

The study on the influence of fibre diameter on bacterial behaviour was per-

formed using meshes fabricated through the electrospinning of PS solutions

in DMF at concentrations 15, 20, and 30% w/v. To fabricate nanofibres

without inducing the formations of defects within the mesh, such as beads or

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polymer agglomerates, the 20% w/v solution was increased by adding 0.1%

w/v SDS.

The details on the optimisation of the process parameters for the fabrica-

tion of fibres with controlled diameter are reported in chapter 3, section 3.1.

Electrospinning was performed using the plate collector. To fabricate meshes

with suitable thickness (∼ 1 mm) to enable handling during the bacterial as-

says, the electrospinning of the solutions at concentrations 10% and 20% w/v

was performed for 4 hours; 2 hours and 30 minutes for the 15% w/v solution

and 1 hour for the 30% w/v solution. The side of the mesh which was ad-

hering onto the aluminium foil will be referred as the back of the mesh.

The conductivity of the polymer solutions that were electrospun was mea-

sured using the SevenCompact S230 conductivity meter from Mettler Toledo

(VIC, Australia).

After electrospinning, the foil was cut into 2 x 2 cm2 squares which were

immersed into ethanol aqueous solution (70 % v/v) for 30 seconds, with the

mesh facing down. The immersion in ethanol solution allowed the gentle

removal of the aluminium foil from the back of each mesh using tweezers.

Meshes were then transferred into 6-well plates, facing up. The meshes fab-

ricated from the 20% w/v PS solution in DMF (voltage = 20 kV; flow rate =

800 µL/h; needle-collector distance = 20 cm; needle inner diameter = 24 ga)

were used for the fibre surface modification through plasma polymerisation

and for the study on the influence of fibre surface chemistry on bacterial

behaviour and skin cell transfer.

Prior to undertaking any biological assay, the electrospun meshes were ster-

ilised in ethanol solution (70% v/v in Milli-Q (MQ) water) for 30 minutes

and rinsed 3 times with MQ water.

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2.2 Plasma polymerisation

Chemicals For the plasma polymer coating of the electrospun meshes the

following monomers were used: acrylic acid (molecular formula C3H4O2,

99.5% purity, MW 72.06) and 1,7-octadiene (molecular formula C8H14, 98.50%

purity, MW 110.20), supplied by Acros Organics (VIC, Australia); allylamine

(molecular formula C3H7N , > 99.5% purity), purchased from Sigma-Aldrich

(NSW, Australia); and eucalyptol or 1,8-cineole (molecular formula C10H18O,

99.5% purity) supplied by FGB Natural Products (VIC, Australia). The

molecular formula and structure of the monomers are summarised in Table

2.1.

Monomer Molecular formula Molecular structure

Acrylic acid C3H4O2

1,8-Cineole C10H18O

1,7-Octadiene C8H14

Allylamine C3H5NH2

Table 2.1: Molecular formula and structure of the monomers used for theplasma polymerisation of the PS meshes.

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Plasma polymer coating of polystyrene meshes Plasma polymerisa-

tion was carried out in a custom-built stainless steel T-shaped reactor (Figure

2.4) with stainless steel end plates sealed with Viton O-rings, as previously

reported by Salim et al. [135]. The reactor consisted of an aluminium disc

electrode (170 mm diameter) aligned perpendicular to the T-shaped cham-

ber. The plasma was ignited within the reactor using a 13.56 MHz genera-

tor coupled to an aluminium internal disc electrode via a matching network

(Coaxial Power Systems, UK). The radio frequency power source was coupled

to the reactor via an impedance matching network (Coaxial Power Systems,

UK). The pumping system consisted of an Edwards RV12 single stage rotary

pump. To prevent damage of the pump due to the monomer fragments, a

cold trap cooled by liquid nitrogen was placed between the chamber and the

pump. The flow rate of the monomer was controlled using a fine or ultra-

fine gas flow control needle valve (Chell Instruments Ltd, England) and the

plasma unit pressure monitored using a Pirani gauge (Edwards, U.K.).

Figure 2.4: Photograph of the plasma polymerisation reactor. The pressure inthe reactor chamber is brought to 1x10−3 mbar through the vacuum pump. Theneedle valve constitutes the inlet for the volatilised liquid monomer into thechamber. The plasma is generated when an electric field at radio frequency(13.56 MHz) is ignited through the electrode, producing a glow discharge that

ionises a fraction of the molecules

The schematic representation of the plasma polymerisation process is shown

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in Figure 2.5. The meshes were placed in the reactor on an alluminium

support, with glass slides positioned at the corners of each mesh to hold

them in place during the coating process. The reactor chamber was evacu-

ated down to base pressure (1x10−3 mbar). Prior to plasma polymerisation,

each monomer was degassed using three freeze-pump-thaw cycles. Monomer

vapour was then introduced to the chamber and an operating pressure of

2x10−2 mbar was maintained at the defined monomer flow rates. After de-

position, monomer flow was maintained for another 5 min to quench any

radicals on the surface [136]. The monomer valve was then closed and the

plasma unit evacuated to remove any residual monomer from the system.

Plasma deposition conditions were 20 W deposition power, 2 sccm monomer

flow rate, and 20 minutes deposition time for each monomer. Air plasma

was also performed at 20 W for 5 minutes. The side of the mesh which was

plasma coated will be referred as the front of the mesh.

Figure 2.5: Schematic representation of the plasma polymerisation process ofthe electrospun meshes. When the monomer is introduced in the chamber, theignition of the electric field generates electrons, ions, free radicals, photons andmolecules in both ground and excited states. The reactive species impinge on thesurface of the substrate creating reactive sites within the plasma zone which are

available for the covalent attachment of other species.

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2.3 Bacterial culture techniques

Chemicals 1.5% w/v tryptic soy agar (TSA) and tryptic soy broth (TSB)

were purchased from ThermoFisher Scientific (VIC, Australia) and used for

the culture of E. coli, S. aureus, and P. aeruginosa (bacterial strains were

clinical isolates purchased from American Type Culture Collection: E. coli

ATCC 25922 (Seattle 1946); P.aeruginosa ATCC 10145 (MDB strain BU

277); S. aureus ATCC 25923 (Seattle 1945)). Brain heart infusion agar was

obtained from Oxoid Limited (UK) and used for the culture of P. aeruginosa

(bacterial strain SOM-1) on the infected wound model. Phosphate buffered

saline (PBS) and glycerol (99.5 % purity, MW 92.09) were purchased from

Sigma-Aldrich (NSW, Australia). Sodium chloride (NaCl) 99.7 % purity

grade was obtained from Honeywell Riedel-de Han (UK).

Bacterial culture To start the experiments, a loop of bacteria was ob-

tained from the original ATCC stock solution kept in a -80 ◦C freezer. Liquid

broth media were inoculated with the bacteria and incubated overnight at

37 ◦C and 100 r.p.m. The overnight inoculum was diluted 1:1 using a glyc-

erol solution (30 % v/v) and stored in at -80◦C for further experiments. To

investigate the influence of fibre diameter on bacterial behaviour 1.5% TSA

plate was inoculated with E. coli thawed from the -80◦C glycerol stock. The

plate was incubated for 18 hours at 37◦C. A single bacterial colony was trans-

ferred from the agar plate into 30 ml of TSB in a 50 ml tube. The culture

was incubated for 18 hours (37◦C, 120 rpm) and consequently centrifuged for

15 minutes (25◦C, 2480 relative centrifugal force (rcf)). The obtained pel-

let was resuspended in 30 ml of clean TSB and diluted up to optical density

O.D.600nm=0.3. For the solution experiment, 3 ml of the inoculated TSB was

transferred on the electrospun meshes in six well plates and incubated (37◦C,

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120 rpm) for 1 hour. For the agar experiments, 100 µL of the broth culture

was transferred onto an agar plate and spread onto the surface of the plate

through a sterile spreader. The plate was incubated for 18 hours at 37◦C.

After incubation a confluent biofilm of E. coli cells was obtained. As the

fibres were spun onto a collector surface, the resulting mesh had an orienta-

tion, with the fibres that were in immediate contact with the collector being

slightly deformed due to the contact. As such, in the study, the meshes were

placed on the agar with the front of the mesh facing down onto the biofilm

and the collector-surface facing up. Meshes were incubated facing down on

the biofilm for 1 hour at 37◦C. After incubation, the culture was observed

for the presence of an inhibitory ring around the meshes. A commercially

available antibacterial silver releasing mesh (UrgoTul Ag/Silver, France) was

used as control.

A schematic representation of the agar experiment is provided in Figure 2.6.

The same procedure was repeated for P. aeruginosa and S. aureus.

Figure 2.6: Schematic representation of the bacterial agar culture experimentdesigned to investigate the transfer of bacterial cells onto and within electrospun

meshes.

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The agar experiment was also performed to investigate the influence of the

plasma coated meshes on E. coli attachment and growth. The plasma coated

meshes and PS control mesh were incubated facing down on the biofilm for

1 hour at 37 ◦C. After incubation, the culture was observed for the presence

of an inhibitory ring around the meshes.

For the infection of the 3-dimensional wound model, P. aeruginosa was

thawed from a -80 ◦C stock and centrifuged for 15 minutes (25 ◦C, 2480

relative centrifugal force (rcf)). The pellet was then resuspended in PBS

and diluted up to 109 CFU/mL. 100 µL of the bacterial dilution were then

transferred onto the wound model described below, which was incubated for

18 hours at 37 ◦C.

2.4 Cell culture techniques

The work involving human cell culture and tissue engineering techniques

was designed and performed during a 3 month overseas research placement

at the Kroto Research Institute (The University of Sheffield, UK), under the

supervision of Prof. Sheila MacNeil, Prof. Ian Douglas, Dr. Anthony Bullock

and Dr. Marc Daigneault.

Materials Human dermal fibroblasts and human dermal kertinocytes were

primary cells isolated from human skin biopsy.

Human skin for cell isolation was obtained from the trimmed edges of spit-

thickness skin grafts in the treatment of major burns or from patients under-

going breast reductions and abdominoplasties (Northern General Hospital,

Sheffield, UK) who gave consent for the use of their skin for research pur-

poses. All tissue was banked and used on an anonymous basis under HTA

Research Tissue Bank License Number 12179.

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The de-epidermised and de-cellularised dermis (DED) that was used for the

3-dimensional infected wound model was also sourced from human skin biop-

sies (Northern General Hospital, Sheffield, UK). DED varied in thickness due

to harvesting methods but was generally 2-3 mm deep. The Split Thickness

Skin (STS) that was used to produce the other wound models was obtained

from European Skin Tissue Bank, The Netherlands. The thickness of the

graft was ∼ 1 mm.

Fibroblasts were cultured in fibroblasts culture medium (DMEM high glu-

cose (4500 mg/L glucose), Sigma-Aldrich, UK) supplemented with 10% v/v

foetal calf serum (FCS) (Biowest Biosera, uk), 2x10−3 M l-glutamine (Sigma-

Aldrich, UK), 0.625 µg/mL amphotericin B (Sigma-Aldrich, UK), 100 IU/mL

penicillin and 100 µg/mL streptomycin (Sigma-Aldrich, UK)).

The culture of keratinocytes required the use of Green’s medium (DMEM

high glucose, Sigma-Aldrich, UK) and Ham’s F12 medium (Sigma-Aldrich,

UK) in a 3:1 ratio supplemented with 10% v/v foetal calf serum (FCS) (Biow-

est Biosera, UK), 10 ng/mL human recombinant epidermal growth factor

(Sigma-Aldrich, UK), 0.4 µg/mL hydrocortisone (Sigma-Aldrich, UK), 10−10

M cholera toxin (Sigma, UK), 1.8x10−4 M adenine (Sigma-Aldrich, UK),

5 mg/mL insulin (Sigma-Aldrich, UK), 5 µg/mL apo-transferrin (Sigma-

Aldrich, UK), 2x10−7 M 3,3,5-tri-idothyronine (Sigma-Aldrich, UK), 2x10−3

M glutamine (Sigma-Aldrich, UK), 0.625 µg/mL amphotericin B (Sigma-

Aldrich, UK), 100 IU/mL penicillin and 1000 µg/mL streptomycin (Sigma-

Aldrich, UK).

Phosphate buffered saline (PBS) was obtained from Oxoid, Unipath Ltd,

UK; trypsin/EDTA and thiazolyl blue tetrazolium bromide were purchased

from Sigma-Aldrich, UK; collagenase A was purchased from Boehringer-

Mannheim, UK.

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Cell isolation Fibroblasts and keratinocytes were isolated as described by

Ghosh et al. [137]. Briefly, for the keratinocyte culture, the skin graft sam-

ples were cut into about 0.5 cm2 pieces and incubated overnight at 4◦C in

0.1% w/v trypsin/EDTA solution. Epidermal and dermal layers were sep-

arated and the keratinocytes collected in FCS by scraping the upper layer

of the dermis and the lower layer of the epidermis. The cell suspension was

centrifuged at 200 g for 5 minutes and the pellet resuspended in Green’s

medium. The cells were transferred in T75 flasks that were seeded approx-

imately 1 hour earlier with 1x106 irradiated 3T3 (i3T3) murine fibroblasts

used as a feeder layer. Keratinocytes were subsequently cultured at 37◦C, 5%

CO2 in a humidified atmosphere until 70-80% confluent. Cells were serially

passaged to a maximum of passage 3. Kertinocyte suspensions used for the

wound models were obtained by trypsin/EDTA release [133].

For the fibroblast culture, cells were harvested from the skin graft via colla-

genase digestion. The dermal specimens were washed in PBS solution and

minced finely. The mince was incubated overnight in 10 ml of 0.05% collage-

nase A solution at 37◦C, 5% CO2 in a humidified atmosphere. The next day

the suspension was centrifuged at 200 g for 5 minutes. The obtained pellet

was resuspended in fibroblast culture medium and placed in T75 flasks. Fi-

broblasts were cultured at 37◦C in a 5% CO2 humidified atmosphere, cells

were used in experiments between passage 4 and 9. Fibrobast suspensions

used for the wound models were obtained by trypsin/EDTA release [133].

Well plate transfer experiment 2 mL of the fibroblast cell culture

(density 20000 cells/mL) was transferred into six well plate tissue culture

polystyrene (TCPS) dishes. The plate was incubated at 37 ◦C, 5% CO2 in

a humidified atmosphere until 70-80% confluency was reached. The 2x2 cm2

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sterilized plasma coated meshes and PS control mesh were gently laid facing

down on the confluent layer of cells and incubated at 37 ◦C, 5% CO2 in a

humidified atmosphere for 3 days. A sterile 2 x 2 cm2 metal grid was used to

maintain the meshes in contact with the bottom of each plate (Figure 2.7a).

1 mL of the keratinocyte cell culture (density 20000 cells/mL) containing

irradiated i3T3 feeder cells in same density was transferred into TCPS 12

well plates in Green’s medium and cultured until 70-80% confluence. The

plasma coated meshes and PS control mesh were laid facing down on the

cell culture. A sterile metal ring (1 cm inner diameter) was used to hold the

meshes in contact with the confluent layer of cells (Figure 2.7b).

After incubation, meshes were removed with sterile tweezers and transferred

into clean plates. Meshes and cultured plates were rinsed three times with

sterile PBS. The MTT assay was then performed on both meshes and cul-

tured plates to investigate the transfer of viable cells from the bottom of the

plates onto the meshes (see section 2.6.2 for details on the MTT assay).

Experiments were repeated three times and a different cell donor was used

for each experiment.

2.5 Wound models

Four different in vitro tissue engineered models were developed to reproduce

wounds at different depth and complexity. The models were developed by

culturing fibroblasts and/or keratinocytes in DED by adapting and optimis-

ing the protocols reported by Gosh et al. [137] and MacNeil et al. [133].

Experiments were repeated three times for each model using different cells

donors each time.

To remove the epidermis from split thickness human skin the protocol re-

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Figure 2.7: Schematic illustrating the well plate transfer experiment: (a) meshexposed to a confluent layer of primary human dermal fibroblasts. A metal gridwas used to hold the mesh in contact with the culture; (b) mesh exposed to a

confluent layer of primary human dermal keratinocytes. A metal ring was used tohold the mesh in contact with the culture

ported by Chakrabarty et al. was followed [138]. Briefly, the glycerol pre-

served skin was washed several times with PBS until most of the glycerol

was washed away. The skin was then soaked in 1 M NaCl overnight at 37

◦C and the epidermis was separated from the skin surface using blunt sterile

forceps. The obtained de-epidermised derims (DED) was then washed thor-

oughly with PBS and then placed in medium at ◦C for 23 days to confirm

sterility.

2.5.1 De-epidermisation of STS

To remove the epidermis from STS the protocol reported by Chakrabarty et

al. was followed [138]. Briefly, the skin was washed several times with PBS

and subsequently soaked in 1 M NaCl overnight at 37 ◦C. The epidermis was

separated from the skin surface using blunt sterile forceps. The obtained

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de-epidermised dermis (DED) was then washed thoroughly with PBS and

incubated in fibroblast culture medium at 37◦C, 5% CO2 in a humidified

atmosphere for a minimum of 48 hours to confirm sterility [137, 139].

2.5.2 Decellularisation of STS

The DED was washed with PBS solution and incubated in 200 ml of PBS at

37◦C for 4 weeks. The PBS solution was changed twice a week. Hematoxylin

and Eosin (H&E) staining was performed to assess the extent of clearing

of the dermis of cellular material. The acellular DED was stored in PBS

solution at -20◦C [137].

2.5.3 Model of superficial partially de-epidermised

wounds

The DED was cut into 1 cm round specimens, which were placed into 12

well plates with the basement membrane of the skin facing up (Figure 2.8a).

A sterile metal ring (1 cm inner diameter / 0.79 cm2) was used to hold the

specimens at the bottom of the plates and create a culture well to retain cells.

1x105 fibroblasts and 3x105 keratinocytes in Green’s medium (500 µL) were

seeded inside the ring, while 800 µL of Green’s medium was added around

the ring. The specimens were incubated at 37 ◦C, 5% CO2 in a humidified

atmosphere for 48 hours.

Medium and rings were removed and the plasma polymer coated meshes

and PS control mesh were gently laid facing down on the skin specimens

(Figure 2.8a). The rings were re-positioned on the meshes and 500 µL of

Green’s medium was added inside the ring and 800 µL outside. The sixth

skin specimen was used as control, without deposing any mesh. The model

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Figure 2.8: Schematic illustrating the preparation of the tissue engineered skinmodels of: (a) superficial partially de-epidermised wound; (b) superficial

de-epidermised wound; (c) deep wound.

was incubated at 37 ◦C, 5% CO2 in a humidified atmosphere for 3 days.

Meshes were gently removed and transferred into clean plates. Meshes and

skin specimens were rinsed three times with PBS. The MTT assay was sub-

sequently performed on both meshes and skin specimens. The experiment

was repeated three times, developing the model with different fibroblast and

keratinocyte donors each time. A fourth model was developed for histology

characterisation of the skin specimens.

2.5.4 Model of superficial de-epidermised wounds

The DED was cut into six 1 cm round specimens, which were placed into

12 well plates with the basement membrane of the skin facing the bottom

of the plate (Figure 2.8b). A sterile metal ring (1 cm inner diameter / 0.79

cm2) was used to hold the specimens at the bottom of the plates and create a

culture well to retain cells. 1x105 fibroblasts cultured in fibroblasts medium

(500 µL) were seeded on the reticular dermis inside the ring, while 800 µL

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of medium was added around the ring. The specimens were incubated at 37

◦C, 5% CO2 in a humidified atmosphere for 48 hours.

Medium and rings were removed and the skin specimens were flipped in each

plate to have the BM facing up. The plasma coated meshes and PS control

mesh were gently laid facing down on the skin specimens (Figure 2.8b). The

rings were re-positioned on the meshes and 500 µL of medium was added

inside the ring and 800 µL outside. The sixth skin specimen was used as

control, without deposing any mesh. The model was incubated at 37 ◦C, 5%

CO2 in a humidified atmosphere for 3 days. Meshes were gently removed and

transferred into clean plates. Meshes and skin specimens were rinsed three

times with PBS.

The MTT assay was performed on both meshes and skin specimens.

The experiment was repeated three times, developing the model with differ-

ent fibroblast donors each time. A fourth model was developed for histology


2.5.5 Model of deep wounds

Dispase II (Roche Diagnostics Ltd., Burgess Hill, UK) was dissolved in

DMEM AQmedia (Sigma-Aldrich, UK) at concentration 2.5 mg/mL. The

solution was filtered with a 0.45 µm syringe filter. To digest the basement

membrane, the DED was incubated in dipase solution at 37 ◦C for 4 hours.

After incubation the DED was rinsed with PBS to remove any remaining

dispase.

The skin was cut into six 1 cm round specimens, which were placed into

12 well plates with the papillary dermis facing up (Figure 2.8c). A sterile

metal ring (1 cm inner diameter / 0.79 cm2) was used to hold the specimens

at the bottom of the plates and create a culture well to retain cells. 1x105

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fibroblasts cultured in fibroblasts medium (500 µL) were seeded inside the

ring, while 800 µL of medium was added around the ring. The specimens

were incubated at 37 ◦C, 5% CO2 in a humidified atmosphere for 48 hours.

Medium and rings were removed. The plasma coated meshes and PS control

mesh (1x1 cm2 squared samples) were gently laid facing down on the skin

specimens (Figure 2.8c). The rings were re-positioned on the meshes and

500 µL of medium was added inside the ring and 800 µL outside. The sixth

skin specimen was used as control, without deposing any mesh. The model

was incubated at 37 ◦C, 5% CO2 in a humidified atmosphere for 3 days.

Meshes were gently removed and transferred into clean plates. Meshes and

skin specimens were rinsed three times with PBS.

