BIO-FUNCTIONALIZATION OF ELECTROSPUN NANOFIBRESCAFFOLDS FOR
CELL CULTURE APPLICATIONSCHUA KIAN NGIAP
B. Eng. (Hons.), NUS

A THESIS SUBMITTEDFOR THE DEGREE OF DOCTOR OF PHILOSOPHYGRADUATE PROGRAMME IN BIOENGINEERINGNATIONAL UNIVERSITY OF SINGAPORE
2006

Acknowledgements

ACKNOWLEDGMENTSFirst of all, I would like to thank my project supervisors Professor Seeram
Ramakrishna, Professor Kam W. Leong and Assistant Professor Hai-Quan Mao for
their constant support and guidance, and for all the opportunities that they have given
me in my education and research. I have learnt to become a better researcher and also
a better person. A simple “thank you” will not be enough to express my gratitude.
I would like to thank all my colleagues at the Tissue and TherapeuticEngineering Laboratory, Division of Johns Hopkins in Singapore for all the assistancethat they provide for the completion of this thesis. My special thanks to Dr. Chou
Chai, Dr. Hong-Fang Lu, Dr. Xue-Song Jiang and Dr. Chao Yin for imparting mewith their skills and knowledge. My sincere appreciation is also given to Mr. PengChou Lee, Ms. Yen-Ni Tang, Mr. Wei-Seng Lim, Ms. Chai-Hoon Quek, Dr. Peng-Chi
Zhang, Mr. Justin Gorham, Ms. Ai-Cheng Tan and Mr. Teck-Jin Tan for all the
precious technical support that they have provided through these years.
I would also like to thank all my colleagues in the Nanobioengineering
Laboratory, NUSNNI and Graduate Programme in Bioengineering. I express mydeepest gratitude to Dr. Kazutoshi Fujihara, Dr. Joon-Kin Yong, Ms. Satinderpal Kaur,Ms. Yan-Ping Wang, Mr. Daniel Wong, Mr. Ramakrishnan Ramaseshan, Mr. ChunWai Ng, Ms. Puay-Joo Low, Ms. Siew-Teng Yeo and Ms. Soo-Hoon Pang for all theassistance that they have given me in many different ways.
Finally, I am greatly indebted to my family for their constant support andencouragement throughout these long thesis years.

ii

Table of Contents

TABLE OF CONTENTS
Title

i

Acknowledgementsii

Table of Contentsiii

Summary

viii

List of Publications

xi

List of Figuresxii

List of Tables

xvi

Chapter 1

General Overview

1.1

Background1

1.2

Thesis Objectives

3

1.3

Thesis Scope

4

Chapter 2
2.1

Literature Review
Electrospun Nanofibers

6

2.1.1

Principles and Mechanisms

7

2.1.2

Parameters that Control the Electrospinning Process

8

2.1.2.1

Effect of Polymer Concentration in Electrospinning Solution 92.1.2.2

Effect of Ionic Additives in Electrospinning Solution

10

2.1.2.3

Collector Design11

2.1.2.4

Spinneret Design11

2.1.2.5

Other Miscellaneous Parameters

12

2.1.3

Electrospun Nanofibers in Cell Culture Applications

13

2.1.4

Nanofiber Modification for Cell Culture Applications

15

2.1.4.1

Doping of Bioactive Molecules

16

2.1.4.2

Nanofiber Surface Modification

17

2.2

Biomaterials Design for Primary Hepatocyte Culture

18

2.2.1

Hepatocyte Function Maintenance through Spheroid Formation

21

2.2.2

Hepatocyte Cultures on Galactosylated Scaffolds

23

2.2.3

Galactosylated Nanofiber Scaffolds for Hepatocyte Cultures

24

2.3

Biomaterials Design for Ex Vivo HSPC Expansion

24
iii

Table of Contents

2.3.1

The Hematopoietic System

26

2.3.2

Hematopoietic Stem/Progenitor Cell Sources

27

2.3.3

Hematopoietic Stem/Progenitor Cell Characterization Techniques 28

2.3.4

Hematopoietic Stem/Progenitor Cell Expansion Cultures30

2.3.4.1

HSPC Cultures with Stromal Cells or Conditioned Medium

30

2.3.4.2

HSPC Cultures with Human Recombinant Cytokines

32

2.3.4.3

HSPC Cultures on Scaffolds

34

2.4
Chapter 3

Concluding Remarks

35

Stable Immobilization of Hepatocyte Spheroids on Galactosylated
Nanofiber Scaffolds for Liver Cell Culture