The MTT assay was performed on both meshes and skin specimens. The

experiment was repeated three times, developing the model with different

fibroblast donors each time. A fourth model was developed for histology


2.5.6 3-Dimensional deep infected wound

The 3 dimensional infected wound model was developed following the pro-

tocol reported by MacNeil et al. [133]. Briefly, the de-epidermised skin (∼

5 mm thick) was cut into six 1 cm2 round specimens, which were placed

into a 12 well plate within 12 mm tissue culture inserts with 4 µm pores in

the base (Greiner, UK). The inserts were suspended from the edges of 12

well plates into the wells. 800 µL of Green’s medium (bathing medium) was

added at the bottom of each well, surrounding the inserts; 100,000 fibrob-

lasts and 500,000 keratinocytes in 500 µL of antibiotic-free Green’s medium

(seeding medium), were seeded on each skin specimen. After 24 hours at 37

◦C, seeding and bathing media were replaced with fresh Green’s medium.

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After further 24 hours, the seeding medium was removed leaving the surface

of the specimens at the air-liquid interface. The bathing medium (800 µL)

was periodically changed every 48 hours. After 14 days at the air-liquid in-

terface (37 ◦C, 5% CO2), a thermal burn was generated in the center of each

skin specimen using an incandescent metal stick that was kept pressed onto

the specimens for 6 seconds.

The P. aeruginosa solution in PBS (100 µL, 109 CFU/mL) was added to

each specimen and the model was incubated at 37 ◦C for 18 hours. The

bacterial solution was removed and the specimens rinsed three times with

PBS. The plasma coated meshes and PS control mesh were gently laid facing

down on the infected skin specimens and 100 µL PBS was added on the top

of each mesh. The sixth specimen was left with no mesh, as control. The

model was incubated at 37 ◦C for 1 hour.

The meshes were gently removed from the specimens. Meshes and speci-

mens were rinsed three times with PBS. Meshes were immersed in 2 mL

of formaldehyde solution in PBS (3.4% v/v) for 10 minutes for cell fixation

and consequently immersed in propidium iodide (PI) (Life Technologies Ltd,

UK) solution (1:3000 in PBS; 15 minutes; room temperature) for fluorescent

labelling. PI is a fluorescent molecule that binds cellular DNA and emits red

fluorescence (emission wavelength 635 nm) when excited at 480-490 nm.

Each skin specimen was cut in half with a sterile scalpel. One half of each

specimen was immersed in 1 mL formalin solution (10% v/v in PBS) for 10

minutes and rinsed once with PBS, for histology characterisation. The other

half of each specimen was weighted and then immersed in 1 mL collagenase A

(Roche Diagnostics, Germany) and incubated at 37 ◦C for 2 hours, to break

the bonds of the bacterial cells attached to the skin. Specimens were then

finely minced with a scalpel and transferred into 1 mL PBS. TissueRuptor

75 / 223


(Qiagen) was used to further disrupt the tissues of the skin inducing the

detachment of the bacteria. The remaining bacterial solution in PBS from

each specimen was diluted 1:10 in PBS eight times; 10 µL of each dilution

was plated twice on an agar plate and incubated overnight at room temper-

ature. The plate was then transferred at 37 ◦C for 4 hours. Finally, the

colony-forming unit (CFU) of P. aeruginosa left by each mesh on the skin

specimens were counted. The described model was developed three times,

using different cell donors each time.

2.6 Characterisation

To achieve a control over the electrospinning parameters affecting the mor-

phological properties of the electrospun structures, meshes were characterised

through a combination of imaging techniques. Scanning electron microscopy

(SEM) , bright microscopy and atomic force microscopy (AFM) were used to

analyse fibre diameter, diameter distribution and presence of defects. X-ray

photoelectron spectroscopy (XPS) analysis was performed before any biolog-

ical assay to characterise fibre surface chemistry.

To qualitatively and quantitatively detect bacteria and skin cells transferred

onto and within electrospun meshes with different morphological and chemi-

cal properties various colourimetric and viability microbiological assays were

combined with SEM images.

Histology images were acquired to characterise the tissue engineered skin

models of wounds at different depths and severity.

76 / 223


2.6.1 Physico-chemical characterisation

AFM An Asylum Research MFP-3D atomic force microscope (CA, USA)

was used to image the electrospun meshes using tapping mode with ultra-

sharp silicon nitride tips (DNP-10 non-contact silicon cantilevers, Mikro-

Masch, Spain). Tips were ozone cleaned for 20 minutes prior use to remove

organic contaminants using UV-ozone Procleaner (BioForce Nanosciences,

Inc.). All images were processed (1st order flattening algorithm) and line

scans generated using NanoScope Analysis 1.40 software (Bruker AFM Probes,

MA, USA).

Bright microscopy Electrospun meshes were imaged using a LEICA DM

LB2 Light Microscope connected to a CCD camera (Spot 3-Shut Insight

QE, SciTech, WA, Australia). Images were processed using Spot Analysis

software (SPOT Imaging Solutions, MI, USA)

SEM The morphological properties of PS fibres electrospun with different

process paramters were investigated using SEM (ZEISS Supra 40 VP Carl

Zeiss SMT, Germany. EHT = 3kV). Prior to the SEM analyses, fibres were

gold coated (10-15 nm) using a Dynavac CS300 thermal deposition cham-

ber (Dynavac, MA, USA). For the measurement of fibre average diameter,

for each set of electrospinning parameters three meshes were electrospun; on

each mesh 3 SEM images were collected at the same magnification. On each

image 10 fibres were randomly selected and the diameter measured using

Image J software (ImageJ, http://rsb.info.nih.gov/ij/index.html). Therefore

the average fibre diameter of the meshes was calculated from 90 fibres in

total for each set of parameters tested.

After plasma polymerisation, SEM images of the coated meshes were ac-

77 / 223


quired to address any morphological modification induced by the plasma

polymerisation process.

SEM images were also used to explore initial bacterial adhesion and progres-

sive spreading and colonisation of PS electrospun meshes exposed to E.coli

culture. After incubation meshes were rinsed 3 times with MQ water, to

remove the bacteria which were not attached to the fibres. 2 ml of formalde-

hyde solution in PBS (3.4 % v/v) was added to the samples for 10 minutes for

bacterial fixation. Samples were rinsed once with MQ water and exposed to 3

subsequent rinses in ethanol aqueous solutions (EtOH) at increasing concen-

trations for dehydrolysis purpose; samples were finally left in pure ethanol for

5 minutes. When dry, meshes were mounted on the SEM support and gold

coated. Experiments were performed in triplicate; 10 images were acquired

on each mesh using a digital grid to select the areas to be imaged in an un-

biased manner. The same protocol was undertaken for imaging meshes with

different average fibre diameter and surface chemistry exposed to solution

and agar cultures of E. coli, S. aureus and P. aeruginosa.

XPS XPS analysis of the plasma coated meshes and PS control mesh was

performed using a Kratos AXIS NOVA spectrometer (Kratos Analytical Inc.,

Manchester, UK) using a monochromated Alkα X-ray source at a power of

150 Watts. An elliptical area with approximate dimensions of 0.3 mm 0.7

mm was analysed on each sample. All elements present were identified from

survey spectra (acquired at a pass energy of 160 eV). To obtain more detailed

information about chemical structure, high resolution spectra were recorded

from individual peaks at 20 eV pass energy. The atomic concentrations of

the detected elements were calculated using integral peak intensities and

the sensitivity factors supplied by the manufacturer. Data processing was

78 / 223


performed using CasaXPS processing software version 2.3.16 (Casa Software

Ltd., Teignmouth, UK).

High resolution C1s spectra were fitted with Gaussian-broadened Lorentzian

functions and peaks were charge corrected relative to the CHx component

at 285.0 eV. Peaks were restricted to full width half maximum (FWHM)

between 1.2-1.6 eV. For the C1s high resolution spectra of ppaAAc the peak

positions were fixed to a minimum position of 287.95 eV for carbonyl com-

ponent at 289.9 eV for carboxyl component. Five distinctive binding energy

shifts were used in curve fitting. Hydrocarbon component (C-C/C-H) is

found at 285.0 eV, hydroxyl component (C-OH, C-O-C) at 286.5 eV, car-

bonyl component (C=O, O-C-O) at 288 eV and carboxylic component (HO-

C*=O) at 289 eV. The carbon adjacent to the carboxylic group is shifted due

to the electronegativity of the carboxylic acid. Experimentally, a β shift or

secondary shift at 285.6 eV (C*COO) is incorporated in to the curve fitting

with peak area equivalent to the carboxylic group [140, 141].

2.6.2 Biological characterisation

Confocal microscopy OLYMPUS FV1000D Laser Confocal Scanning Mi-

croscope with OLYMPUS x40 and x100 objectives was used to investigate

the influence of fibre diameter and fibre surface chemistry on bacterial be-

haviour. After exposure to the bacterial cultures, the meshes were rinsed 3

times with NaCl 0.85 % w/v in MQ water, to remove the bacteria which were

not adhered. LIVE/DEAD assay was performed using the LIVE/DEAD Ba-

cLight, Bacterial Viability Kits (3.34 mM PI in dimethyl sulfoxide (DMSO)

and 20 mM of SYTO 9 in DMSO) purchased from Invitrogen Life Technolo-

gies (VIC, Australia). LIVE/DEAD is a two colours fluorescence assay of

bacterial viability. After staining, when excited at 480-490 nm, bacteria emit

79 / 223


green fluorescence (emission wavelength 500 nm) if alive, and red fluorescence

(emission wavelength 635 nm) if dead. LIVE/DEAD staining solution was

prepared according to the product instructions. 2 mL of the LIVE/DEAD

solution were added to each sample for 30 minutes (37◦C in dark). The stain-

ing solution was removed and samples rinsed once with PBS. For imaging,

selected filters were U-MNIBA filter (excitation 470-490 nm, green emission)

and U-MWIG2 filter (excitation 510-550 nm, red emission). Olympus Flu-

oViewer software (Olympus, NSW, Australia) was used for image capturing

and processing. The experiment was performed in triplicate; 10 images were

acquired on each mesh using a digital grid to select the areas to be imaged

in an unbiased manner.

The same microscope was used for imaging the meshes that were tested on

the 3 dimensional infected wound model and stained with PI. In this case

the U-MWIG2 filter only was used. 10 z-stack images were acquired on each

mesh using a digital grid to select the areas to be imaged in an unbiased

manner.

Bacterial counting ImageJ software (ImageJ, http://rsb.info.nih.gov/ij/

index.html) was used for the quantification of the PI stained bacteria present

on the meshes after exposure to the infected wound model. Each image stack

was projected along the z axis perpendicular to the image plane. The max-

imum intensity projection option was used, thus creating an output image

each of whose pixels contained the maximum value over all images in the stack

at the particular pixel location. Subsequently the threshold was adjusted on

each image to isolate the fluorescing bacteria while excluding background

signal and autofluorescence from the fibres. The ”Analyse particle” function

of the ImageJ software was used to count the number of fluorescing particles

80 / 223


present on the images. Figure 2.9 summarises the image processing steps

that were undertaken to quantify the number of bacterial cells present onto

the meshes. After projecting each z-stack image along the z axis (Figure

2.9a), a threshold was manually selected and applied to the image to isolate

the bacterial cells from the underlying fibres and the image noise (Figure

2.9b). Figure 2.9c shows the outline of the bacterial cells that were counted

using the ”Analyse particles” function of the ImageJ software.

A programming code capable of automatically performing the mentioned

analysis steps was designed and compared to the manual counting. The code

allowed the user to manually adjust the threshold for the images while auto-

matically implementing the z-projection command, followed by the particle

counting. In output the code provided the z-projected images; a summary

containing the total number of particles counted; and images showing the

outlines of the counted particles. The code is reported in Appendix A.

Figure 2.9: Image processing steps for the quantification of bacterial cellsattached onto the meshes: (a) projection of the z-stack image along the z-axis; (b)

application of threshold to isolate bacteria from fibres and noise; (c) particlecounting and outlines. Scale bar µm.

MTT andMTS assays MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium

bromide) and MTS (3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-

2-(4-sulfophenyl)-2H-tetrazolium) are colourimetric assays for assessing the

81 / 223


viability of bacterial and mammalian cells. They are performed by the addi-

tion of a premixed optimised dye solution to a cell culture. Enzymes produced

by living cells convert the yellow tetrazolium component of the dye solution

into a formazan purple product. In this work, these assays were adapted for

assessing the viability of bacteria adhering onto electrospun fibres.

CellTiter 96 Non-Radioactive Cell Proliferation Assay and CellTiter 96 AQue-

ous One Solution Cell Proliferation Assay supplied from Promega were used

for the MTT and MTS experiments respectively.

The MTT assay was performed to qualitatively confirm the presence of vi-

able bacteria within the meshes. A mesh not exposed to the bacterial culture

was stained with MTT as control. To remove bacteria which were not stably

adhering onto the fibres, the samples were rinsed 3 times with PBS after

incubation. Samples were transferred into clean 6-well plates. For the qual-

itative detection of bacteria, 2 mL of MTT dye diluted in PBS (1:10) were

added to the samples. Meshes were incubated at 37◦C in dark for 2 hrs.

After incubation, the MTT dilution was removed and pictures of the stained

meshes were taken.

The MTS assay allowed the quantification of viable microbial cells present

within the fibrous structure. A mesh not exposed to the bacterial culture

and stained with MTS was used as control. 2 mL of MTS dye diluted in

PBS (1:10) were added to the samples. Samples were incubated in the same

conditions used for the MTT assay. After incubation, 100 µL of MTS solu-

tion were transferred from the samples into a 96 well plate and absorbance

was measured at 490 nm through a FLUOstar Galaxy plate reader (BMG

Labtech, VIC, AU). Experiments were performed in triplicate.

The MTT assay was also used to investigate the mechanisms of transfer of fi-

broblasts and keratinocytes onto electrospun meshes with different chemistries

82 / 223


from cell culture plates and wound models. In this case, the MTT solution

was prepared by dissolving Thiazolyl Blue Tetrazolium Bromide (Sigma-

Aldrich, UK) in sterile PBS (0.5 mg/mL). In the well plate transfer ex-

periments, meshes and cultured plates were rinsed three times with PBS and

1 mL of MTT solution was added. Samples were incubated at 37◦C in dark

for 40 minutes. After incubation, the MTT solution was removed, samples

rinsed once with PBS and photographs of the stained materials were taken.

After taking the photographs, the MTT solution was dissolved from the sam-

ples (both meshes and substrates) by adding 1 mL of 2-Ethoxyethanol. The

system was incubated for 2 hours at room temperature. 100 µL of the sol-

ubilised MTT was transferred from each sample into a 96 well plate and

absorbance was measured at 570 nm using a plate reader.

For the wound model transfer experiments, the MTT assay was performed

by adding 500 µL of the dye solution to the meshes and the skin specimen.

Crystal violet assay The Crystal Violet (CV) Aqueous Stain (ProSciTech,

QLD, Australia) is a colourimetric assay based on the dissociation of CV in

aqueous solutions into CV+ and chloride (Cl-) ions. These ions penetrate

through the cell wall and cell membrane of both gram-positive and gram-

negative cells, staining the cells purple.

The assay was performed to investigate the transfer of E. coli, S. aureus and

P. aeruginosa cells from agar cultures onto electrospun meshes with different

average fibre diameter. After 1 hour incubation on the agar cultures, meshes

were removed from the plates. 5 mL of 1:100 dilution of crystal violet aque-

ous solution in NaCl 0.85% were added to each plate and incubated for 10

minutes in dark. The dilution was then removed and plates were rinsed three

times with NaCl 0.85%. Photographs of the plates were taken.

83 / 223


Histology on wound models Histology of tissue sections of the vari-

ous wound models was performed. Specimens were processed, embedded in

paraffin wax, sectioned to a thickness of 6 µm, mounted, and stained follow-

ing the Haematoxylin and Eosin (H & E) protocol previously reported by

McLaughlin [142]. Briefly, tissue sections were immersed in xylene followed

by 100% denatured alcohol (IMS); 70% IMS; and distilled water; samples

were then immersed in hematoxylin for 1 min 30 sec, rinsed with tap water

and dipped into eosin for 5 min. A series of consequent rinsing in tap water,

70% IMS, 100% IMS and xylene was carried out. DPX mounting medium

(Fisher Scientific, UK) was used to position glass cover slips over the stained

tissue sections prior to imaging. In addition, the histology sections of the skin

composites that were developed for the 3 dimensional infected wound model

were stained using the Gram staining protocol [143], that results by staining

purple the Gram-positive bacterial cells and pink the Gram-negative ones.

Tissue sections were first exposed to crystal violet for 1 minute followed by

iodide for binding the CV and trapping it in the cells. Rapid decolourisa-

tion was carried out using acetone, and carbol fuchsin was added as counter

staining.

84 / 223

Chapter 3

Electrospinning of polystyrene

meshes

Contents3.1 Optimisation of electrospinning parameters . . . 86

3.2 Electrospinning of nanofibres . . . . . . . . . . . 100

3.3 Characterisation of electrospinning apparatus per-formance . . . . . . . . . . . . . . . . . . . . . . . . 106

3.4 Electrospinning of aligned fibres . . . . . . . . . 109

3.5 Conclusions . . . . . . . . . . . . . . . . . . . . . . 111

This chapter aims to describe the home-built electrospinning set up and

optimise the process parameters for the fabrication of fibres with controlled

morphological properties. Polystyrene was selected as polymer to be electro-

spun due to the availability of several examples in the literature based on the

electrospinning of this material. The chapter describes the optimal solvents,

solution concentration and conductivity values to achieve continuous and

uniform spinnability; the influence of applied voltage, flow rate and needle-

collector distance on the spinnability of the solution and on the properties of

85

3. Electrospinning of polystyrene meshes

the resultant fibres is discussed. The optimal process and solution parameters

to fabricate smooth, round fibres with controlled diameter are highlighted;

the strategies to prevent the formation of defects in form of beads or polymer

agglomerate within the meshes are presented. The present chapter consti-

tutes a systematic study of a variety of variables and parameters that need

to be strategically tailored and tuned to control the electrospinning process

and fabricate meshes with the desired properties.

The work presented in this chapter has been partially published in the

journal article: M. Abrigo, P. Kingshott, S. L. McArthur, ”Electrospun

Polystyrene Fiber Diameter Influencing Bacterial Attachment, Proliferation

and Growth,” ACS Appl. Mater. Interfaces., Vol. 7, no. 14, pp. 7644-52,

2015

3.1 Optimisation of electrospinning parame-

ters

For the electrospinning of PS, process and solution parameters were initially

selected according to different studies found in literature [144–147]. The

choice of the optimal solvent for achieving a controlled spinnability of the

polymer was the first challenge faced. PS was initially dissolved in chloro-

form and four polymer concentrations were tested (20%, 30% , 35% and 40%

w/v). The electrospinning of the solutions was performed using a home-built

apparatus in the horizontal configuration (see chapter 2, section 2.1.1 for de-

tails on the electrospinning apparatus). A 22 Gauge needle was used for the

20% and 30% w/v solutions and a 18 Gauge needle for the 35% w/v and 40%

w/v solutions. The selected needle inner diameters allowed a continuous ex-

trusion of the solution.

86 / 223






Different values of the electrospinning process parameters were tested before

finding the best combinations that allowed continuous spinnability of the PS

solutions. Table 3.1 is a complete summary of the values of applied voltage,

needle-collector distance (N-C) and solution flow rate that were tested for

the four polymer concentrations. The spinnability and fibre morphological

properties resulting from each set of parameter combinations are also high-

lighted in Table 3.1.

The electrospinning of PS in chloroform at concentrations 20% and 30% w/v

was found unsatisfactory regardless of the process parameters. In fact, due

to the low viscosity of the solutions, the polymer droplet broke at the tip of

the needle and no polymeric filament could be formed. To increase solution

viscosity the polymer concentration was increased to 35% w/v. This solution

was successfully electrospun applying a voltage between 15 and 20 kV and

positioning the collector at 15-18 cm from the tip of the needle. A flow rate

of 800 µl/h induced the continuous extrusion of a polymer droplet at the

tip of the needle that led to the formation of a continuous filament from the

spinneret to the collector.

To compare the influence of the selected solvent on the spinnability of the

polymer, PS was also dissolved in DMF at various concentrations (Table

3.1). Due to its lower volatility, DMF allowed a better spinnability than

chloroform, resulting in an easier formation of the filament at the tip of the

spinneret. The optimal spinnability was achieved for the intermediate con-

centrations (15%, 20%, 30% , and 35% w/v); the lowest concentration (10%

w/v) could be electrospun at low voltages (10-15 kV) and the formation of

the mesh was not always continuous. No process parameter combination was

found satisfactory for the electrospinning of the highest concentration (40%

w/v), which always induced needle clogging.

87 / 223


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88 / 223


Con

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89 / 223


Figure 3.1: SEM images of PS fibres electrospun from 35% w/v solution inchloroform at different magnification. Scale bar: (a) 10 µm; (b) 2 µm; and (c) 1

µm.

For the intermediate concentrations, voltage values between 15 and 20 kV

successfully allowed filament formation and fibre deposition; values below this

optimal range caused needle clogging while higher values resulted in breakage

of the polymer droplet (electrospraying). The N-C was selected between 15

and 25 cm; lower distance did not allow the complete evaporation of the

solvent inducing the deposition of polymer droplets within the mesh. If the

N-C was over 25-30 cm the filament never reached the collector as it was

attracted towards other closer metallic grounded components present in the

electrospinning box (i.e. parts of the interlock safety system).

SEM images were used to study the surface morphology of the fibres obtained

from the electrospinning of the PS solutions in chloroform and DMF. Table

3.1 summarizes the morphological properties of the fibres obtained from the

electrospinning of the PS dissolved in the two solvents. Figure 3.1 shows the

fibres obtained from the 35% w/v PS solution in chloroform. The surface

of the fibres was not perfectly round (Figure 3.1a), it appeared flattened

and affected by the presence of rifts that induced a distortion of the fibre

morphology. Figure 3.1b exhibits the cracks present on the fibre surface as

well as a widespread porosity. The higher magnification provided by Figure

3.1c shows pores of a few nanometers in size spread over the entire surface

of fibre.

90 / 223


The use of DMF as solvent (Figure 3.2) resulted in smoother and more uni-

form fibres when PS was dissolved at concetrations 15%, 20%, 30% and 35%

w/v. Figure 3.2a shows electrospun fibres from the 35% w/v PS solution

with uniform round shape and even surface. No rifts or pores were present

on the fibre surface (Figures 3.2b and 3.2c). Due to the easier spinnability

and the more controllable surface properties of the fibres obtained when us-

ing DMF, this solvent was chosen for the experiments performed after these

results.

Figure 3.2: SEM images of PS fibres electrospun from 35% w/v solution inDMF at different magnifications. Scale bar: (a) 10 µm; (b) 2 µm; and (c) 1 µm.

A photograph of a PS mesh on aluminium foil placed on the plate collector

is shown in Figure 3.3. The mesh is pink in colour because rhodamine was

added to the polymer solution (1% w/v) for imaging purposes. The image

shows that after 5 minute electrospinning the mesh had an approximate

square shape, with a surface of about 30 x 30 cm2 and a thickness of 1-2

mm.

To understand the level of details on the morphological properties of the fi-

bres that various characterisation techniques can provide, non-woven meshes

fabricated through the electrospinning of 35% w/v PS solution in DMF were

imaged through bright microscopy, SEM and AFM.

Figure 3.4a shows a bright microscopy image of the PS mesh, where it is

possible to visualise the density of the fibrous structure, the organisation of

the fibres and the presence of defects, such as beads or polymer agglomerates

91 / 223


Figure 3.3: Photograph of the electrospun mesh obtained from PS solution inDMF (35% w/v). Rhodamine was added to DMF (1% w/v) prior electrospinningfor imaging purposes. After 5 minute electrospinning a mesh with approximate

square shape, surface of about 30 x 30 cm2 and thickness of 1-2 mm wasobtained. Scale bar 10 mm.

that can form within the fibrous network. In Figure 3.4b a single PS fibre is

imaged through SEM. The diameter of the fibre (Φ) can be precisely mea-

sured and the morphology of the surface be visualised. AFM images (Figure

3.4c) allow to accurately visualise an area of 1 x 1 µm2 of the surface of the

fibre for analysing morphological properties, including texture and roughness.

To evaluate the influence of polymer concentration on fibre morphological

properties, meshes electrospun from PS solutions in DMF at concentrations

10 %, 15%, 20%, 30% w/v and 35% w/v were analysed with SEM. SEM

images (Figure 3.5) were used to characterise fibre diameter, diameter dis-

tribution and presence of defects within the fibrous substrates. Table 3.1

summarizes the values of the average fibre diameter of the meshes fabricated

from the polymer solutions at different concentrations.

The lowest polymer concentration (10% w/v) resulted in the smallest fibre

diameter, in the range Φ = 300 ± 200 nm; intermediate concentrations of

15% and 20% induced an increase of the diameter to Φ = 900 ± 200 nm and

Φ = 1800 ± 200 nm respectively; the highest concentrations tested (30%

and 35% w/v) resulted in a noticeable increase of average fibre diameter and

92 / 223


Figure 3.4: (a) Bright microscopy, (b) SEM and (c) AFM images of fibreselectrospun from 35% w/v PS solution in DMF. Scale bar (a) 50 µm; (b) 10 µm;

and (c) 100 nm.

diameter distribution (Φ = 3000 ± 1000 nm).

The polymer concentration was also shown to affect the number and size of

beads within the meshes. In fact, a progressive increase in polymer concen-

tration resulted in the formation of fewer and smaller defects (Figure 3.5).

The lowest polymer concentration (10% w/v) induced the formation of large

defects along the fibres, spread throughout the mesh (Figure 3.5a), while the

number and size of defects progressively decreased with the increase of the

concentration to 15% and 20% w/v (Figures 3.5b and 3.5c). The electro-

spinning of the solution at the highest concentration (30% w/v) allowed the

fabrication of defect-free fibres (Figure 3.5d).

The morphology of the beads was also found to depend on the concentra-

tion of the polymer solution. Figure 3.6 shows a magnification of the beads

formed along fibres electrospun from PS solutions at different concentrations.

The 10% w/v concentration induced the formation of round defects, with the

bead diameter reaching up to 10 µm (Figure 3.6a); the surface of the beads

appeared covered by a widespread porosity with pores of few nanometers in

size. Porosity was also found on the surface of the beads formed from the

15% w/v concentration, as shown in Figure 3.6b. With the increase of the

polymer concentration, the beads appeared smaller (1-3 µm wide) with a

93 / 223


Figure 3.5: SEM images of PS meshes electrospun from solutions in DMF atdifferent concentrations: (a) C = 10% w/v, Φ = 300 ± 200 nm; (b) C = 15%

w/v, Φ = 900 ± 200 nm; C = 20% w/v, Φ = 1000 ± 100 nm; C = 30% w/v, Φ= 3000 ± 1000 nm. Scale bar 2 µm.

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Figure 3.6: SEM images showing the morphology and size of the beads alongfibres electrospun from (a) 10% w/v; (b) 15% w/v; and (c) 20% w/v PS solution

in DMF. Scale bar 1 µm.

more elongated rather than a spherical shape. With the 20% w/v concentra-

tion, the beads lost shape and appeared to have wider fibre diameters of few

hundreds nanometers. The yellow line in Figure 3.6c represents the edge of

the fibre, with the red dashed line showing the 400 nm widening of the fibre

diameter induced by the defect. The 20% w/v concentration also resulted in

the disappearance of the porosity on the bead surface.

The influence of applied voltage and N-C distance on average fibre diameter

was also investigated. The influence of flow rate could not be evaluated as

a significant change in the flow rate was found to impair the formation of

the fibres. The 10% w/v PS solution in DMF had to be excluded from this

study as significant changes in voltage and N-C distance from the optimal

values reported in Table 3.1 were found to impair the spinnability of the so-

lution. For the other three sets of concentrations (15%, 20% and 30% w/v)

two values of voltage (15 kV and 20 kV) and two values of N-C distance (15

cm and 20 cm) were tested and for each of them the average fibre diameter

was measured. These values were chosen as they allowed the spinnability of

the solutions at the concentrations of interest. As shown in Table 3.1 values

of voltage below 15 kV or above 20 kV were not satisfactory as they resulted

in needle clogging or electrospraying respectively. The N-C had to be kept

between 15 and 20 cm to ensure the deposition of fibres on the collector.

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For each polymer concentration, applied voltage and N-C distance did not

significantly affect average fibre diameter and diameter distribution (Figure

3.7). When applying different voltages keeping constant flow rate, N-C dis-

tance and polymer concentration, the average diameter did not significantly

change. The same response was obtained when the applied voltage was kept

constant and the N-C distance was varied. For this reason, for the experi-

ments performed subsequently, the electrospinning parameters were selected

to ensure the best spinnability, in terms of continuous and uniform forma-

tion of the fibres (Table 3.1). The concentration of the polymer solution was

tailored to match the desired morphological properties of the fibres.

Figure 3.7: Influence of applied voltage and N-C distance on the average fibrediameter of electrospun PS meshes.

Discussion The first challenge faced for optimising the electrospinning pa-

rameters consisted in choosing the most appropriate solvent for fabricating

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meshes in a reproducible manner, with controlled fibre morphology. PS scaf-

folds have been electrospun from a wide variety of solvents, including chlo-

roform, DMF, ethylacetate, methylethylketone, tetrahydrofuran and toluene

[148]. Chloroform was chosen as first solvent to test due to its high volatility

that reduces the chances of remaining trapped into the fibres [148]. This sol-

vent however resulted in flattened ribbon-like fibres with a significant porosity

spread out onto the fibre surface. Fibres in the form of ribbons with various

cross sections have been associated to the rapid evaporation of the solvent

[149]. The porosity of the fibres have been found to be dependent on atmo-

sphere humidity, polymer molecular weight [150] as well as solvent volatility

[151].

Chloroform was replaced with DMF, which is less volatile and allowed the

formation of round shaped fibres without superficial porosity.

To investigate the morphological properties of the PS fibres three imaging

techniques were chosen, thus defining the level of information that each tech-

nique can provide. Bright microscopy images constituted a valid tool to

visualise mesh density, defects (beads, polymer agglomerates and pores) and

fibre organisation (non-woven or aligned), while a more detailed analysis of

the morphology of the fibre surface was provided by SEM and AFM images.

SEM images offered the best magnification and resolution to measure the

fibre diameter and consequently calculate the average diameter of the fibres

forming a mesh. A better understanding of defect types and distribution was

also achieved using SEM.

SEM images were used to compare the average diameter and defect distribu-

tion in meshes electrospun under different process parameters.

The concentration of the polymer solution was found to be the key param-

eter affecting fibre diameter, which progressively increased with increasing

97 / 223


the concentration. As Huang Z.-M et al. [1] reported in their review, this

result is supported by various experimental studies available in the literature,

showing that solution concentration is one of the main parameters affecting

fibre diameter. Higher values of polymer concentration were shown to pro-

duce larger fibres according to a power law relationship [1, 152].

In the present study, the polymer concentration was also found to be a signif-

icant parameter affecting the formation of polymer agglomerates and beads

along the fibres. The number and size of beads decreased with the increase of

PS concentration. This result is confirmed by other studies found in the lit-

erature [1, 153, 154]. Electrospinning of PS nanofibres has been reported by

various authors [1, 155, 156], who showed the presence of beads or necklace-

like fibres often present within the meshes; these defects adversely affected

the reproducibility of the electrospinning system and the evenness of the re-

sultant fabrics. The formation of beads along electrospun fibres is due to

instability of the polymer jet and has been shown to depend on three key

factors: the viscosity of the solution which is proportional to the polymer

concentration; the charge density carried by the jet; and the surface tension

of the solution [157]. Eda et al. [154] showed the morphology of electrospun

PS fibres being significantly dependent on polymer molecular weight, con-

centration, and solvent. At a certain molecular weight (393,400 g/mol), as

the PS concentration in tetrahydrofuran (THF) was increased, the morphol-

ogy of the fibres progressively transitioned from beads only, to beads with

incipient fibres, elongated beads along the fibres, and bead-free fibres at the

highest concentration (21% w/v) [154].

The polymer concentration was also found to affect the spinnability of the

solution, in terms of fibre formation and deposition [9]. If the concentration

was not sufficient, the solution broke up into droplets before reaching the

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collector; if the solution was too concentrated the fibres could not form due

to excessive viscosity, which resulted in the clogging of the needle [9]. The

15%, 20%, 30% and 35% w/v concentrations of PS in DMF were all found to

allow the spinnability of the solutions; the 10% w/v solution could be elec-

trospun only at low voltage (10-15 kV) and droplet breakage often occurred,

while the 40% w/v concentration was too high and caused needle clogging.

For each concentration a specific set of optimal electrospinning parameters

(applied voltage, flow rate and N-C distance) for the formation of the fibres

was identified.

Voltage values below 10 kV resulted in most cases in needle clogging since

the applied electric field was not sufficient to induce the continuous forma-

tion of the polymer filament and this resulted in rapid solvent evaporation

in the needle. High voltage values, over 20 kV caused electrospraying, which

consists in the breakage of the polymer droplet at the tip of the needle, re-

sulting with the deposition of smaller droplets on the surface of the collector

[9, 152].

The applied voltage and N-C distance were found to not significantly affect

the fibre diameter. In the literature, the N-C distance is reported to play a

smaller role in the resulting morphological properties of the fibres, as long as

the collector is far enough from the spinneret to allow the complete evapora-

tion of the solvent. The applied voltage is known to control fibre formation

and size [45]. The fact that in this work the applied voltage was found to

induce no significant change in the fibre size could be due to the fact that the

tested voltage values (V = 15 kV and V = 20 kV) were not different enough.

However values of voltage significantly lower than 15 kV or higher than 20

kV were found to impair the spinnability of the solutions. An important

aspect to keep in consideration consists in the fact that although general

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relationships between electrospinning parameters and fibre morphology have

been drawn in the literature, the exact relationship will always be affected

by the chosen polymer/solvent system and the electrospinning setup in use

[9].

3.2 Electrospinning of nanofibres

The studies on the influence of electrospinning process parameters and poly-

mer concentration previously reported showed the possibility of fabricating

PS fibres in the nanometre scale by dissolving PS in DMF at a concentration

of 10% w/v (Φ = 300 ± 200 nm). However, as shown in Figure 3.5a, this

concentration resulted in the formation of defects along the fibres in form of

beads and polymer agglomerates that highly compromised the morphological

properties of the mesh.

A strategy to reduce the fibre diameter while preventing formation of defects

consists of increasing the conductivity of solutions at higher concentrations

through the addition of surfactants to the polymer solution [158]. Several au-

thors have successfully fabricated nanofibres by adding bromide salts, includ-

ing trabutylammonium bromide and hexadecyltrimethylammonium bromide

(HTAB), to the polymer solution to be electrospun [159, 160]. In the present

work HTAB was added to the 20% w/v polymer solution. Electrospinning

was performed by setting the voltage at 18-20 kV, flow rate 700-1000 µl/h

and needle-collector distance 20 cm. A blunt 24 Gauge needle was used.

The salt was added to the polymer solution in different concentrations, as

reported in Table 3.2. Resulting solution conductivity and average fibre di-

ameter are also shown in Table 3.2.

The addition of the HTAB surfactant to the the 20% w/v PS solution did

100 / 223


HTAB Conductivity Φ(mM) (µS/cm) (µm)

3 62 1000±2005 70 900±2008 93 800±10014 120 1100±10027 160 1200±600

Table 3.2: Solution parameters and average fibre diameter of electrospunpolystyrene solutions containing hexadecyltrimethylammonium bromide.

not lead to a significant reduction of the average fibre diameter. In fact, the

electrospinning of solutions containing 3 mM and 5 mM HTAB produced

meshes with the same Φ as the ones fabricated from pure PS (Φ = 1000 ±

100 nm); the 8 mM solution resulted in a small decrease of Φ to 800±100

µm, maintaining the mesh defect-free. Higher surfacatant concentrations (14

mM and 27 mM) induced an increase of Φ to 1100±100 µm and 1200±600

µm respectively. The high conductivity of these solutions (120-160 µS/cm)

resulted in unstable processability, with electrospraying often occurring.

Since the HTAB surfacant at different concentrations did not allow to fab-

ricate nanofibres with diameter smaller than 800±100 nm, two other surfac-

tants, cetyltrimethylammonium bromide (CTAB) and sodium dodecyl sulfate

(SDS), were tested [161, 162]. The surfactants were added to the 15% and

20% w/v PS solutions at a concentration of 0.1% w/v. The solutions were

electrospun by selecting the following process parameters: V = 14 kV; Flow

rate: 500 µl/h; N-C distance = 15 cm, which allowed stable spinnability.

A blunt 24 Gauge needle was used. The addition of the surfactants to the

PS solutions resulted in a significant increase of solution conductivity. The

conductivity of the 15% and 20% w/v PS solutions was measured to be 0.5

and 0.3 µS/cm respectively. After the addition of CTAB and SDS to the

101 / 223


15% w/v PS solution, the solution conductivity increased to 105 µS/cm and

70 µS/cm respectively; the average fibre diameter decreased to 400±80 nm

after the addition of CTAB and 450±90 nm after the addition of SDS. Con-

ductivity values of 92 µS/cm and 57 µS/cm were recorded after the addition

of CTAB and SDS respectively to the 20% w/v solution. The average fibre

diameter decreased to 700±100 nm after the addition of CTAB and 400±100

nm after the addition of SDS. A summary of the solution conductivity and

average fibre diameter values is reported in Table 3.3.

PS Salt Conductivity Φ(% w/v) (µS/cm) (nm)

15 - 0.5 900±20015 CTAB 105 400±8015 SDS 70 450±9020 - 0.3 1800±20020 CTAB 92 700±10020 SDS 57 400±100

Table 3.3: Solution conductivity and average fibre diameter obtained after theaddition of CTAB and SDS surfactants to 15% and 20% w/v PS solutions in

DMF.

Figure 3.8 shows the significant decrease in average fibre diameter induced

by the addition of the surfactants in both the 15 and 20% w/v PS solutions.

The smallest diameter was obtained with the 15% w/v solution containing

CTAB (Φ = 400±80 nm) followed by the solutions containing the SDS sur-

factant, that also resulted in average fibre diameter of 400 nm. The addition

of CTAB to the 20% PS solution also induced a decrease in fibre diameter

from 1800±200 to 700±100 nm, but this result was less significant compared

to the values obtained with the other solutions tested.

All the meshes were composed of uniform smooth round fibres, with no beads

or polymer agglomerates (Figure 3.9). The addition of CTAB to the 20%

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Figure 3.8: Average fibre diameter of the meshes electrospun from 15 and 20%w/v PS solutions in DMF before and after the addition of CTAB and SDS.

w/v PS solution (Figure 3.9c) induced the minor decrease of fibre diameter

compared to the addition of CTAB to the 15% w/v solution (Figure 3.9a)

and the addition of SDS to both the 15 and 20% w/v solutions (Figures 3.9b

and 3.9d).

Discussion To fabricate nanofibres while preventing the formation of de-

fects in the form of beads and polymer agglomerates within the mesh, the

conductivity of the 15% and 20% w/v PS solutions in DMF was increased.

Solution electrical conductivity is one of the major factors that affect the

diameter of electrospun fibres [163]. Uyar et al. carried out a systematic

study on the effect of solution conductivity on the electrospinning of PS fi-

bres and showed that even slight changes in the conductivity of the solution

can greatly affect the morphology of the resulting fibres [164].

In the present study, the increase in conductivity was achieved by adding

103 / 223


Figure 3.9: SEM micrographs of electrospun meshes obtained from: 15% w/vPS in DMF with the addition of (a) CTAB; (b) SDS; and 20% w/v PS in DMFwith the addition of (c) CTAB; (d) SDS. The diameter of the single fibres (in

red) is expressed in nm. Scale bar 1 µm.

104 / 223


ionic surfactants to the polymer solution. This approach has been shown to

be a good strategy to reduce the fibre diameter while preventing the forma-

tion of defects [158].

The addition of CTAB and SDS to the 15% and 20% w/v PS solutions

in DMF induced a significant increase of the conductivity and resulted in

smooth bead-free fibres with reduced average diameter. The reduction in

fibre diameter is due to the presence of the ionic surfactant that, having an

ionic hydrophilic head, induces an increase in surface tension and conduc-

tivity, which in turns increases the net charge density of the polymer jet

[158, 165]. This causes the jet being stretched under stronger force, resulting

in the exhaustion of any bead-like fluid; the larger charge repulsion in the

polymer jet due to the increased conductivity results in stretching the thread

thinner, thus obtaining smaller fibres [158].

When the HTAB surfactant was added to the 20% w/v PS solution an in-

crease of solution conductivity was measured but the fibre diameter did not

significantly change. This could be due to the incapability of the polymer

to associate with this specific surfactant and generate the so-called poly-

mer/surfactant interaction. The phenomenon of polymer/surfactant interac-

tion has been extensively studied by Lindman et al. [166] and Friberg [167]

and occurs when a non-ionic polymer associates with ionic surfactants by

wrapping the individual polymer chain around the surfactant molecules. Lin

et al. demonstrated that not all the ionic surfactants can be used to stop

bead formation and to tune fibre diameter, as only those that generate a

strong polymer/surfactant interaction are effective [158].

The addition of surfactants to polymer solution is a good strategy to con-

trol fibre morphology, however this approach needs to be carefully controlled

when the fibres are designed for medical applications, as surfactants are po-

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tentially toxic [168]. Fibre surface chemistry needs to be charatcerised prior

to any biological study, to investigate residual traces of surfactants that could

affect the responses biological molecules and cells.

3.3 Characterisation of electrospinning appa-

ratus performance

Reproducibility The reproducibility of the morphological properties of

the fibres fabricated through the electrospinning process was investigated.

The 15% w/v PS solution in DMF was electrospun selecting two values of

the applied voltage (15 and 20 kV). A flow rate of 800 µl/h was selected and

N-C distance was set at 20 cm. A month later the same experiment was per-

formed by using freshly prepared PS solution and selecting the same process

parameters. The conductivity of the solutions was measured to be 0.3-0.5

µS/cm. During the two sets of experiments the temperature was between 20

and 23◦C and the ambient humidity was measured at 40%. The average fibre

diameter of the meshes fabricated during the experiments is reported in Fig-

ure 3.10. The values of average fibre diameter were found to be reproducible,

ranging between 800 and 1200 nm, regardless the time when the electrospin-

ning was performed. As previosuly reported (section 3.1), the voltage was

found not to affect the resulting average fibre diameter both times. Fibre

morphological properties were reproducible in time, resulting round-shaped,

uniformly distributed within the mesh and defect-free, regardless when the

electrospinning was performed.

Spinning rate To calculate the spinning rate of the electrospinning appa-

ratus, fibres were fabricated from a 20% w/v PS solution in DMF containing

106 / 223


Figure 3.10: Average fibre diameter of the meshes electrospun from 15% w/vPS solutions in DMF at a time distance of one month.

1% w/v SDS. The process parameters that were selected are reported in sec-

tion 3.2. The electrospinning was performed for 30 seconds; 2; 5; 10; 30;

120; 240; and 360 minutes. After electrospinning, meshes were weighed on

a precision balance (XS603S, Mettler Toledo, Australia). The values of the

weight of the meshes measured after each spinning time is reported in Fig-

ure 3.11. Mesh weight was found to be constant (1.2 g) until 300 seconds

of electrospinning; at that time the weight began to progressively increase

reaching 1.8 g after 600 seconds. The fact that mesh weight does not vary

during the first 300 seconds of electrospinning could be due to an insufficient

sensitivity of the scale to detect minor changes of mesh weight. Moreover,

once the electrospinning is initiated, the fibres are initially deposited on a

spread area over the collector. Fibre deposition progressively narrows down,

covering a smaller, central portion of the collector, and the mesh starts to

form.

The morphological properties of the meshes were investigated by comparing

107 / 223


Figure 3.11: Graph showing the weight of the meshes after different times ofelectrospinning

SEM images of fibres fabricated from 30 seconds up to 6 hours of electro-

spinning. Average fibre diameter was found to be constant (400 ± 100 nm)

from 30 to 4 hours. After this time threshold fibre diameter slightly increased

reaching 500 ± 100 nm and after 6 hours it was measured 600 ± 100 nm.

Discussion The homebuilt electrospinning apparatus was characterised by

measuring the spinning rate and analysisng the reproducibility of the fibre

morphological properties. The system was found to be reproducible when

looking at the average diameter of fibres electrospun in the same conditions

at a time distance of one month.

The average fibre diameter was found to change when the PS solution was

electrospun continuously for longer than 4 hours. After this time threshold,

fibre diameter increased with the increase of spinning time. This could be

caused by a progressive evaporation of the solvent of the solution in the spin-

neret, that results in a slow increase of solution concentration, that in turns

108 / 223


induces the formation of larger fibres. The characterisation of the electro-

spinning apparatus was a necessary step in this work due to the high number

of variables that are involved in the process. Although there are multiple

studies in the literature investigating the electrospinning phenomenon [169–

171], defining process parameters and solution properties prior fabrication

depending on the desired properties of the meshes, remains a challenge [172].

It is not always possible to exactly predict the features of the electrospun

meshes on the basis of the selected parameters. The current state of the art

consists in finding the best range for each electrospinning parameter for a

given polymer/solvent system and for the apparatus in use. The parameters

have to be continuously adjusted during the process to achieve uniform and

continuous spinnability. The reproducibility study was performed to ensure

that the optimal values of the parameters initially found for the electrospin-

ning of the PS/DMF system were reproducibly providing fibres with specific

morphological characteristics.

3.4 Electrospinning of aligned fibres

The home-built rotating mandrel (see chapter 2, section 2.1.1 for details on

the collector) was used to collect PS fibres along a preferential direction. The

electrospinning of the 20% w/v polymer solution was performed at a voltage

of 18 kV, flow rate of 800 µl/h and N-C distance of 20 cm. Two rotational

speeds of the rotating mandrel were tested: 500 rpm and 2500 rpm. Fibre

diameter and diameter distribution resulted not to be affected by the type of

collector used. In fact, the meshes electrospun on the rotating mandrel had

an average fibre diameter of 1600±200 µm, regardless the selected rotational

speed, which matches the average fibre diameter measured on the meshes

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Figure 3.12: SEM images of PS meshes electrospun on the rotating mandrel attwo rotational speeds: (a) 500 rpm; (b) 2500 rpm. Scale bar 10 µm.

electrospun on the metal plate from the 20% w/v PS solution.

The presence of beads and defects within the mesh was found to be affected

by the rotational speed of the mandrel. In fact, when using the lowest speed,

the mesh resulted to be defect-free, as shown in Figure 3.12a. When the

rotational speed was increased to 2500 rpm some beads and polymer ag-

glomerates formed along the fibres (Figure 3.12b).

The rotational speed of the cylinder was found to be a significant parameter

affecting the alignment of the fibres. With the low rotational speed (500

rpm), no preferential fibre alignment was found to occur in the mesh; when

the rotational speed was increased to 2500 rpm a visible degree of alignment

of the fibres oriented along the rotational direction of the cylinder occurred.

Discussion The rotating mandrel was designed and fabricated to provide

the electrospinning setup for an additional tool, that can be used for a variety

of tissue engineering applications that require the use of meshes or scaffolds

composed of aligned fibres (vascular and cardiac grafts [83], nerve regenera-

tion [83, 173], skeletal muscle regeneration [174]).

A minimum value of the mandrel rotational speed of 2500 rpm was found

110 / 223


necessary to achieve the alignement of the fibres along the rotational direc-

tion of the cylinder. Lower values of the rotational speed resulted in the

deposition of non-woven meshes.

The dependence of the degree of fibre alignment on the mandrel rotational

speed has been reported by Teo et al., who reviewed the different designs of

collectors that can be used for the electrospinning of non-woven and aligned

meshes [175]. Matthews et al. demonstrated the effect of the mandrel rota-

tional speed on the alignment degree of collagen fibres. Below 500 rpm, a

random non-woven mix of fibres was collected; to achieve a visible alignment

the rotational speed had to be increased at 4500 rpm [176].

A specific rotational speed of the collector is necessary to align the fibres

due to the high travel speed of the polymer jet from the needle towards the

collector [177]. The mandrel rational speed has to be sufficiently high so that

the fibres can reach the mandrel surface and be wounded around it [177].

3.5 Conclusions

The present chapter focuses on the characterisation of the home-built elec-

trospinning apparatus and the optimisation of the process parameters for the

fabrication of polystyrene fibres with controlled morphological properties.

The solvent used to prepare the PS solution was found to significantly af-

fect the spinnability and the surface properties of the fibres. DMF was the

optimal solvent for producing smooth round fibres with no porosity on the

surface.

Multiple combinations of process parameters (applied voltage, flow rate, and

needle-collector distance) were tested for a variety of polymer concentrations.

An ideal set of process parameters to fabricate fibres with controlled diameter

111 / 223


and surface morphology was defined where the fibres can then be confidently

used in subsequent experiments throughout this thesis. The 10% w/v PS

solution was electrospun only at low voltage (10-15 kV) and nanofibre (Φ =

300 ± 200 nm) meshes affected by the presence of beads and polymer agglom-

erates were obtained. Higher concentrations (15%, 20%, 30%, and 35% w/v)

resulted in fibres with larger diameters and fewer and progressively smaller

defects. The highest concentration tested (40% w/v) was not electrospun as

needle clogging always occurred due to the high viscosity of the solution.

The addition of CTAB and SDS surfactants to the 20% w/v PS solution was

proven to be a good strategy to increase the conductivity of the solution for

fabricating nanofibres while preventing the formation of defects within the

mesh.

to ensure the spinnability of PS solutions, the applied voltage was tuned at

15-20 kV. In fact, voltage values below 10 kV resulted in most cases in needle

clogging as no filament formed from the polymer droplet; voltage values over

20 kV caused the breakage of the polymer droplet and electrospraying.

The home-build electrospinning apparatus was proven to produce meshes

with reproducible average fibre diameter and surface morphology when the

process was continued for less than 4 hours. After this time, fibre diameter

was measured to increase with the increase of spinning time, possibly due to

the evaporation of the solvent in the spinneret.

112 / 223

Chapter 4

Plasma polymerisation of

electrospun meshes

Contents4.1 Characterisation of plasma polymerised meshes 114

4.1.1 Surface morphology of plasma polymerised meshes 114

4.1.2 Surface chemistry of plasma polymerised meshes . 116

4.1.3 Aging study on ppAAm coating . . . . . . . . . . 122

4.2 Conclusion . . . . . . . . . . . . . . . . . . . . . . 124

To investigate the response of bacteria to different fibre surface chemistries,

surface modification was carried out using plasma polymerisation. The present

chapter aims to characterise the surface chemistry and morphology of the

plasma modified fibre meshes. Plasma polymerised films were deposed us-

ing four different monomers: acrylic acid (ppAAc); 1,7-octadiene (ppOct);

allylamine (ppAAm); and 1,8-cineole (ppCo). These monomers were chosen

because they generate different chemical functionalities with various degrees

of wettability and surface charge [107, 178]. In addition, air plasma treatment

was performed to increase the hydrophilicity of the fibre surfaces [179]. To

113

4. Plasma polymerisation of electrospun meshes

investigate changes in fibre morphology induced by the surface modification

process SEM images were acquired. XPS analysis was performed to anal-

yse the chemical composition of the fibre surfaces and verify the successful

deposition of the coatings.

4.1 Characterisation of plasma polymerised

meshes

4.1.1 Surface morphology of plasma polymerised

meshes

Air plasma treatment was performed on the PS meshes to increase the wet-

tability of the fibre surface [179]. To investigate any changes in fibre surface

morphology induced by the treatment SEM images were acquired. Figure 4.1

shows a comparison between the untreated PS, air plasma treated and plasma

polymerised fibres. The air plasma treatment caused significant etching of

the fibres. After 5 minute treatment time the fibre surface had an increased

roughness with spread out porosity on the individual fibres (Figure 4.1b)

compared to the untreated mesh (Figure 4.1a) that is composed of smoother

fibres with no pores. The plasma polymerisation processes did not induce

significant changes in fibre surface morphology. The ppAAc, ppCo, ppOct

and ppAAm plasma coated meshes shown in Figures 4.1c, 4.1d, 4.1e, and 4.1f

respectively are composed of uniform fibres with surface morphology com-

parable to the untreated sample (Figure 4.1a), indicating that no significant

etching occurred during the coating processes. The average fibre diameter

remained unaltered (500 ± 200 nm) after the plasma polymerisation of the

monomers, further confirming that no polymer material was etched from the

114 / 223


Figure 4.1: SEM microgaphs of (a) untreated; (b) air plasma treated; (c)ppAAc; (d) ppCo; (e) ppOct; and (f) ppAAm plasma coated PS fibres. Scale bar

1 µm

fibre surface.

Discussion A significant increase in surface roughness and a spread out

porosity was found to occur after the air plasma treatment. The change in

the surface morphology is due to the substrate etching associated with the

plasma process itself and is believed to result mainly from the bombardment

of the surface by the energetic particles present in the plasma [180]. Cui

et al. modified the surface properties of polypropylene (PP) films using air

plasma and AFM analysis showed an increased roughness and the formation

of annular features on the surface after 2 minute treatment [180]. Yang et

al. used low-pressure air plasma to modify surface properties of polyethy-

lene terephthalate films and obtained a significant decrease of contact angle

accompanied by the formation of conical protuberances on the surface [181].

The roughness increase induced by the air plasma is due to the removal of

top few monolayers of the substrate surface, caused by the impact of plasma

species on the surface [181].

115 / 223


Due to the significant etching of the surface, the air plasma treated meshes

were not further characterised and excluded from the present study to pre-

vent additional variables to interfere with the interactions of bacterial cells

with the fibre surface. The change in fibre surface wettability was achieved

through the plasma polymerisation of the acrylic acid and allylamine monomers,

that generate hydrophilic films, and the 1,7-cotadiene and 1,8-cineole monomers

that increases the hydrophobicity of the fibres [22, 182].

4.1.2 Surface chemistry of plasma polymerised meshes

To confirm the successful deposition of the plasma coatings onto the PS

meshes, XPS survey and high resolution C1s analysis were performed. Fig-

ure 4.3 and Figure 4.2 show respectively the wide scans and the C1s high

resolution spectra obtained for each coating as well as the untreated mesh.

The elemental composition and atomic ratios of the analysed meshes are

compared with the theoretical values derived from the molecular formula of

the monomers in Table 4.1.

On the wide scan spectrum (Figure 4.3), the elemental analysis of the PS

mesh showed the surface composition to be 99-100% carbon, nearly match-

ing the theoretical values derived from the molecular formula of PS (Table

4.1). Trace amounts of oxygen and nitrogen (0.2 and 0.1% respectively) were

detected on the fibre surface, due to the exposure of the fibres to impurities

during the electrospinning process.

The C1s spectrum of the PS mesh was resolved into two components (Fig-

ure 4.4a): an intense peak at 285.0 eV characteristic of hydrocarbon groups

(C-C/CH and C=C) and small peaks shift at about 291 eV attributed to a

shake-up satellite peak from the aromatic π–π* transition .

116 / 223


Figure 4.2: XPS high-resolution carbon 1s spectra of untreated and plasmapolymerised PS meshes

117 / 223


Sam

ple

Theo

reti

cal

atom

icco

mp

osit

ion

(%)

Theo

reti

cal

atom

icra

tio

(%)

(Mon

omer

form

ula

)C

1sO

1sN

1sT

race

ofO

/CN

/C

PS

(C8H

8)

100

00

-0

0

ppA

Ac

(C3H

4O

2)

6040

0-

0.66

0

ppC

o(C

10H

18O

)91

90

-0.

090

ppO

ct(C

8H

14)

100

00

-0

0

ppA

Am

(C3H

5N

H2)

600

40-

00.

66

Sam

ple

Mea

sure

dat

omic

com

pos

itio

n(%

)M

easu

red

atom

icra

tio

(%)

C1s

O1s

N1s

Tra

ceof

O/C

N/C

PS

99.7±

0.2

0.2±

0.2

-N

0.1

00

ppA

Ac

75.6±

0.1

24.4±

0.1

--

0.32

0

ppC

o95

.1±

0.1

4.9±

0.1

--

0.05

0

ppO

ct97

.7±

02.

3±

0-

-0.

020

ppA

Am

84.2±

0.1

3.2±

012

.6±

0.1

-0.

040.

15

Tab

le4.1

:X

PS

theo

reti

cal

an

dm

easu

red

ato

mic

com

posi

tion

an

dato

mic

rati

os

rela

tive

toth

eto

tal

con

cen

trati

on

of

carb

on

(O/C

an

dN

/C

)of

the

un

coate

dan

dpla

sma

coate

dm

eshes

.T

he

mea

sure

dva

lues

are

the

mea

nva

lues±

dev

iati

on

base

don

+3

an

aly

ses

perf

orm

edon

each

sam

ple

.

118 / 223


Figure 4.3: XPS wide scan spectra of the uncoated PS mesh and the plasmapolymerised meshes

The high resolution C1s spectra of all the plasma polymerised meshes

(Figure 4.2) showed the disappearance of the shake-up satellite peak compo-

nent characteristic of PS, indicating that the meshes were uniformly coated.

The wide scan spectrum (Figure 4.3) and elemental composition analysis of

ppAAc indicated that the films were rich in carbon and oxygen. The atomic

composition showed a considerable retention of oxygen within the films (24.4

± 0.1%) and a O/C ratio of 0.32. As shown in Table 4.1, the elemental

composition of the ppAAc did not exactly reproduce the theoretical values

that were expected based on the the molecular formula of the acrylic acid

monomer (O/C ratio of 0.66%). The curve fitted C1s spectrum of the ppAAc

coated mesh is shown in Figure 4.4b. Five component peaks were used to

fit the spectrum, including 55.1 ± 1.1% hydrocarbon group (C-C, C-H) at

285.0 eV; 14.1 ± 1.4% hydroxyl component (C-O) at 286.6 eV; and 3.8 ± 1.0

% carbonyl (C=O, O-C-O) at 287.9 eV. A distinctive binding energy shift

119 / 223


Figure 4.4: XPS high-resolution carbon 1s spectra of (a) untreated PS and (b)ppAAc coated meshes with fitted curves

to approximately 289 eV was indicative of the C-C*-O=O component of the

ppAAc film. The C*-C-O=O component was found to be 13.5 ± 0.4 %. The

associated β shift (C*-C-O=O) was placed at 285.6 eV, in line with literature

standards [183].

The XPS survey for the ppCo coating revealed a film rich in carbon (95.1 ±

0.1 %) with an amount of oxygen of 4.9 ± 0.1 %, which is lower than the

theoretical oxygen content of 9% (Table 4.1). The spectrum of the ppOct

coated meshes showed similar results, with a oxygen content of 2.3 ± 0 %.

The ppAAm was composed of carbon (84.2 ± 0.1 %); oxygen (3.2 ± 0 %);

and nitrogen (12.6 ± 0.1 %). The amount of nitrogen in the ppAAm coating

was lower than the expected theoretical amount (40%); oxygen was detected

in both the ppOct and ppAAm coatings, although this element is not present

in the molecular structure of the original monomers (Table 4.1).

Discussion The XPS analysis performed on the meshes obtained from the

20% w/v PS solution in DMF containing 0.1% w/v SDS did not detect sulfur

on the surface of the meshes, indicating that the addition of the surfactant

120 / 223


to the polymer solution did not affect the surface chemistry of the resultant

fibres. The XPS survey and C1s high resolution spectrum of the PS meshes

showed the fibre surface being uncontaminated, matching the XPS data on

PS films or scaffolds available in the literature [148, 184, 185]. Trace amounts

of nitrogen and oxygen were detected on the fibre surface; the presence of

these elements is possibly due to impurities deriving from the polymer or the

electrospinning apparatus [148]. In addition, the DMF solvent used to pre-

pare the PS solution might not completely evaporate during the fabrication

process resulting with impurities trapped on the fibre surface.

The XPS analysis performed on the plasma coatings on the fibre surface

matched the results previously reported in the literature for flat uniform

coatings [22, 186].

All the high resolution C1s spectra (Figure 4.2) showed the disappearance of

the shake-up satellite peak characteristic of the untreated PS, indicating that

the films were uniformly covering the fibres and the thickness of the coatings

was higher than the XPS analysis depth (about 10 nm at the selected take-off

angle of 90◦).

The XPS survey on the ppAAc coated meshes showed about 14 % of car-

boxyl component, that correlates with the literature and confirms that the

desired chemistry was achieved, being the carboxyl component the finger

print of ppAAc [187, 188]. The significant amount of nitrogen (12.6 %) in

the ppAAm films was expected as it is indicative of presence of amines and

amides [189, 190] and the calculated N/C ratio (0.15) correlates with values

reported in the literature [135, 191]. The elemental composition of the ppOct

and ppCo coating matched the XPS analysis reported by Pegalajar-Jurado

et al. [22].

The elemental composition of the plasma polymerised films did not entirely

121 / 223


reproduce the values that were expected from the molecular formula of the

original monomers (Table 4.1). The O/C ratio for the ppAAc and ppCo films

as well as the N/C ratio for the ppAAm coating were lower than expected.

The loss in chemical functionality of the plasma polymer films is known to

be due to the complexity of the chemistry in the gas phase and to the frag-

mentation of the monomers and some loss of functionality that occurs during

the polymerisation process [192, 193]. The retention of chemical function-

ality within plasma polymers has been shown to be dependent on several

factors, including reactor design and deposition parameters, such flow rate

and treatment duration [194].

Although the 1,7-octadiene monomer does not contain oxygen, this element

was detected in the ppOct plasma polymer. The presence of oxygen in the

coating is due to the presence of free radicals in the film that induce oxidation

upon exposure to the atmosphere. These results correlate with the existing

literature on the plasma polymerisation of 1,7-octadiene[135, 191].

Similarly, the oxygen content found in the elemental composition of the

ppAAm coating is due to the post-plasma oxidation associated with amine

containing plasma polymers [191].

4.1.3 Aging study on ppAAm coating

Plasma polymers are known to undergo a range of oxidation and ageing

processes [191, 195]. This is particularly important for ppAAm as the films

are known to incorporate oxygen and form amides over time [191]. The

oxidation rate of the ppAAm films was investigated by analysing the surface

chemistry of the coated meshes after different days from the time when the

plasma coating was generated. The XPS survey was undertaken immediately

after the plasma polymerisation process and subsequently after 1, 2, 5, 7, 14,

122 / 223


and 22 days. The oxygen, nitrogen, and carbon content of the coating that

was recorded each day is shown in Figure 4.5a. The carbon content was

found to slightly decrease in the tested time frame, from 83.9 ± 0.2 % at day

0 to 81.3 ± 0.3 % at day 22; while the nitrogen content was constant until

the 14th day (about 13%) and then decreased to 12%. A more significant

change was obtained in the content of oxygen, which was 2.1 ± 0.1 % the

day of the coating and then progressively increased with a linear trend to 3.3

± 0.5 % on day 1; 3.8 ± 0.1 % on day 2; 4.2 ± 0.2 % on day 5; 5.2 ± 0.3 %

on day 7; and 5.6 ± 0.7 % and 6.7 ± 0.2 % on days 14 and 22, respectively.

This increase is shown in Figure 4.5b, where the variation of the O/C and

N/C atomic ratios in the considered time frame is reported. The O/C ratio

increased progressively from 0.03 at day 0 to 0.08 at day 22, while the N/C

ratio slightly varied in the range 0.15-0.17.

Figure 4.5: (a) Elemental composition and (b) oxygen/carbon andnitrogen/carbon atomic ratios of the ppAAm caoted meshes from day 0 until 22

days after plasma polymerisation

Discussion The intrinsic reactivity of the amine groups present in the

ppAAm films induced changes in the elemental composition of the coating

123 / 223


over time. The surface oxidation was found to be progressively more signifi-

cant, with about 5% oxygen content increase 22 days after plasma polymeri-

sation. Several studies in the literature have shown that plasma polymer

films undergo aging upon storage [191, 195, 196]. Whittle et al. investigated

the changes with sample age in the surface chemistry of various plasma poly-

merised films, including ppAAm. Authors showed a sharp uptake of oxygen

in the ppAAm coatings during the 30 days following the deposition [191].

The mechanism by which the oxygen is incorporated over time in the plasma

polymer films is driven by the reaction of oxygen with long-lived reactive

species present within the surface. These species that are trapped in the

films can react with oxygen upon exposure to air, resulting in oxygen incor-

poration into the film. The most likely form of these reactive species are

radicals trapped in the plasma polymer during the treatment [195].

4.2 Conclusion

The present chapter focuses on the morphological and chemical characteri-

sation of the air plasma treated and the plasma coated meshes.

The air plasma treatment resulted in a significant etching of the PS fibres,

compromising the fibre surface morphology. On the contrary, the deposition

of the plasma polymers did not induce significant changes in fibre surface

roughness and average fibre diameter.

The XPS analysis of the plasma modified meshes confirmed the successful

deposition of the coatings uniformly on the fibre surface, matching studies

in the literature based on the plasma polymerisation of the same monomers

on flat substrates.

The ppOct and ppAAm coatings were found to incorporate oxygen upon

124 / 223


exposure to air. Due to the high reactivity of the amine groups present in

the ppAAm films, the surface oxidation upon aging was characterised, show-

ing an increase of 5% in oxygen incorporation over 22 days. This change in

surface chemical composition needs to be considered when the responses of

biological molecules or cells to ppAAm meshes are investigated.

The results reported in the present chapter demonstrate that the plasma

polymerisation of different monomers allowed to control and tune the chem-

istry properties, including surface charge, wettability and functional groups,

of the PS fibres. These properties have been previously shown to affect and

potentially drive bacterial attachment and proliferation on flat surfaces [85].

The need now exists to extend these studies to fibrous three-dimensional

substrates. The following chapter report the results obtained investigating

the response of E.coli cells to the plasma polymerised PS meshes.

125 / 223


126 / 223

Chapter 5

Interactions of wound bacteria

with electrospun meshes

Contents5.1 Bacterial colonisation of electrospun meshes . . 129

5.2 Influence of fibre diameter on bacterial behaviour133

5.3 Influence of fibre surface chemistry on bacterialbehaviour . . . . . . . . . . . . . . . . . . . . . . . 146

5.3.1 Bacterial transfer onto ppAAm coated meshes . . 154

5.4 Conclusions . . . . . . . . . . . . . . . . . . . . . . 156

The mechanisms of interactions of wound bacteria with fibrous substrates

with different morphological and surface chemistry properties is still an open

question in the literature and needs to be addressed to develop effective

strategies for controlling the bacterial load in the wound bed.

In the present chapter the response of bacteria commonly involved in chronic

wound infections to electrospun PS meshes with different morphological and

chemical properties was investigated. The experiments were designed to

understand the interactions occurring between bacteria and electrospun fibres

127

5. Interactions of wound bacteria with electrospun meshes

in the short term, during the first hour of contact, since initial bacterial

attachment constitutes the critical initial step towards biofilm formation and

infection development:

• In section 1 the colonisation of PS electrospun meshes by E.coli from

a suspension culture was assessed using SEM images and the MTT

viability assay.

• Section 2 explored the influence of fibre size on the capacity of wound

bacteria (E.coli, S.aureus, and P.aeruginosa) to transfer, attach and

colonise PS meshes. Experiments included attachment studies in liquid

medium but also directly onto agar plates; the latter was aimed at

mimicking a chronic wound environment.

• In section 3 plasma modified meshes (ppAAc, ppCo, ppOct, ppAAm),

exposing different surface wettability and functional groups, were ex-

posed to E.coli agar cultures and the transfer mechanisms of the bac-

teria onto and within the meshes was assessed.

The work presented in this chapter has been published in the journal articles:

M. Abrigo, P. Kingshott, S. L. McArthur, ”Electrospun Polystyrene Fiber

Diameter Influencing Bacterial Attachment, Proliferation and Growth,” ACS

Appl. Mater. Interfaces., vol. 7, no. 14, pp. 7644-52, 2015

M. Abrigo, P. Kingshott, S. L. McArthur, ”Bacterial response to differ-

ent surface chemistries fabricated by plasma polymerization on electrospun

nanofibers,” Biointerphases, vol. 10, no. 4 pp. 04A301-9, 2015

128 / 223








5.1 Bacterial colonisation of electrospun meshes

In the initial studies, MTT and MTS assays were performed to assess the

attachment and viability of E.coli cells as they interact with electrospun

meshes.

15% w/v PS solution in DMF (Φ=900±200 nm) was electrospun into meshes

and exposed to an E.coli culture. Samples were then stained using MTT to

determine the cell viability; one mesh was not exposed to the bacterial cul-

ture before staining. Figure 5.1 shows the photographs of the meshes after

the staining. The MTT solution is yellow in colour, due to the tertrazolium

compound. Originally electrospun meshes were white in colour, but after the

staining and multiple washing steps, the control mesh (Figure 5.1a) appeared

yellow, indicating that the MTT solution penetrated through the fibrous sub-

strates and adsorbed to the surface of the PS fibres. The mesh exposed to the

bacteria (Figure 5.1b) appeared purple after contact with the MTT solution,

indicating that viable bacterial cells were present within the mesh.

Figure 5.1: Electrospun PS meshes stained through the MTT assay. (a) Controlmesh, not exposed to bacterial culture; (b) Mesh exposed to bacterial culture for 1

hour. Scale bar 1 cm

As the MTT assay produced a solid product, the solution form of the assay

(MTS) was performed to provide quantitative information on the number of

129 / 223


viable cells attached onto and within the meshes after they were exposed to

the bacterial culture. No significant change in signal from the control mesh

compared to the background signal of the MTS solution occurred (0.20±0.02,

p<0.005). When the meshes were exposed to media containing bacteria, the

absorbance values rose significantly to 0.33±0.01, p<0.05.

To explore initial bacterial adhesion, proliferation and colonisation of the PS

electrospun meshes (Φ=900±200 nm), they were exposed to E.coli for 30

minutes, 1 hour, 2 hours, 4 hours and 6 hours. Adherent bacteria were fixed

and imaged using SEM. The progressive colonisation of the meshes is shown

in Figure 5.2.

Figure 5.2: E.coli cells onto electrospun PS fibres after incubation for (a) 30min; (b) 1hr; (c) 2hrs; (d) 4 hrs; and (e) 6 hrs. Scale bar: (a) and (b) 1 µm; (c),

(d) and (e) 2 µm.

After 30 minutes of incubation (Figure 5.2a) single bacterial cells adhered

onto the surface of individual fibres. After one hour, single cells were still

130 / 223


visible along fibres with small clusters of bacteria forming at points where

several fibres crossed (Figure 5.2b). Figure 5.2c and 5.2d show that after 2

and 4 hour exposure to the bacterial culture the initial clusters had rapidly

spread and bridged across fibres colonising a progressively larger area of the

mesh. After 6 hours (Figure 5.2e), colonies appeared to have spread estab-

lishing structure using the fibres as a supportive frame. The SEM images

obtained after different times of incubation suggest that in 30 minutes E.coli

cells were capable of irreversibly adhering onto the fibres and within 6 hours

they spread within the mesh forming progressively bigger colonies.

In Figure 5.3a a bacterial cell in the typical elongated configuration occurring

during microbial division is pointed by the red arrow; Figure 5.3b shows a

cell that completed or is completing the fission process, with the formation

of two separate cells ready to divide.

Figure 5.3: SEM of bacteria colonising PS electrospun mesh. (a) The elongatedbacterium pointed by the red arrow is in the elongation configuration occurring

during the binary fission process; (b) two bacterial cells after cell fission, ready todivide. Scale bar 1 µm.

131 / 223


Discussion MTT and MTS colourimetric assays are traditionally performed

on mammalian cell cultures for determining cell viability in proliferation, cy-

totoxicity, cell attachment, chemotaxis and apoptosis. In this study, the

assays were adapted to bacterial culture to visualise and assess the viability

of E. coli in electrospun PS meshes. These assays were found to be effec-

tive tools for this purpose, providing information on the viable bacterial cells

colonising the fibrous structure.

The MTT experiments qualitatively demonstrated the viability of E.coli cells

throughout electrospun meshes, while the MTS assay provided a more quan-

titative approach to study the viability of bacterial cells through the PS

meshes.

Results showed that the assays did not detect any viable cells in the control

mesh, which was expected since the mesh was not exposed to the bacterial

culture. On the contrary, the absorbance of the MTS solution added to the

mesh exposed to the bacterial culture resulted to be statistically significantly

higher than the absorbance obtained from the control, confirming the pres-

ence of viable cells within the mesh.

While MTT and MTS assays provide useful initial information regarding the

viability of the bacteria within the substrate, a better understanding of the

behaviour of the cells when exposed to a fibrous substrate was required. To

explore initial bacterial attachment and progressive spreading within electro-

spun meshes, SEM images were used. The colonisation of the meshes in time

resulted in the progressive formation of stable and compact aggregates of

bacteria using the fibres as a scaffold to support attachment and spreading.

On the SEM images of bacteria colonising the fibres it was possible to distin-

guish the typical configuration of the cells occurring during microbial division

through binary fission mechanisms [197] (Figure 5.3). The presence of the

132 / 223


fibres and the empty spaces within the mesh did not impair the capacity of

the bacterial cells to attach and divide. The mesh appeared to encourage the

cells to spread across the interstices and use the fibres as a support to create

bridges and form compact agglomerates.

Many authors in the tissue engineering literature have shown that fibrous

substrates can be used to encourage the adhesion, proliferation, and activa-

tion of mammalian cells for tissue reconstruction [198]; similarly, the present

results show that bacterial cells are able to use the fibres in similar ways. This

knowledge constitutes an essential achievements particularly for those fibrous

devices designed to interact with an environment intrinsically populated by

mammalian and bacterial cells, such as chronic wound beds.

5.2 Influence of fibre diameter on bacterial

behaviour

As the interaction of mammalian cells with fibrous substrates have been

shown to strictly depend on the size of the fibres [82], the obtained results

led to the next question regarding the influence of fibre diameter on the ca-

pability of bacteria to colonise the meshes.

The work that most closely approaches this problem in the literature was

provided by Kargar et al. [89], who investigated the state of adhesion of

P. aeruginosa bacteria to flat PS surfaces texturized with aligned PS fibres

with different diameters and spacing. The minimum value of bacterial ad-

hesion density was found to occur for fibres with a diameter close to the

bacterial diameter at a spacing less than bacterial diameter; the highest den-

sity was measured when the spacing between fibres and fibre diameter were

bigger than the bacterial size [89]. These results were obtained by studying

133 / 223


the single-cell level, thus bringing up the question about the influence of fi-

bre diameter on the capacity of bacteria to spread and colonise micro and

nanofibrous meshes.

E.coli attachment in solution To investigate the influence of fibre di-

ameter on the initial attachment and spreading of bacteria, meshes in three

average fibre diameter ranges were used: Φ1 = 500 ± 200 nm; Φ2 = 1000 ±

100 nm; and Φ3 = 3000 ± 1000 nm.

The Φ1 mesh was fabricated by electrospinning 20% w/v PS in DMF contain-

ing 0.1% w/v SDS. To ensure that the mesh did not leach residual surfactant

when exposed to a culture of bacteria, an inhibitory zone experiment was per-

formed. The mesh was incubated on confluent layers of E. coli cells cultured

on agar and incubated overnight at 37◦C. While the UrgoCell Ag/SilverR con-

trol sample (silver impregnated mesh obtained from Urgo Medaical, France)

produced a clear zone of inhibition, killing the cells it came in contact with

on the agar, no ring was detected around the Φ1 mesh.

The meshes were exposed to a culture of planktonic E.coli bacteria in growth

media with an O.D. of 0.3. Samples were immersed in the culture for 1 hour

at 37◦C. SEM images were used to monitor the spreading of bacteria within

the mesh; confocal images were taken after performing the LIVE/DEAD as-

say to visualise the distribution of live and dead cells within the samples.

The influence of fibre diameter on the ability of E. coli cells to spread within

the mesh and form colonies is shown in Figure 5.4. In the confocal images

(Figure 5.4a, 5.4c and 5.4e) fibres are fluorescing green due to the autoflu-

orescence of the PS. Bright green and red spots corresponding to live and

dead bacterial cells respectively can be visualised. When Φ = Φ1 (Figure

5.4a), a high prevalence of dead bacterial cells were present within the mesh.

134 / 223


Bacteria appeared to be mainly isolated cells adhering onto the surface of

the fibres. This was confirmed by SEM images (Figure 5.4b), where cells

were found to adhere onto and wrap around the surface of the fibres. Few

cells appeared to bridge from the surface of the fibres towards the interstices.

Clusters of bacteria formed within the mesh, but due to the small size of the

fibres, bacteria appeared to find it difficult to create compact colonies on the

fibrous network during the 1 hour exposure time. When the fibre diameter

was increased to Φ2, bacteria appeared to be able to bridge across fibres and

create colonies that used the fibrous substrate as a scaffold, supporting and

encouraging cell growth and spreading. Figure 5.4c shows a colony of E. coli

cells adhered over tens of fibres. A high proportion of live cells could be seen

on the surface of the fibres as well as throughout the colony in the interstices.

The SEM image (Figure 5.4d) confirmed the presence of a compact colony

spread throughout the mesh. The darker agglomerates on the surface of the

mesh corresponded with the extracellular polymeric substance (EPS) that

bacteria themselves produce to ensure adhesion onto a surface and to each

other. When the fibre diameter was greater than the bacterial size (Φ = Φ3),

E. coli cells appeared to consider each fibre surface as a flat substrate. Most

of the bacteria were aligned along single fibres in the form of a train (Figures

5.4e and 5.4f). In response to crossing over points among two or more fibres,

bacteria were able to proliferate across the fibres, producing agglomerates of

cells.

The response of isolated E. coli cells to single fibres in the three diame-

ter ranges is shown in Figure 5.5, where the cells are false coloured in red.

When Φ = 300 nm (Figure 5.5a), the cell appeared to wrap around the fi-

bre to achieve complete adhesion, thus assuming a round shape. When Φ

= 1 µm (Figure 5.5b) or bigger (Figure 5.5c), the cells could easily adhere

135 / 223


Figure 5.4: Bacterial solution culture experiment. Confocal (a; c; e) and SEM(b; d; f) images of E. coli cells colonising PS electrospun meshes with fibre

diameter ranges: (a, b) Φ1 = 500 ± 200 nm; (c; d) Φ2 = 1000 ± 100 nm; (e; f)Φ3 = 3000 ± 1000 nm. Scale bar: (a), (c), and (e) 5 µm; (b), (d), and (f) 2 µm.

136 / 223


Figure 5.5: SEM of single E. coli cells (false coloured in red) adhered onto PSelectrospun fibres with diameter: (a) 0.3 µm (b); 1 µm; (c) 5 µm. Scale bar 1

µm.

onto the surface maintaining their original rod-like shape. This behaviour

suggests that the distortion of E. coli cells required to adhere onto the Φ1

fibres affects bacterial function and viability, resulting in a prevalence of dead

isolated cells. The change of bacterial shape induced by the small size of the

fibres could impair the ability of bacteria to bridge across fibres and produce

EPS for developing colonies.

137 / 223


Agar-mesh cell transfer Colonised or infected wounds are solid sub-

strates contaminated by a variety of bacterial species. In an attempt to at

least in part mimic this environment, confluent biofilms of E. coli, P. aerug-

inosa and S. aureus were grown on agar plates. The meshes with the three

fibre diameter ranges were placed on top of the bacterial cultures for 1 hour

at 37◦C. The capacity of the meshes to attract and remove the bacterial cells

from the agar plates was tested by staining the plates with crystal violet

(CV) after mesh removal. Figure 5.6 shows (a) E. coli, (b) P. aeruginosa

and (c) S. aureus agar cultures where the mesh was in contact for 1 hour

and then removed. The images clearly show that the meshes attracted and

removed most of the bacteria cells present on the three agar cultures.

The meshes were also analysed after removal from the agar plates for the

presence of bacteria, using SEM and confocal microscopy. Figure 5.7a shows

LIVE/DEAD stained E. coli cells colonising the Φ1 mesh. The image shows

a high prevalence of dead cells, and a few clusters of live bacteria. In the

SEM image (Figure 5.7b) single bacterial cells can be seen to have adhered

onto and wrapped around the surface of the fibres; cell clusters correspond

with fibre crossover or adjacent fibres. When the fibre diameter was in the

range Φ2 bacterial cells proliferated within the fibrous network, using the

fibres as a support to move across the interstices of the mesh. The confocal

image (Figure 5.7c) reveals a high prevalence of live bacteria, both adhered

onto the surface of the fibres and clustered within the colony. The SEM

micrograph (Figure 5.7d) shows that bacteria were capable of progressively

colonising a region of the mesh composed of tens of fibres, developing a com-

pact system in which each cell was supporting the adjacent ones. On the

larger diameter fibres (Φ3), bacteria tended to adhere onto the fibre surface

and preferentially proliferate along single fibres, creating trains of aligned

138 / 223


Figure 5.6: Bacterial agar culture experiment. Crystal violet staining of agarcultures after mesh removal: (a) E. coli; (b) P. aeruginosa; and (c) S. aureus.

cells. Aligned bacterial colonies were also found between two adjacent fibres;

agglomerates of cells can be seen between two or more fibres in areas of fi-

bre cross over (Figure 5.7f). The confocal image (Figure 5.7e) shows a high

prevalence of live bacteria proliferating along the fibres. Results showed that

the Φ2 meshes, when exposed to E.coli solution or agar cultures, acted as a

scaffold, supporting and encouraging cell proliferation along the fibres and

through the interstices.

P. aeruginosa response to different fibre sizes (Figure 5.8) was similar to

E.coli, where cells colonising the Φ1 meshes were prevalently dead and iso-

lated onto the fibres (Figure 5.8a).

The SEM magnification of the mesh (Figure 5.8b) shows single cells adhered

and wrapped around the fibres as well as small agglomerates of cells at the

crossing over points between fibres. The Φ2 (1000 ± 100 nm) mesh provided

the best support for bacteria to adhere, spread and proliferate. Figure 5.8c

shows a high prevalence of live cells not only colonising the fibre surface but

also spread throughout the fibrous network; Figure 5.8d shows that after 1

hour incubation the cells that were attached onto the fibre surface, were in

the process of creating bridges among the fibres and creating a progressively

spread out colony. Cells appeared embedded in EPS presumably, this assists

139 / 223


Figure 5.7: Bacterial agar culture experiment. Confocal (a; c; e) and SEM (b;d; f) images of E. coli cells colonising PS electrospun meshes with fibre diameterranges: (a; b) Φ1 = 500 ± 200 nm; (c; d) Φ2 = 1000 ± 100 nm; (e; f) Φ3 =3000 ± 1000 nm. Scale bar: (a), (c), and (e) 5 µm; (b), (d), and (f) 2 µm.

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Figure 5.8: Bacterial agar culture experiment. Confocal (a; c; e) and SEM (b;d; f) images of P. aeruginosa cells colonising PS electrospun meshes with fibre

diameter ranges: (a; b) Φ1 = 500 ± 200 nm; (c; d) Φ2 = 1000 ± 100 nm; (e; f)Φ3 = 3000 ± 1000 nm. Scale bar: (a), (c), and (e) 5 µm; (b), (d), and (f) 2 µm.

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and ensures the support of the colony and the adhesion between adjacent

cells. As previously found for E. coli, when Φ = Φ3, P. aeruginosa preferen-

tially proliferated along the fibre surface. A high prevalence of live cells are

present in the confocal images (Figure 5.8e), indicating that bacterial adhe-

sion and proliferation were not impaired; SEM images showed that bacteria

tended to proliferate randomly on the fibre surface, without following the

alignment trend that was found recurrent for E. coli. Small cell agglomer-

ates can be seen between fibres crossing over each other, although most cells

covered the fibre surface (Figure 5.8f).

S. aureus cells were found to proliferate and cover the entire Φ1 mesh after

1 hour of incubation. Figure 5.9a shows colonies of live bacteria throughout

the fibrous network, with a very low percentage of dead cells. The SEM mi-

crograph (Figure 5.9b) confirms that S. aureus cells adhered onto the fibre

surface and proliferated within the mesh forming a compact system of cells,

supporting each other. When the larger fibres (Φ2 and Φ3) were exposed to

the bacterial culture, cells adhered and proliferated predominantly along the

fibre surface. Figures 5.9c and 5.9e show a high prevalence of live bacteria

attached onto the Φ2 and Φ3 fibres, respectively. The tendency of bacteria to

attach and proliferate onto the fibres is evident from Figures 5.9d and 5.9f,

where little evidence of bridging among fibres and colonising the interstices

of the meshes can be seen.

Discussion The influence of the average fibre diameter of PS electrospun

meshes on bacterial behaviour was investigated by initially culturing E. coli

cells in solution growth media, which is a most simplistic model of the wound

environment. The agar experiment was subsequently designed as this consti-

tutes a more realistic model that may better mimic a wound bed. The agar

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Figure 5.9: Bacterial agar culture experiment. Confocal (a; c; e) and SEM (b;d; f) images of S. aureus cells colonising PS electrospun meshes with fibre

diameter ranges: (a; b) Φ1 = 500 ± 200 nm; (c; d) Φ2 = 1000 ± 100 nm; (e; f)Φ3 = 3000 ± 1000 nm. Scale bar: (a), (c), and (e) 5 µm; (b), (d), and (f) 2 µm.

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culture experiment allowed the investigation of bacterial cell transfer from

the culture onto and within the meshes with different average fibre diameter.

When comparing the two methods of bacterial culture (suspension or agar),

the results illustrated that for each of the fibre sizes, the bacterial responses

were similar independent of the format of the bacterial exposure. This may

be in part due to the fact that in both cases, bacteria had all the nutrients

they required to survive available over the relatively short terms of these as-

says. To explore the general applicability of these results, future work would

need to look at longer term cultures and lower nutrient media and agar.

E. coli and P. aeruginosa are Gram-negative rod shaped bacteria 1-2 µm

long, that were found to have similar responses when interacting with the

three sets of meshes tested. The Φ2 mesh was composed of fibres with di-

ameter close to the cell length. In this case, the mesh was found to act as

a scaffold, encouraging bacterial growth throughout the fibrous substrate.

Bacteria were glued together in the empty spaces among the fibres through-

out the mesh where there is no other support than the EPS to sustain them.

On the Φ1 mesh, where fibre size was smaller than the bacterial length, a

distortion of the cell shape occurred, resulting in a high prevalence of dead

cells. Fibre diameters larger than the bacterial length (Φ3), resulted in cells

predominantly proliferating onto the surface of the fibres following aligned

or random directions, with a low degree of bridging among and across fibres.

S. aureus, a Gram-positive round shape bacterium 0.5-1 µm diameter, showed

the highest proliferation rate with the Φ1 mesh, where fibre diameter was

close to bacterial size. On the Φ2 and Φ3 meshes, where fibre diameter was

bigger than the bacterial size, S. aureus preferentially proliferated onto the

surface of the fibres.

These results show that the average fibre diameter of the mesh does in fact

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influence the capacity of bacteria to adhere, proliferate and form colonies.

This influence is directly linked to bacterial size and shape. In fact, results

show that for the three bacterial species considered, the highest spreading

and proliferation was found to occur when the fibre diameter was close to

bacterial size. These findings can be related to the ”attachment points”

theory, according to which organisms smaller than the scale of the surface

texture have greater adhesion strength due to the availability of multiple at-

tachment points, in comparison to microorganisms that are larger than the

surface texture [199, 200]. The theory also states that small round shape

bacteria, such as S. aureus, exhibit a different attachment pattern compared

to the bigger, elongated cells, due to the different number of accessible at-

tachment points. This is in agreement with the observation that S. aureus

had the highest attachment and proliferation rate on the smallest Φ1 fibres,

while the rod shape bacteria colonised the Φ2 mesh preferentially [201].

These findings open up the possibility to design fibrous meshes of hetero-

geneous morphology to suppress the growth of different bacterial species in

complex environment such as a chronic wound bed. These results suggest

the possibility of designing an innovative wound dressing that, instead of

killing the bacteria present in the wound bed by releasing an antimicrobial

agent, could attract the microbial cells from the wound bed. Once attracted

towards the mesh, the bacteria would find a fibrous substrate with suitable

fibre size to ensure bacterial anchoring and cell morphology distortion to sup-

press their proliferation, thus protecting the wound bed. Future studies will

be undertaken to investigate the relative rate of attachment on the fibres of

each bacterial species present within a wound, as this information will affect

the design of a dressing capable of trapping bacteria with different size and

shape and attachment mechanisms.

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To attract the bacteria from the wound bed towards the mesh, one strategy

could be strategically designing the fibre surface chemistry. However, a sys-

tematic study on the influence of fibre surface chemistry properties on the

capacity of bacteria to attach and proliferate does not yet exist.

5.3 Influence of fibre surface chemistry on

bacterial behaviour

To investigate the influence of fibre surface wettability, chemical functional

groups and surface charge on bacterial behaviour, the plasma modified meshes

(ppAAc, ppCo, ppOct, and ppAAm) were exposed to confluent layers of

E.coli cells grown on agar plates and an inhibitory zone experiment was per-

formed. For these studies the Φ1 = 500 ± 200 nm meshes were used due to

the high potential of nanofibrous meshes as wound dressings, as described in

chapter 1, section 1.3.

A silver impregnated mesh used as control produced a clear zone of inhi-

bition on the agar culture. Figure 5.10 shows the transparent area around

the mesh where the bacterial proliferation was impaired by the release of

the antibacterial compounds. No inhibitory ring was induced by the plasma

polymerised meshes. It is clear from the images that the bacteria reached

the edges of all the plasma coated meshes (Figure 5.10), indicating that the

bacteria proliferated around the meshes and no inhibition occurred.

To investigate the transfer of bacterial cells onto the meshes a combination

of SEM and confocal imaging after viability staining was used. To obtain

robust results, for each surface chemistry including the untreated PS, three

meshes were characterized with SEM and the other three meshes with con-

focal microscopy.

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Figure 5.10: Photograph of the plasma coated and silver releasing meshesexposed to E. coli layer.

Figure 5.11 shows LIVE/DEAD stained E.coli cells colonising the plasma

coated meshes and the PS control mesh. The presented images are repre-

sentative of the results. The green colour of the fibres is due to the intrinsic

autofluorescence of the PS material at the selected excitation wavelengths

(480-490 nm). As shown in the previous section, a significant proportion of

dead bacterial cells were adhered on or wrapped around the fibre surface of

the untreated PS mesh (Figure 5.11a). Dead bacteria, fluorescing red, were

predominantly isolated onto the fibres or aggregated in small clusters at the

crossover between fibres. On the ppAAc mesh (Figure 5.11b) a lower number

of bacteria was present, with few isolated dead cells attached onto the surface

of the fibres. The highest proportion of live cells was found on the ppAAm

coated fibres. Figure 5.11c shows very few dead cells adhered onto the fibres

and large clusters of live bacteria, fluorescing green, that colonised the mesh

by bridging and spreading across the fibres. Cells not only attached onto

the fibre surface but also spread in the interstices and empty spaces within

the mesh forming compact colonies. The ppOct coating (Figure 5.11d) had

a higher proportion of live cells compared to the untreated PS with the cells

clustering at fibre crossovers or between adjacent fibres. The ppCo coatings

induced a minor attachment of bacterial cells compared to the untreated

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Figure 5.11: Confocal images of LIVE/DEAD stained E.coli cells onto (a)untreated PS mesh; (b) ppAAc; (c) ppAAm; (d) ppOct; and (e) ppCo meshes

after removal from the E.coli agar culture. Scale bar 5 µm.

PS. Most cells appeared dead, isolated onto the fibre surface. No bacterial

clusters were found on the mesh (Figure 5.11e). The reported results were

reproducible across 3 separate experiments, during which 10 images were ac-

quired on each mesh.

SEM micrographs of the untreated PS and plasma coated meshes exposed to

the E.coli cultures were compared. Figure 5.12 shows representative images

belonging to the set of 15 SEM micrographs that were acquired for each sur-

face chemistry.

The images confirmed that on the untreated PS mesh (Figure 5.12a) bacterial

cells were predominantly isolated, wrapped around the fibres, or embedded in

small clusters composed of few cells at fibre crossovers. The ppAAc coating

resulted with the lowest proportion of cells attached onto the fibres com-

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pared to the untreated mesh as well as the other plasma coatings. Figure

5.12b shows single isolated cells attached onto the fibre surface or in the in-

terstices between close fibres.

The SEM of the ppAAm coated meshes (Figure 5.12c) confirmed the results

shown by the confocal images. The coating resulted with the highest propor-

tion of cells that spread throughout the fibrous network forming colonies and

compact clusters across the interstices of the mesh. Figure 5.12d shows the

bacterial cells that transferred from the agar culture onto the ppOct mesh.

There was a proportion of cells attached onto the fibres, forming clusters at

fibre crossovers or in the interstices between few fibres. Bacteria were capa-

ble of bridging across fibres and spreading through the empty spaces of the

mesh, forming an agglomerate of tens of cells. The ppCo coating induced a

lower attachment of E.coli cells compared to the ppOct and ppAAm meshes.

Figure 5.12e shows a prevalence of isolated cells wrapped around the fibres,

with no clusters at fibre crossovers or across the interstices of the mesh.

Figure 5.12: SEM images of (a) untreated PS; (b) ppAAc; (c) ppAAm; (d)ppOct; and (e) ppCo coated meshes after removal from the E.coli agar culture.

E.coli cells were false coloured in red. Scale bar 2 µm.

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Discussion Plasma polymerisation was used to modify the surface chem-

istry of the electrospun PS meshes through the deposition of thin polymeric

films onto the fibres. The surface chemistry was confirmed in chapter 4,

showing that the fibres had the chemistry typically associated with each of

the selected plasma polymers.

The monomers used for the surface modification of the fibres were chosen

to generate different chemical functionalities. The ppAAc coating is a hy-

drophilic carboxyl rich film, negatively charged under neutral pH [178, 202],

while ppAAm is an aminated, positively charged coating [182]. The ppOct

and ppCo coatings are hydrophobic hydrocarbon rich films [22].

Physicochemical factors, including roughness, wettability and chemistry of

the substrate surface have all been found to be drivers of bacterial attach-

ment. In addition, the surface properties of the bacterial cells, including

surface hydrophobicity and surface charge, were shown to play a significant

role in the attachment processes [85, 203]. In the literature, several au-

thors have investigated the attachment and spreading of bacteria onto flat

surfaces with different degrees of wettability. The attachment of different

bacterial species including isolates of Staphylococcus epidermidis and E.coli

was shown to be more effective on hydrophobic substrates [178, 204–206] due

to the so-called ”hydrophobic effect” occurring between the substrate surface

and the hydrophobic residues present on the bacterial cell surface [85]. The

hydrophobic effect has been considered to be nonspecific and the literature

provides evidence that a large number of bacteria and fungal pathogens de-

pend on hydrophobic interactions for the successful colonization of a surface

[207]. Doyle et al. listed the most common structures contributing to the

hydrophobicity of the bacterial cell surface, including nonpolar groups on

fimbriae, lipopolysaccharides, and outer membrane proteins [207].

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The hydrophobicity of the ppOct coating could be one of the reasons for the

high proportion of clustered live cells found on the mesh. However, other fac-

tors must come into play since the characterisation of the other hydrophobic

material, the ppCo coating [22], showed less transferal of bacteria and the

few cells that attached onto the fibres were dead and isolated. The PS fibres,

which are also hydrophobic [208], exhibited a similar proportion of attached

bacteria compared to the ppOct coating, but the cells found on the untreated

fibres were predominantly dead.

In the inhibition assay, the ppCo coating along side all the other plasma

polymers, did not inhibit bacterial growth, indicating that, as shown in the

previous work from Pegalajar-Jurado et al., the cineole film was not leach-

ing any antimicrobial agents [22]. In the same work, Pegalajar-Jurado et al.

demonstrated the antibacterial activity of cineole against E.coli cultured in

solution. Authors showed that plasma polymerised films produced from 1,8-

cineole on flat substrates retained part of the antimicrobial activity against

suspension culture of E.coli and S.aureus, by resisting bacterial attachment

after 18 hours and biofilm formation after 5 days of incubation. The ppCo

deposited onto the nanofibres did not significantly reduce bacterial attach-

ment and transfer when compared to the untreated fibres, suggesting that

the coating did not retain the antibacterial activity of the original monomer

in the first hour of exposure to the culture. These results indicate that the

ppCo chemistry on the PS fibres would not significantly contribute to impair

initial bacterial attachment for short term contact applications; the coating

might be beneficial in the long term, if the fibrous PS substrate will reproduce

the same results obtained by Pegalajar-Jurado et al. on flat glass surfaces.

Future work will include studies to investigate bacterial transfer onto ppCo

coated meshes in the long term.

151 / 223


The ppAAc coating resulted in the lowest proportion of bacterial cells trans-

ferred from the culture onto the fibres compared to the untreated PS and

the other coatings. The hydrophilicity and negative charge of the carboxyl

groups in the coating could be one of the main reasons for the low attraction

towards the bacterial cells. As most bacteria, including E.coli, carry a net

negative surface charge, the initial attachment of the cells is discouraged on

negatively charged surfaces by electrostatic repulsion [209]. Phosphodiester

bonds of teichoic acids are responsible for the net negative charge of the

Gram-positive cell wall while lipopolysaccharides impart a strongly negative

charge to surface of Gram-negative bacterial cells [210].

The ppAAm coating, which is also moderately hydrophilic, showed the high-

est proportion of live cells. This could be due to the presence of amine

groups and/or the positive charge carried by these functional groups. In

fact, amine groups have been found to promote bacterial and protein inter-

actions, thus encouraging the attachment of the cells onto the surface [211].

Moreover, bacterial attachment could be promoted by the positive charge of

these groups due to the attraction towards the net negative charge of the

bacterial membrane [209].

These results were obtained on the nanofibre meshes which were previously

shown to induce the death of elongated bacteria due to the cell morphological

changes induced by the small fibre diameter. The high proprtion of viable

cells on the ppAAm mesh suggests that the coating overtakes the effects of

fibre morphology on bacterial behaviour, at least in the short term. Further

studies will be performed to investigate the responses of E.coli to ppAAm

coated nanofibre meshes in the long term (12-24 hour), to understand if the

high proportion of viable cells is maintained over time.

These results underline the complexity of the mechanisms involved in the

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attachment of bacteria onto substrates, highlighting that this process is gov-

erned by a combination of surface properties of both substrate and cells. The

hydrophobic effect could be one significant driver encouraging the initial at-

tachment attachment onto the ppOct mesh. However surface hydrophobicity

is not the only parameter affecting the attachment process since a signifi-

cantly lower proportion or viable clustered bacteria was found on the other

hydrophobic surface chemistries (PS and ppCo). Chemical functionalities

and surface charge could be significant parameters affecting bacterial trans-

fer and attachment onto the ppAAc and ppAAm coated meshes.

The results suggest that the possibility exists for using fibre surface chem-

istry as a tool to control bacterial interactions, at least in the short term. For

instance, the ppAAc coating could be a suitable candidate for those devices

that need to minimise the attachment of bacterial cells onto the surface, in

the short term, during the initial exposure. The ppAAm chemistry could

instead be used for systems that need to be attractive towards bacteria, such

as an innovative wound dressing that can clean up the wound bed by attract-

ing and trapping the bacterial cells during the initial contact.

The present work also highlights that plasma polymers constitute an advan-

tageous approach for controlling and tailoring the surface chemistry of any

substrate. In fact, the plasma polymerisation process is transferable and re-

producible between a large range of different materials without the need for

specific substrates. A wide variety of chemical functionalities can be gener-

ated onto traditional materials without affecting the bulk properties nor the

surface morphological features.

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5.3.1 Bacterial transfer onto ppAAm coated meshes

The ppAAm coating was shown to be a promising candidate for developing

attractive surfaces that could potentially be used to clean up the wound bed

from bacterial contamination. The question arises if the bacteria found on

the ppAAm meshes were mostly coming from the agar culture or from the

proliferation of the cells that initially attached onto the fibres or a combina-

tion of the two phenomena.

The design of a specific experiment was required to understand if the ppAAm

coating encourages bacterial transfer and attachment or bacterial prolifera-

tion or both.

Control ppAAm meshes were incubated on E.coli agar cultures for 1 hour at

37◦C; a second set of ppAAm meshes was incubated on the same cultures for

30 minutes only. After 30 minute incubation on the E.coli biofilm, a third

set of meshes was transferred onto clear agar plates (no bacteria present) and

maintained at 37◦C for other 30 minutes. The third set of meshes allowed to

evaluate the proliferation of the cells that initially transferred and attached

onto the ppAAm meshes from the agar culture.

Figure 5.13 shows the number of bacteria quantified on the three sets of

meshes.

The control ppAAm meshes (Set 1) had the highest number of bacterial cells

attached onto the fibres (46x103 ± 20x103 bacteria/mm2). On the meshes ex-

posed to same culture for 30 minutes (Set 2), 22x103 ± 9x103 bacteria/mm2

were quantified. Similar numbers (21x103 ± 8x103 bacteria/mm2) were ob-

tained from the meshes that were transferred onto the clear agar plates (Set

3). The number of bacteria on the Set 1 meshes had a statistically significant

difference from the numbers obtained on the Set 2 and 3 (according to the

t-test for two independent samples, for p<0.005)

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Figure 5.13: Quantification of E.coli cells that transferred onto the ppAAmcoated meshes from the agar plates at different culturing conditions.

Discussion The number of bacteria quantified on the control ppAAmmeshes

(Set 1) was almost double the number obtained on the meshes incubated on

the E.coli culture for 30 minutes (Set 2). This result suggests that with the

additional 30 minutes of contact with the agar, the number of bacteria within

the meshes doubled either because more cells transferred from the agar cul-

ture or because the cells initially transferred to the mesh proliferated.

The numbers of bacteria quantified on the ppAAm meshes exposed to the

biofilm for 30 minutes (Set 2) were not significantly different from the num-

bers obtained on the meshes transferred onto the clear agar plates (Set 3).

These results show that when the bacterial cells were put in the condition

to proliferate but not transfer (Set 3), the number of cells present within

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the meshes did not significantly increase. This suggests that the majority of

the cells found on the control ppAAm meshes (Set 1) were transferred from

the agar culture instead of deriving from the proliferation of the cells that

initially attached onto the fibres.

These results indicate that the ppAAm coating encourages predominantly

cell transfer and adhesion rather than cell proliferation. As previously men-

tioned, these results were obtained on the nanofibre meshes, which were

previously shown to discourage cell proliferation throughout the mesh due

to the small diameter of the fibres compared to the bacterial length. The

ppAAm coating could constitute an attractive chemistry for initial bacterial

adhesion, while the fibre size could be responsible for the slow proliferation

of the cells throughout the mesh.

The obtained results confirm that the ppAAm coating encourages the trans-

fer of the bacterial cells from the underlying culture onto the coated fibres.

The possibility exists to combine the ppAAm coating with nanoscale fibres

to develop a dressing that attracts rod shape bacteria from the wound bed

and traps them within the fibrous network.

5.4 Conclusions

The diameter of electrospun PS fibres was shown to influence the ability of

E.coli, S.aureus, and P.aeruginosa to proliferate and colonise the fibrous sub-

strate. SEM and confocal images indicated that bacterial spreading through-

out the mesh depended on fibre diameter and bacterial size and shape.

Meshes with an average fibre diameter close to bacterial size were found

to offer the best support for bacterial adhesion and spreading, constituting a

scaffold that bacteria use as a framework for forming colonies. For rod shape

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elongated cells (E.coli and P.aeruginosa), fibre diameters smaller than the

bacterial length resulted in most cells wrapping around each fibre, limiting

the ability of bacteria to easily create bridges across fibres and form colonies.

These bacteria exhibited similar behaviour, colonising preferentially the 1

µm meshes.

Round S.aureus cells showed the highest proliferation throughout the nanofi-

brous substrates; in the presence of bigger fibres, the cells preferentially ad-

hered on the fibre surface, without spreading throughout the mesh.

Controlled fibre morphology could be combined with fibre surface chemistry

to control the bacterial load in chronic wounds. In fact, the strategic com-

bination of a variety of surface chemistry features (fibre surface wettability,

functional groups and surface charge) could allow the control over the bac-

terial attachment and proliferation processes onto fibrous substrates in the

short term.

Fibre surface chemistry was shown to influence the ability of E.coli cells to

adhere and proliferate within the electrospun meshes. The ppAAm coating,

hydrophilic and rich in amine postively charged groups, induced the highest

attraction of viable cells from the underlying agar culture. A significantly

lower number of E.coli cells were found to adhere onto the hydrophilic ppAAc

meshes, possibly due to the negative charge of the coating, while the ppOct

meshes resulted with a higher proportion of clustered live bacteria when com-

pared to the untreated PS mesh. The ppCo did not show any inhibitory effect

on the bacteria growing in the surrounding culture but a high proportion of

dead isolated cells were found adhered to the fibres.

The presented results were obtained investigating the interactions of bacteria

with fibres in the short term, tackling initial bacterial attachment and prolif-

eration. This stage needed to be explored as it constitutes the beginning of

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bacterial infection: the discovery of solutions to control bacterial behaviour

in the first few hours of contact with the fibre surface could allow the preven-

tion of wound infection, thus significantly improving the treatments outcomes

and patient lifestyle.

The results presented in this chapter underline the complexity of the envi-

ronment that wound dressings are designed to interact with. Bacteria with

different morphologies were shown to respond in a distinctive manner to

different fibre sizes; since chronic wounds are contaminated by a variety of

bacteria, with different sizes and shapes, fibre diameter may not be the only

strategy used to limit the bacterial load in the wound bed. The control over

fibre size could potentially be combined with strategically designed addi-

tional fibre properties, such as surface chemistry or controlled release. Apart

from providing a device that could minimise bacterial growth in a wound

bed, the possibility also exists for developing a mesh capable of ”attracting

and trapping” bacteria from the wound bed. The ppAAm coating could be

a suitable chemistry for attracting the bacteria from the wound bed towards

the dressing, while the controlled fibre diameter could constitute a trap to

be used to clean up the wound.

158 / 223

Chapter 6

Skin and bacterial cells transfer

onto electrospun meshes

Contents6.1 Cell transfer studies . . . . . . . . . . . . . . . . . 160

6.2 Wound models . . . . . . . . . . . . . . . . . . . . 168

6.2.1 Superficial partially de-epidermised wound . . . . . 170

6.2.2 Superficial de-epidermised wound . . . . . . . . . . 173

6.2.3 Deep wound . . . . . . . . . . . . . . . . . . . . . . 175

6.2.4 3-Dimensional deep infected wound . . . . . . . . . 178

6.3 Conclusions . . . . . . . . . . . . . . . . . . . . . . 185

The work presented in this chapter was performed in Prof. Sheila Mac-

Neil’s laboratories at the Kroto Research Institute at The University of

Sheffield, UK, a leader in the development of tissue-engineered models of

human skin [117, 137, 138, 212]. This work was supervised by Prof. Sheila

MacNeil, Prof. Ian Douglas, Dr. Anthony Bullock, and Dr. Marc Daigneault.

The chapter aims to investigate the mechanisms of skin cell and bacterial

transfer onto and within the ppAAc, ppCo, ppOct, and ppAAm plasma

159

6. Skin and bacterial cells transfer onto electrospun meshes

coated meshes. The initial in vitro assay was developed by culturing fibrob-

lasts and keratinocytes on tissue culture plates and studying cell transfer

from the surface into the meshes, in a method that was analogous to the

bacterial techniques used in chapter 5.

More complex tissue engineered models of wounds at different skin depths

and degrees of severity were fabricated by co-culturing skin cells (fibroblasts

and keratinocytes) in decellularised human skin, that was then subject to a

wound simulation and infected with bacteria (P.aeruginosa). The models of

superficial and deep wounds, infected and not, were characterised through

histological analysis and cell viability assays. The transfer of the bacterial

and skin cells from the models onto the plasma modified meshes was investi-

gated with a combination of the MTT viability assay and confocal microscopy

after fluorescent staining of the cells.

6.1 Cell transfer studies

To investigate the mechanisms of transfer of skin cells onto and into the elec-

trospun meshes, the plasma coated meshes and PS control mesh were placed

on sub-confluent layers of fibroblasts and keratinocytes separately cultured.

The MTT assay was performed on both meshes and well plates after mesh

removal to investigate the transfer of viable cells that occurred from the bot-

tom of the plates onto the meshes.

Figures 6.1a and 6.1b show the microscopy images of the 70-80% confluent

fibroblast and keratinocyte cultures respectively that were used for the exper-

iment. When confluent, fibroblasts appeared large and flat, with elongated

processes protruding from the cell body, creating a spindle-like appearance.

Keratinocyte cells when confluent displayed their typical cobblestone pat-

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tern with tight cell-cell junctions. The brighter spots that can be seen on the

culture (Figure 6.1b) correspond to patches of differentiated cornified (or ker-

atinazed) keratinocyte cells, which are dead squamous cells that do no longer

multiply [213]. Figure 6.2 shows the photographs of the electrospun meshes

Figure 6.1: Bright-field optical microscopy images images of 70-80% confluentcultures of (a) human dermal fibroblasts; (b) human dermal keratinocytes. Scale

bar 100 µm.

and the 6 well plate after the MTT assay was performed on the fibroblast

culture. It can be observed that the plasma coated meshes and PS control

mesh (Figure 6.2a), originally white in colour, presented a patterned purple

area that is particularly evident on the ppAAc, ppCo and ppAAm meshes.

The purple areas are indicative of the presence of viable cells that transferred

from the bottom of the well plate onto the meshes. The patterned area of

circles replicates the geometry of the grid that was used to hold the meshes

in contact with the bottom of the plate. The untreated PS mesh showed

viable fibroblast cells spread across the surface of the mesh. On the ppOct

coated mesh small purple patches were predominantly in the centre of the

mesh. The ppAAc, ppCo and ppAAm appeared yellow in colour, while the

untreated and ppOct meshes remained white. This discolouration is due to

the absorption into the mesh of the original MTT solution, which is yellow

in colour.

The images of the MTT stained wells after mesh removal (Figure 6.2b)

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Figure 6.2: Photographs of (a) electrospun meshes; (b) 6 well plate afterfibroblast transfer experiment. The viable fibroblast cells that transferred on the

meshes or remained on the plates are stained purple.

showed both the level of cell transfer and the viability of cells that remained

attached to the well after mesh removal. In the control TCPS well, where no

mesh was present, a significant portion of the bottom area stained purple,

illustrating the high viability of the cells across the surface of the well. The

wells where the untreated PS, ppAAc, ppCo and ppAAm meshes were incu-

bated clearly show regions of viable cells remaining outside the area where

the meshes were placed, with little or no staining underneath the meshes.

This indicates that cells had either transferred to the meshes or were still on

the well surface but not viable after mesh removal. The well corresponding

to the ppOct mesh showed a stained area comparable to the control well,

indicating that viable cells were still attached to the well bottom after mesh

removal.

The solubilisation of the MTT dye from both the meshes and the wells al-

lowed the quantification of the viable cells that transferred onto the meshes

and of those that remained attached to the wells (Figure 6.3). Considering

the absorbance of the MTT dye from the well plates, it can be noted that

where the ppOct mesh was placed resulted with the closest value (0.24 ±

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Figure 6.3: Absorbance at 570 nm of the MTT dye dissolved from the fibroblastcultured well plates and dressings.

0.12) to the control well (0.31 ± 0.10), where no mesh was placed. The wells

corresponding to all the other meshes produced lower values (0.12-0.15 ±

0.02-0.06). This result suggests that the ppOct mesh had the least cell trans-

fer, while the other coatings had similar numbers of viable cells transferred

from the wells. The MTT absorbance values from the meshes reproduced

the same trend. In fact, the MTT absorbance from the ppOct mesh (0.02 ±

0.02) was the lowest compared to all other meshes, indicating that the ppOct

coating had the least transfer of viable cells. The very low absorbance values

obtained from the meshes (0.02 - 0.06) highlight the challenge associated to

solubilise the MTT from these substrates. The 3-dimensional fibrous struc-

ture of the meshes make the MTT solubilisation an empirical process with

the intrinsic possibility that some MTT material may remain trapped in the

meshes, reducing the total signal. In addition, the volume of the solubilis-

ing solution needs to be sufficient to cover the entire surface of the meshes;

however this results in a significant dilution of the extracted MTT material,

thus the low absorbance measures.

The high variability encountered in the absorbance values of the MTT dye is

due to the variability that intrinsically characterises the experiment, due to

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the different patient donors used for the isolations of the cells used in each

experiment. Although a statistically significant difference amongst the MTT

absorbance values was not found, the data show a specific trend that con-

firms the results derived from the photographs of the MTT stained meshes

and plates (Figure 6.2).

Figure 6.4 shows the electrospun meshes and 12 well plate after the MTT as-

say was performed on the keratinocyte culture. Also in this case the ppAAc,

ppCo and ppAAm meshes absorbed the MTT solution and were discoloured,

while the PS and ppOct re-achieved the original white colour after the rins-

ing steps of the MTT protocol (Figure 6.4a). None of the meshes showed

Figure 6.4: Photographs of (a) electrospun meshes; (b) 12 well plate afterkeratinocyte transfer experiment. The viable keratinocyte cells that transferred on

the meshes or remained on the plates are stained purple.

any purple area, thus suggesting that no viable cells were transferred from

the well plate into the meshes. This is confirmed by the photograph of the

MTT stained 12 well plate (Figure 6.4b). Each well showed a stained area

comparable to the control well, where no mesh was incubated. In each well

except the control one, a thin round transparent circle can be seen. This

corresponds to the edges of the metal ring that was used to hold the meshes

at the bottom of the wells during the experiment.

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The absorbance values of the MTT dye dissolved from the cultured plates

and the meshes are shown in Figure 6.5. No significant difference in the

MTT absorbance from the well plates was found to occur between each well

and the control, where no mesh was placed, confirming that most viable ker-

atinocytes were left at the bottom of the wells. The same result was further

confirmed by the MTT absorbance values from the meshes, which were zero

for all the plasma coated meshes and the PS control mesh, indicating that

no viable cells were detected on the dressings.

Figure 6.5: Absorbance at 570 nm of the MTT dye dissolved from the HDKcultured well plates and dressings.

Discussion The cell transfer studies from the bottom of well plates onto

and into the plasma coated meshes were performed with the ultimate goal

of understanding if a device capable of preventing cell ingrowth could be

designed. This is an essential requirement to avoid wound damage upon

dressing removal.

All the tested meshes were found to remove fibroblast cells from the bottom

of the well. In fact all the meshes showed purple areas or patches, indicative

of the presence of viable cells. The ppOct mesh was found to remove the

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least number of cells, while the other hydrophobic materials (PS and ppCo

coating) showed a significant proportion of cell removal, comparable with the

hydrophilic surfaces (ppAAm and ppAAc).

The attraction of the ppAAm and ppAAc coatings towards fibroblasts was

hypothesised due to the number of studies in the literature that prove the en-

couraged attachment of mammalian cells onto hydrophilic surfaces. Several

authors investigated the influence of various surface chemistries generated

through plasma polymerisation on the attachment and spreading of epithe-

lial cells in solution cultures. Hamerli et al. showed that human dermal

fibroblasts tend to attach and spread much more widely on hydrophilic sur-

faces than on hydrophobic ones. In particular, the attachment and viability

of human dermal fibroblasts on allylamine plasma coated polyester surfaces

were found to be more extensive than on the untreated control hydrophobic

substrates [108]. In another work, Hamerli et al. demonstrated that amine

groups are highly attractive for fibroblasts adhesion. In fact plasma-modified

polyethylenterephtalate (PET) membranes with amine containing coatings

were found to offer better adhesion and growth of fibroblasts than untreated

surfaces. Other authors have demonstrated that amine and amide containing

plasma polymers are capable of encouraging the attachment and growth of

endothelial cells [214]. The carboxylic acid functionality introduced by the

ppAAc coating has also been shown to promote cell adhesion [187, 215, 216].

The higher cell transfer onto the untreated PS and ppCo compared to the

ppOct coating was not expected given the hydrophobicity of these surfaces.

In fact, it has been shown that the extracellular matrix (ECM) proteins re-

quired for cell adhesion tend to adsorb in low quantities and/or denature

on hydrophobic surfaces [22]. However, this assessment is contradictory in

the literature as other authors have demonstrated that depending on the

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cell type, higher cell adhesion strength onto hydrophobic surfaces can occur

[217, 218]. In addition to surface wettability, other properties of the sub-

strate such as surface charge, rigidity, and the specific chemical composition

of the materials can come into play during the cell-surface interaction pro-

cesses [218]. Untreated PS, ppOct and ppCo have been previously shown

to limit cell attachment when exposed to a culture of fibroblasts [22, 219].

However, these studies focused on the response of solution cultures of cells

seeded onto flat substrates. In the present study the meshes were exposed to

70-80% confluent layers of fibroblasts and the transfer mechanisms were in-

vestigated. In this type of experimental setup different phenomena affecting

cell attachment might come into play. The results suggest that cell attach-

ment behaviour might be driven by different factors when cells are given a

choice to transfer from a culture or attach from a solution. Currently the

mechanisms of mammalian cell attachment onto a substrate are well inves-

tigated, but this knowledge only covers the case of the substrate exposed

to a solution culture of cells. The present results suggest that fundamental

studies need to be performed to understand which mechanisms affect cell be-

haviour when a substrate is exposed to cells that already have attached onto

a different surface and are given the choice to transfer or remain attached.

The second part of the experiment consisted in evaluating the transfer of

keratinocytes onto the plasma coated meshes from the bottom of well plates.

No viable cells were detected on the meshes and there was little evidence

of disruption of the cells on the TCPS surfaces, suggesting that no transfer

occurred.

In the literature, keratinocyte cells were found to have similar behaviour to

the fibroblasts, in terms of adhesion and proliferation when seeded onto the

plasma coated substrates. Various studies demonstrated that keratinocytes

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preferentially adhere onto ppAAc and ppAAm coated surfaces in comparison

to the the ppOct and other hydrophobic chemistries [106, 107, 220]. These

results were found when a solution culture of keratinocytes was seeded onto

the coated surfaces; in a ”transfer” condition, when the cells are cultured

onto a solid substrate and are given the choice to transfer on the meshes,

no transfer was found to occur on any of the tested meshes. As previously

observed for the fibroblasts, likewise keratinocyte attachment onto a given

surface chemistry was shown to be different depending on the culture condi-

tions (transfer or solution). Keratinocyte cells tend to develop tight junctions

between cells when adhered onto a surface, strongly binding amongst each

other and to the bottom of the substrate. This could be a significant reason

that prevented the transfer of the cells onto the meshes.

6.2 Wound models

To mimic wounds at different depths and stages of healing, four in vitro

wound models were developed. The models were used to further investigate

the transfer of skin cells and bacteria onto the plasma coated meshes.

Skin wound models were created by culturing human dermal fibroblasts

and/or keratinocytes in de-epidermised and de-cellularised split thickness

skin grafts. Figure 6.6 shows the histology images of the H&E stained

split thickness skin graft before (Figure 6.6a) and after (Figure 6.6b) de-

epidermisation and de-cellularisation.

Figure 6.6a exhibits the three layers forming the human skin. The outermost

layer, or stratum corneum, consists of layered differentiated keratinocytes

(bright pink on Figure 6.6a); the epidermis, composed of proliferating and

differentiating keratinocytes, is the intermediate violet layer and the darker

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spots correspond to the cellular components present within the tissue; the

deepest layer of the skin, the dermis, is formed by the more superficial, dense

papillary region and a deeper and thicker area known as the reticular der-

mis. The dermis corresponds to the clear pink layer on Figure 6.6a and the

darker areas visible within the tissue are the cellular components, including

fibroblasts, that were originally present in the STS [27].

The skin specimen after de-cellularisation (Figure 6.6b) was composed of the

dermis only (pink structure), with the outermost layers entirely removed as

well as the cellular components. In fact, the darker spots are no longer visible

throughout the tissue.

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Figure 6.6: H & E histology images of the skin specimens (a) before and (b)after the de-cellularization and de-epidermization of the split thickness skin

grafts. Scale bar 20 µm

6.2.1 Superficial partially de-epidermised wound

Superficial partially de-epidermized wound model corresponds to a wound

that lost the stratum corneum and part of the epidermis, maintaining a su-

perficial layer of differentiating keratinocytes and an intact basement mem-

brane (BM). According to Ghosh et al. [137] the keratinocytes tend to attach

and grow on the top of the BM, initiating the process of restoration of the

epidermis, while the fibroblasts spread throughout the dermis. Figure 6.7

shows the H & E histology image of the reconstructed skin mode; the thick

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Figure 6.7: H & E histology image of the superficial partially de-epidermisedwound model. Scale bar 20 µm

purple layer on the surface of the dermis correspond to the keratinocyte cells,

that have attached to the basement membrane and proliferated to initiate

the restoration of the epidermis.

The untreated and plasma coated meshes were incubated on the skin model

for 3 days. After mesh removal, a MTT assay was performed on both the skin

specimens and the meshes. Figure 6.8 shows the photograph of the meshes

and skin specimens after the MTT assay. The skin specimens that were in

contact with the meshes showed a visible purple ring in the center (Figure

6.8a), corresponding to the viable cells that were seeded and cultured. The

coloured area of these specimens was similar to the purple spot found on the

control sample, where no mesh was placed. None of the meshes reported

significant purple stained areas (Figure 6.8b), indicating that no viable cells

were present on the meshes.

These results show that viable cells were present in each skin specimen and

no transfer onto the meshes occurred. It was observed that during the three

repeats of the experiment, ppAAc and ppAAm meshes remained partially

attached to the skin specimens. Figure 6.9a is a photograph of the skin

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Figure 6.8: Photographs of (a) skin specimens; (b) electrospun meshes afterMTT assay. No transfer of viable cells occurred from the skin specimens onto the

meshes. The wound model was developed to mimic a superficial partiallyde-epidermized wound, with keratinocytes differentiating above the BM and

fibroblasts spread through the dermis.

Figure 6.9: Photographs of the skin specimen onto which the ppAAc mesh wasplaced. (a) The purple ring corresponding to an area of viable cells can be

visualised; (b) the edges of the skin specimen and of the fibre layers that remainedadhered onto the skin after mesh removal are underlined in blue and red

respectively.

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specimen onto which the ppAAc was placed. Figure 6.9b underlines the

edges of the skin specimen, in blue, and the outermost fibre layers of the

mesh (red) that separated from the fibrous substrate while it was removed

and stayed adhering onto the skin. The violet ring corresponding to the area

containing viable cells is clearly evident.

6.2.2 Superficial de-epidermised wound

The superficial de-epidermized wound model mimics a wound that lost stra-

tum corneum and epidermis, maintaining a intact BM. The H & E histology

image of the model is reported in Figure 6.10. The fibroblast cells, stained in

dark violet on the image, can be distinguished throughout the dermis, from

the reticular layer up to the papillary dermis, remaining underneath the BM.

After the meshes were cultured on the wound model for 3 days, the MTT

assay on the plasma coated meshes revealed that no viable cells were trans-

ferred from the wound model, since none of the meshes placed on the skin

specimens showed purple stains (Figure 6.11b). As previously observed the

ppAAc, ppCo and ppAAm meshes retained the MTT dye after the rinsing

steps, thus appearing yellow coloured, in comparison to the PS and ppOct

meshes that instead re-acquired the original white colour.

All the skin specimens showed a purple circular area, corresponding to the

viable fibroblasts that proliferated underneath the BM (Figure 6.11a) and

did not transfer onto the meshes. The size and colour of the rings on the

specimens in contact with the meshes were similar to the ring on the con-

trol specimen, where no mesh was placed. However, slight variations in the

purple intensity from one skin specimen to the other might be observed in

Figure 6.11a, where the ppCo specimen exhibits the darkest purple spot and

the PS and ppAAm the brightest. This could suggest that the meshes with

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the paler spots caused the death of some fibroblasts. However, the possibility

also exists that the colour intensity differences are related to the complex-

ity of the experiment design. In fact, the same number of fibroblasts were

initially seeded in the skin specimens but there was not control over cell pro-

liferation during the three day culture, resulting with an unknown number

of viable cells present in the models and exposed to the meshes. The cur-

rent experimental approach allows a qualitative evaluation of the transfer of

viable cells onto the meshes from the skin specimens. The obtained results

indicate that no viable cells were present on the meshes and most fibroblasts

remained in the skin specimens after mesh removal. Future studies will in-

clude a viability assay, such as the LIVE/DEAD protocol, performed on the

meshes to investigate the potential presence of dead cells. In addition quan-

titative approaches to compare the number of cells left into the specimens

after mesh removal will be designed and performed.

Figure 6.10: H & E histology image of the superficial de-epidermised woundwound model. Scale bar 20 µm

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Figure 6.11: Photographs of (a) skin specimens; (b) electrospun meshes afterMTT assay. No transfer of fibroblast cells occurred from the skin specimens onto

the meshes. The wound model was developed to mimic a superficialde-epidermized wound, with fibroblasts spread through the dermis, underneath the

BM.

6.2.3 Deep wound

The deep wound model was developed to represent a deep wound that lost

the entire epidermis, including the BM. The papillary layer of the dermis

constitutes the outermost layer of the model, as shown on the H & E his-

tology image (Figure 6.12). The fibroblasts that were seeded on the skin

specimens (dark purple spots on Figure 6.12) prevalently proliferated on the

surface and through the papillary dermis.

Figure 6.13 shows the meshes and skin specimens after the MTT assay. A

dark purple circle was obtained on the control specimen, onto which no mesh

was placed. The specimens incubated with the untreated PS, ppAAc and

ppAAm meshes showed very pale purple rings, while the specimens in con-

tact with ppCo and ppOct exhibited a more intense purple circle (Figure

6.13a), suggesting that the latter specimens retained the highest number of

viable cells.

All the meshes showed a purple ring, corresponding to viable fibroblasts that

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Figure 6.12: H & E histology image of the deep wound wound model. Scale bar20 µm

transferred from the skin model. The ppAAc and ppAAm meshes exhibited

the most intense purple rings, followed by the ppCo mesh; the ppOct coating

and the untreated PS meshes showed rings of lower intensity, not entirely

filled by the purple colour, but rather stained in the form of small purple

patches (Figure 6.13b).

As mentioned in the previous section, with the current experimental approach

only qualitative conclusions can be drawn by comparing the size, shape and

intensity of the purple rings on meshes and skin specimens. The obtained re-

sults indicate that a most significant transfer of viable cells occurred onto the

ppAAm and ppAAc meshes where full and dark purple rings were observed.

The ppOct chemistry induced the least transfer since the skin specimen ex-

hibited a dark and full purple ring and the mesh had small and light purple

patches. Further experiments need to be performed to investigate the pres-

ence of dead cells onto the meshes and to compare the number of viable cells

that transferred and those that remained into the specimens.

Discussion Four wound models were developed to reproduce wounds at

different depths and degree of severity. The plasma coated meshes were

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Figure 6.13: Photographs of (a) skin specimens; (b) electrospun meshes afterMTT assay. Transfer of fibroblast cells from the skin specimens onto the meshesoccurred. The wound model was developed to mimic a deep wound, with loss of

BM.

tested on the models to investigate the mechanisms of skin cells transfer.

In the presence of a superficial partially de-epidermized wound, where layers

of proliferating and differentiating keratinocytes are present on the top of

the basement membrane (BM) of the skin, none of the plasma coatings was

found to induce cell removal from the skin grafts. It was noticed that the

ppAAc and ppAAm meshes left fibrous layers attached to the cells at the

bottom of the wells. This suggests that the keratinocyte cells were strongly

bound within the culture and tended to proliferate throughout the outermost

layers of the ppAAc and ppAAm coated meshes. These results confirm the

findings obtained from the keratinocyte transfer studies from well plates and

are also supported by studies found in the literature [106, 107, 220], where

keratinocytes in solution culture were found to preferentially adhere onto

ppAAc and ppAAm coated samples, compared to the ppOct coating.

The second wound model was designed to reproduce a superficial wound that

lost stratum corneum and epidermis but maintained the BM. In this model

the meshes were in direct contact with the BM of the skin onto which no

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cells were present (Figure 6.10) as the fibroblasts were seeded and cultured

beneath the BM. The BM of the skin is a thin, fibrous, non-cellular region of

tissue that separates the epidermis from the underlying dermis. The major

molecular constituents of the BM are proteins including collagen IV, laminin-

entactin/nidogen complexes, and proteoglycans. In a superficial wound, the

integrity of the BM has to be maintained to avoid the exposure of deeper

tissues [117, 221].

Results showed that none of the tested plasma coatings induced the trans-

fer of viable cells from the skin specimens. This suggests the the BM was

kept intact after mesh removal and prevented the fibroblasts proliferating

throughout the dermis to be removed by the meshes. However, a more quan-

titative approach needs to be undertaken to compare the number of viable

cells cells left in the specimens after mesh removal.

In the third model, where the wound was deep and the BM compromised,

the ppAAc and ppAAm coatings were found to induce the highest removal

of viable fibroblasts from the skin models, while the ppOct coated and the

untreated PS meshes showed a low proportion of viable cells transferred.

Results suggest that the ppOct coating could be a suitable candidate for

developing dressings designed to interact with open deep wounds, where the

fibroblasts are exposed and cell removal has to be prevented. These results

support the findings obtained in the cell transfer studies performed on the

TCPS well plates.

6.2.4 3-Dimensional deep infected wound

The 3-dimensional deep infected wound model was developed by co-culturing

fibroblasts and keratinocytes in ∼ 5mm thick de-epidermized skin. After 14

day culture a thermal burn was generated on the specimens and the wound

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Figure 6.14: Histology images of the 3-dimensional wound model: (a) H & Estained section of the skin composite, before generating the burn. Scale bar 50µm; (b-e) Gram stained tissue sections, showing the progressive development ofthe model, from (b) epidermis formation; (c) thermal burn; (d) P.aeruginosabiofilm formation. Scale bar 100 µm; (e) magnification over the bacteria cells

forming the biofilm. Scale bar 20 µm.

was infected with P.aeruginosa. Figure 6.14 shows the histology images of

the 3-dimensional wound model. Figure 6.14a was obtained after H & E

staining the tissue section. The three characteristic layers of the human

skin can be visualised: the dermis, composed of fine and loosely arranged

collagen fibres in the uppermost layer (papillary dermis) and dense irregu-

lar connective tissue featuring densely packed collagen fibres in the bottom

layer (reticular dermis); the epidermis is the dark purple intermediate layer

of the skin composed of proliferating and differentiating keratinocytes; the

stratum corneum is the outermost layer of the epidermis, in bright pink,

consisting of flattened dead cells. On the gram stained histology section

(Figure 6.14b), the same layered organisation of the skin specimen can be

recognised. The thermal burn caused the loss of the stratum corneum and

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epidermis, as shown in Figure 6.14c. The dermis is exposed with irregular

edges. After seeding the P.aeruginosa culture on the burn for 18 hours, a

biofilm formed on the surface of the exposed dermis. The biofilm is shown in

Figure 6.14d, corresponding to the dark irregular layer on the surface of the

tissue. A magnification image of the infected specimen is provided in Figure

6.14e, where single rod-shaped P.aeruginosa cells can be identified within

the biofilm. The bacteria that were present on the skin specimens after 18

hour incubation were quantified at 10mm9 ± 5*10mm8 bacteria/g of tissue.

The untreated PS and plasma coated meshes were positioned on the infected

skin specimens and incubated for 1 hour. To ensure that the fibroblasts

and bacterial cells that transferred on the meshes could be easily identified,

confocal images of fibroblasts and P.aeruginosa cells attached onto separate

PS meshes were compared (Figure 6.15). Keratinocytes could not trans-

fer on the meshes because the burn of the specimens caused the removal of

the entire epidermis as shown by the histology study (Figure 6.14c). The

confocal images showed that fibroblasts and P.aeruginosa cells could be dis-

tinguished given the significant difference of nucleus size. Fibroblast nuclei

(Figure 6.15a) are about 10 µm wide, with a circular shape, while bacterial

nuclei (Figure 6.15b) are significantly smaller (about 1 µm in length) and

are rod-shaped.

The meshes that came in contact with the infected specimens were PI stained

and confocal z-stack images were acquired to quantify the number of mam-

malian and bacterial cells that transferred from the wound model. No fibrob-

lasts were found, while the number of bacterial cells that were counted on

each mesh are reported on Figure 6.16. In each of the three experiments, the

ppAAm coated mesh was found to have the highest number of bacterial cells

adhered compared to the untreated PS as well as the other tested chemistries.

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Figure 6.15: PI stained (a) fibroblast and (b) P.aeruginosa cells attached ontoPS meshes. Scale bar 20 µm

The first experiment resulted in 40086 ± 14682 bacteria/mm2 on the ppAAm

compared to the untreated PS which showed 24427 ± 9218 bacteria/mm2

(statistical significant difference for p < 0.01). The number of bacteria found

on the ppAAc (15622 ± 7608), ppCo (21451 ± 14882) and ppOct (24239

± 8327) were not statistically significantly different from the untreated PS

mesh. The second experiment exhibited a similar trend, with the ppAAm

mesh showing the highest number of bacteria/mm2 (30608 ± 15718) com-

pared to the uncoated mesh (17429 ± 5040) and the other plasma coatings

(ppAAc, 18823 ± 10617; ppCo, 15178 ± 7082; and ppOct, 23702 ± 13300).

In the last experiment the difference between the ppAAm and the untreated

meshes was statistically significant with p < 0.0001 (ppAAm, 44676 ± 14808;

and PS, 7471 ± 4543 bacteria/mm2); compared to the PS mesh, ppAAc and

ppOct showed a higher number of bacteria/mm2 (20405 ± 9480 and 27423

± 10978 respectively), while the ppCo coating was not significantly different

(10816 ± 8101).

The second part of the experiment consisted in recovering and quantifying

the viable bacterial cells that did not transfer onto the meshes and remained

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Figure 6.16: Graph showing the number of P.aeruginosa cells counted on themeshes on each of the three experiments.

on the skin specimens. Figure 6.17 shows the results expressed in the Log10

of the Colony Forming Units (CFU) recovered from the skin specimens per

gram of tissue. On the graph the individual data from each of the three

experiments are reported. The experiments show a similar trend, with the

lowest number of bacteria left on the wound model by the ppAAm mesh.

In experiment 3 this result is particularly evident, with the Log10(CFU/g)

calculated on the specimen from the ppAAm being 6.7 in comparison to

all the other specimens that had range between 8.9 and 9.7. The high-

est number of CFU was found left on the specimen corresponding to the

ppAAc mesh, with a Log10(CFU/g) = 9.7, followed by untreated PS and

ppCo (Log10(CFU/g)= 9.4) and ppOct (Log10(CFU/g) = 8.9). Experiments

1 and 2 resulted in a Log10(CFU/g) of 7.9 and 7.8 respectively on the spec-

imen from the ppAAm meshes, while the other specimens recorded higher

values, between 8.2 and 9.4 in both experiments. In experiment 1 and 2 most

bacteria (Log10(CFU/g) = 9.4) were found on the specimens onto which the

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Figure 6.17: Graph showing the quantification of P.aeruginosa cells remainedon the 3D wound model specimens after the removal of the plasma coated meshes

and the PS control mesh.

untreated PS meshes were placed, followed by ppOct (Log10(CFU/g) = 8.8),

and ppAAc and ppCo (Log10(CFU/g) = 8.2-8.6).

Discussion The infected wound model was developed to investigate the

transfer onto the plasma coated meshes of bacterial cells (P.aeruginosa) when

co-cultured with fibroblasts and keratinocytes. The protocol to co-culture

skin cells and bacteria in skin grafts to produce this model was originally

developed by Shepherd et al. to study any cutaneous invasive bacterial or

fungal infections [117, 133]. Other authors co-cultured bacteria and cells

to create human skin models for wound dressing testing, but most of these

studies do not involve the complete reconstruction of the layered structure

of the skin and the generation of a dermal injury before seeding the infection

[222–224]. The model used in the present work shares many of the properties

of normal skin such as a well-differentiated keratinocyte layer, a convoluted

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epidermaldermal junction retaining a basement membrane, and a fibroblast

populated dermis, and is histologically similar to human skin [117]. The

generation of the thermal burn on the model prior to seeding the bacterial

culture allowed to closely reproduce the conditions of a human dermal wound

with entire loss of epidermal layers. The number bacteria that was quantified

on the skin specimens prior to positioning the meshes (109 ± 5x108 CFU/g

of tissue) reproduced the conditions of an infected wound. In fact, the litera-

ture reports that acute or chronic wound infection exists when the microbial

load is >105 CFU/g of tissue [225].

The analysis of the meshes after exposure to the infected wound model

showed that no fibroblasts transfer occur on any of the meshes. This re-

sult could be likely due to the burning procedure that can have compromised

the viability of the fibroblast cells present in the outermost layer of the der-

mis that came in contact with the meshes.

The ppAAm coating induced the highest transfer of P.aeruginosa cells. In

parallel, the quantification of the viable bacteria left on the skin specimens af-

ter mesh removal showed that the skin specimen corresponding to the ppAAm

coated mesh had the lowest number of bacteria left. These results confirmed

the findings described in chapter 5 (section 5.3) where the ppAAm coated

meshes tested with an E.coli agar culture were shown to induce the highest

attachment of bacterial cells.

The number of bacteria that transferred from the infected wound model on

the ppAAc and ppCo meshes was similar to the untreated PS mesh, while

the ppOct showed a slightly higher attraction towards the bacterial cells,

reproducing the same trend that was reported in chapter 5 (section 5.3).

Although the number of bacteria left on the skin by the ppAAm mesh was

the lowest compared to the other meshes, this number was not low enough

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to prevent or disrupt the infection (<105 CFU/g). These results indicate

that the ppAAm coating has the potential for developing an attractive sur-

face that can clean the wound bed from bacterial contamination. However,

future studies will be performed to determine whether a reduction of the

bacterial load significant enough to prevent infection can be achieved by, for

instance, leaving the mesh on the infected tissues for a longer period of time.

Alternatively, the periodic replacement of clean ppAAm coated meshes can

be tested to evaluate the possibility of cleaning the wound bed through repet-

itive application of the mesh.

It is important to underline that the high variability that was obtained in the

quantification of the number of bacteria transferred onto the meshes and left

onto the skin model after mesh removal is inevitable given the complexity of

the experiments themselves, which include a combination of steps that could

partially affect bacterial viability.

6.3 Conclusions

To fabricate wound dressings that prevent wound reopening upon dressing

removal, the mechanisms of transfer of skin cells onto electrospun meshes

with different surface chemistries were investigated, by exposing the meshes

to a confluent culture of the cells at the bottom of well plates.

The ppOct coating resulted with the minor transfer of fibroblasts onto the

fibres, while ppAAc and ppAAm showed a significant cell removal from the

culture. These results suggest the possibility of deposing the ppOct coating

onto those wound dressings that are designed for deep wounds, where the

fibroblasts are directly exposed and their ingrowth into the dressings need to

be prevented. To assess this possibility, the coated meshes were exposed to

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a model of a deep wound, obtained by culturing human dermal fibroblasts

into a de-epidermized skin graft. The ppOct mesh was showed to induce

the minor degree of cell removal from the wound model, thus confirming the

suitability of the coating for a dressing capable of preventing fibroblasts in-

growth.

Other wound models were developed for testing the coatings onto more su-

perficial wounds, where the basement membrane is still present as well as part

of the epidermis made of keratinocyte cells. None of the coating was found to

be disruptive towards the basement membrane of the skin. The ppAAc and

ppAAm coated meshes remained partially adhered to the kertinocyte culture

upon mesh removal, thus suggesting that those coatings could be used for

dressings designed for superficial wounds, to bind to the outermost layer of

the wound bed for a period of time in order to encourage the proliferation of

keratinocytes and restoration of the epidermis.

The study on the P.aeruginosa transfer from the three-dimensional infected

wound model onto the plasma polymerised meshes showed that ppAAm in-

duced the highest removal of bacterial cells from the wound bed. This result

supports the findings obtained from the in vitro studies reported in chapter

5 and encourages the possibility of using the ppAAm coating as an ”attract

and trap” surface for the bacteria, to clean up the wound bed.

One significant contribution of the present work consists in adapting and op-

timising the various protocols of tissue-engineered human skin available in the

literature [137, 139] to design and develop innovative models of wounds at dif-

ferent depths (superficial partially de-epidermized; superficial de-epidermized;

and deep). In addition, the study of skin cells and bacterial transfer from

tissue engineered skin models onto and into electrospun meshes has not been

previously reported in the literature. The results suggest that the surface

186 / 223


chemistry of wound dressings could be specifically tailored to treat different

types of wound, simultaneously preventing skin cell ingrowth and controlling

the bacterial load.

187 / 223


188 / 223

Chapter 7

Conclusions

Contents7.1 Conclusions . . . . . . . . . . . . . . . . . . . . . . 189

7.2 Outlook . . . . . . . . . . . . . . . . . . . . . . . . 191

7.1 Conclusions

This thesis provided new insights into some of the key issues associated to

chronic wound management. The mechanisms of interaction of skin cells and

wound bacteria with electrospun materials were investigated and key prop-

erties of wound dressings that can be tuned to control wound healing and

prevent infection were identified.

The average fibre diameter of PS electrospun meshes was found to influence

the ability of three bacterial species (E.coli, P.aeruginosa and S.aureus) to

proliferate and colonise the fibrous substrate. Meshes with an average fibre

diameter close to bacterial size were found to offer the best support for bac-

terial adhesion and spreading, constituting a scaffold that bacteria use as a

189

7. Conclusions

framework for forming colonies. For rod shape elongated cells (E. coli and

P. aeruginosa), fibre diameters smaller than the bacterial length resulted in

most cells to wrap around each fibre, thus limiting the ability of bacteria to

easily create bridges across fibres and form colonies. These bacteria exhib-

ited similar behaviour, colonising preferentially the 1 µm diameter meshes.

Round S. aureus cells showed the highest proliferation throughout the 500

nm diameter fibrous substrates; in the presence of bigger fibres, S. aureus

cells preferentially adhered on the fibre surface, without spreading through-

out the mesh.

The influence of the surface chemistry of plasma modified PS meshes on

bacterial behaviour was also explored. A combination of surface chemistry

features, including surface wettability, charge and functional groups, were

shown to affect the capacity of bacteria to transfer and attach onto the fibres

and proliferate within the mesh. The allylamine plasma coating, that is an

amine rich, hydrophilic positively charged film induced the highest propor-

tion of E.coli and P.aeruginosa cells transferred from an underlying culture

onto and within the coated meshes, in comparison to untrated materials and

the other coatings (ppAAc, ppCo, and ppOct).

Fibre surface chemistry was also found to affect the capacity of skin cells to

transfer and attach onto the electrospun fibres. The ppOct mesh was found

to induce the least degree of cell removal from the tissue engineered model of

a deep wound, suggesting the possibility of deposing this coating onto those

wounds where the fibroblasts are directly exposed and their ingrowth into

the dressings needs to be prevented to avoid wound reopening upon dressing

removal.

Other wound models were developed for testing the plasma coatings onto

more superficial wounds, where the basement membrane and part of the epi-

190 / 223

7. Conclusions

dermis are still intact. The ppAAc and ppAAm coated meshes remained

partially adhered to the kertinocyte culture upon mesh removal, thus sug-

gesting that those coatings could be used for dressings designed for superficial

wounds, to be integrated with the wound bed for encouraging and supporting

the epidermis re-growth.

The presented results showed the possibility of combining controlled fibre

size and surface chemistry to control initial bacterial attachment and spread-

ing into electrospun materials. The ppAAm coating could be a potential

candidate for an ”attract and trap” dressing capable of attracting bacteria

from the wound bed, while controlled fibre size could allow to trap the bac-

teria within the fibrous network. Fibre surface chemistry was also found to

be a key parameter that can be strategically controlled to prevent skin cell

transfer and ingrowth in the dressing.

7.2 Outlook

Overall, the results reported in this thesis highly encourage further studies

on the responses of wound bacteria to electrospun materials. Fibre surface

modification combined with controlled fibre size was found to be a potential

strategy to develop an ”attract and trap” wound dressing capable of attract-

ing the bacteria from the wound bed and trapping them within the fibrous

network. Further studies are necessary to assess the potential efficacy of this

solution. Different bacterial species need to be included and long term exper-

iments should be performed to investigate the possibility of using fibre size

and surface chemistry as tools to control not only initial bacterial attachment

but also biofilm formation and infection development.

Complex three dimensional fibrous meshes could be fabricated and differ-

191 / 223

7. Conclusions

ent strategies could be combined within the same dressing. For instance,

an antibacterial coating or drug release approach could be integrated in the

”attract and trap” dressing to kill the viable bacterial cells that transfer from

the wound bed and remain trapped within the fibres.

The tissue engineered skin models of wounds offered extraordinary tools to

evaluate the interactions of fibroblasts and keratinocytes with the plasma

coated meshes. The same models could be used to test complex wound

dressings obtained by combining the required fibre morphology and surface

chemistry features necessary to reduce the bacterial load in the wound bed

while preventing skin cell ingrowth. For instance, a layered dressings com-

posed of ppAAm coated fibres for attracting the bacteria and ppOct coated

fibres for preventing skin cell ingrowth can be fabricated and tested on the

model of a deep infected wound to evaluate the possibility of achieving a

simultaneous control over bacterial and skin cell behaviour. In addition, the

wound models could be adapted to test the capacity of the plasma modified

meshes to encourage skin cell proliferation in the wound bed and wound clo-

sure. To this end, the models could be further improved to better mimic a

wound environment by adding other molecules that play key roles in wound

healing, including cytokines and macrophages. The culture conditions will

need to be explored and optimised to achieve a correct balance between skin

cells, bacteria and inflammatory cells. The result would constitute a wound-

on-chip platform that could partially reproduce the inflammatory conditions

of real chronic wounds.

192 / 223

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Appendix

Figure 7.1: Example of the imageJ code that was developed for an automaticselection and counting of the particles present on the confocal images of the

meshes tested on the infected wound model. Each particle counted by the softwarecorresponds to a PI stained bacterial cell. The code implements the z-projection

of each image, threshold adjustment and particle counting. The commentsdescribing each function of the code are shown in green.

217

Appendix

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Research achievements

Reviewed publications

M. Abrigo, P. Kingshott, S. L. McArthur, ”Bacterial response to differ-

ent surface chemistries fabricated by plasma polymerization on electrospun

nanofibers,” Bionterphases, vol. 10, no. 4, pp. 04A3011-9, 2015.

M. Abrigo, P. Kingshott, S. L. McArthur, ”Electrospun Polystyrene Fiber

Diameter Influencing Bacterial Attachment, Proliferation and Growth,” ACS

Appl. Mater. Interfaces., vol. 7, no. 14, pp. 7644-52, 2015.

M. Abrigo, S. L. McArthur, P. Kingshott, ”Electrospun Nanofibers as

Dressings for Chronic Wound Care: Advances, Challenges, and Future Prospects,”

Macromol Biosci., vol. 14, no. 6, pp. 772-92, 2014.

Conference presentations

Poster presentation at ANN Nanotechnology Jun 2015

Entrepreneurship Workshop

Gold Coast, Australia

219

Research Achievements

Oral presentation at ISSIB/ASBTE Conference Apr 2015

Sydney, Australia

Poster presentation at Nanolytica Symposium Feb 2015

Melbourne, Australia

Oral presentation at PacSurf Conference Dec 2014

Hawaii, USA

Oral presentation at AVS Conference. Nov 2014

Baltimore, MD, USA

Oral presentation at NanoBIO Conference July 2014

Brisbane, Australia

Poster presentation at ASBTE Conference Apr 2014

Lorne, Australia

Oral presentation at Australian Colloid & Surface Science Feb 2014

Ballarat, Australia

3 Minute Thesis Competition July 2013


Poster presentation at AVS Conference Nov 2013

Long Beach, CA, USA

220 / 223


Attendance to AusMedTech Conference. May 2013


Poster and oral presentation at Swinburne Post Graduate Nov 2012

Conference


Attendance to Swinburne Living Research Conference Sept 2012


Awards

1st prize for Startup pitch presentation at ANN Jun 2015

Nanotechnology Entrepreneurship Workshop (1000$)

Gold Coast, Australia

ISSIB/ASBTE Conference travel grant (270$) Apr 2015

& runner up in best student oral presentation

Sydney, Australia

First place poster prize at Nanolytica symposium (800$) Feb 2015


ANFF Travel grant for NanoBio Conference (500$) July 2014

Brisbane, Australia

221 / 223


First prize poster presentation at AVS Conference Nov 2013

(500$)

Long Beach, California, USA

First prize at the Faculty Final of the 3 Minute Thesis July 2013

Competition (1000$)

Swinburne University of Technology, Australia

First prize for 1 minute thesis presentation (250$) Nov 2012

Swinburne University of Technology, Australia

Online media releases

Smart dressings interview x

Youtube Media - 23 Sept 2015

Science magazine news article x

IFLScience - 7 Sept 2015

Come funziona il cerotto che accelera guarigione x

La Stampa - 3 Sept 2015

A new type of bandage will draw out bacteria and speed up healing x

Science Alert - 2 Sept 2015

Smart dressings speed healing of chronic wounds x

Swinburne Media Center - 21 Aug 2015

Swinburne story x

Swinburne Media Center - 14 Aug 2015

222 / 223

https://www.youtube.com/watch?v=UmaH3xDy7L0

http://www.iflscience.com/health-and-medicine/bandage-sucks-bacteria-and-rips-them

http://www.lastampa.it/2015/09/03/scienza/benessere/come-funziona-il-cerotto-che-accelera-guarigione-Q0bW629r4wjsFRYkUpgbLN/pagina.html

http://www.sciencealert.com/a-new-type-of-bandage-can-draw-out-bacteria-and-speed-up-wound-healing

http://www.swinburne.edu.au/media-centre/news/2015/08/smart-dressings-speed-healing-of-chronic-wounds-.html

http://swinburne.tumblr.com/post/126634227969/my-research-has-been-challenging-demanding-at


Biomedical Picture of the Day (BPoD) news article x

Biomedical picture of the day - 25 Nov 2014

A Band-Aid that could suck bugs out of your wound x

Science mag - 12 Nov 2014

223 / 223

http://www.bpod.mrc.ac.uk/archive/2014/11/25

http://news.sciencemag.org/health/2014/11/band-aid-could-suck-bugs-out-your-wound

Documents

Development of electrospun dressings for infected wounds