3.1

Summary37

3.2

Introduction

38

3.3

Experimental Methods

40

3.3.1

Fabrication of PCLEEP Nanofiber Scaffolds

40

3.3.1.1

Surface Grafting of Scaffolds with Poly(acrylic acid)41

3.3.1.2

Galactosylation of Poly(acrylic acid) Grafted Scaffolds

42

3.3.2

Hepatocyte Culture and Assays

42

3.3.2.1

Hepatocytes Isolation

42

3.3.2.2

Hepatocyte Attachment Assay42

3.3.2.3

Hepatocyte Culture Maintenance43

3.3.2.4

Albumin Secretion Assay43

3.3.2.5

Urea Synthesis Assay44

3.3.2.6

Cytochrome P450 Activity Assay44

3.3.2.7

Preparation for Scanning Electron Microscopy

45

3.3.3
3.4

Statistical Analysis

46

Experimental Results

46

3.4.1

Optimization of PCLEEP Electrospinning

46

3.4.2

Optimization of Scaffold Galactosylation Process

47

3.4.3

Hepatocyte Functional Maintenance

48

3.4.4

Hepatocyte Morphological Changes

52

3.5

Discussion

57

3.6

Concluding Remarks

60

iv

Table of Contents

Chapter 4

Hepatocyte Cytochrome P450 Inducing Dual-Functional Nanofiber
Scaffolds for Hepatocyte Culture

4.1

Summary61

4.2

Introduction

62

4.3

Experimental Methods

65

4.3.1

Fabrication of Dual-Functional Nanofiber Scaffolds

65

4.3.1.1

Electrospinning of Undoped Nanofiber Mesh

65

4.3.1.2

Poly(acrylic acid) Grafting of Undoped Nanofiber Mesh

65

4.3.1.3

Electrospinning of 3-Mc Loaded Nanofiber Mesh

65

4.3.1.4

Galactosylation of Composite Nanofiber Scaffold

66

4.3.2

Hepatocyte Culture and Assays

66

4.3.2.1

Hepatocytes Isolation

66

4.3.2.2

Hepatocyte Attachment Assay66

4.3.2.3

Hepatocyte Culture Maintenance67

4.3.2.4

Cytochrome P450 Activity Assay67

4.3.2.5

Albumin Secretion Assay68

4.3.2.6

Transwell Cultures

68

4.3.3
4.4

Statistical Analysis
Experimental Results and Discussion

69
69

4.4.1

Dual-Functional Nanofiber Scaffold Characterization

69

4.4.2

Hepatocyte Attachment Efficiency

71

4.4.3

Cytochrome P450 Function

72

4.4.4

Albumin Synthesis Function

73

4.4.5

Mechanism of 3-Mc Transport from Nanofiber to Cell

74

4.5
Chapter 5Concluding Remarks

75

Aminated Nanofiber Scaffolds Enhance Adhesion and Expansion of
Human Umbilical Cord Blood Hematopoietic Stem/Progenitor Cells

5.1

Summary77

5.2

Introduction

78

5.3

Experimental Methods

80

5.3.1

Fabrication of PES Nanofiber Scaffolds

80

5.3.1.1

Surface Grafting of Scaffolds with Poly(acrylic acid)81

5.3.1.2

Amination of Poly(acrylic acid) Grafted Scaffolds

81vTable of Contents

5.3.1.3

Surface Analysis of PES Scaffolds

82

5.3.2

Hematopoietic Stem Cell Culture and Assays

82

5.3.2.1

Ex Vivo Hematopoietic Expansion Culture83

5.3.2.2

Flow Cytometry

83

5.3.2.3

Colony-Forming Cell Assay84

5.3.2.4

Preparation for Scanning Electron Microscopy

85

5.3.2.5

Preparation for Laser Scanning Confocal Microscopy

85

5.3.3
5.4

Statistical Analysis

85

Experimental Results

85

5.4.1

Modification of PES Substrates and Surface Characterization

85

5.4.2

Ex Vivo HSPC Expansion on Various PES Substrates

87

5.4.3

Colony-Forming Cell Assay Results

89

5.4.4

Expanded HSPC Surface Marker Expression

91

5.4.5

Imaging of Adherent Cells on Aminated Substrates

93

5.5

Discussion

96

5.6

Concluding Remarks

99

Chapter 6

Nanofiber Scaffolds Modified with Different Spacer-Length Amines
Modulate Hematopoietic Stem/Progenitor Cell Maintenance andProliferation Kinetics

6.1

Summary101

6.2

Introduction

102

6.3

Experimental Methods

103

6.3.1

Fabrication of PES Nanofiber Scaffolds

103

6.3.1.1

Surface Grafting of Scaffolds with Poly(acrylic acid)103

6.3.1.2

Amination of Poly(acrylic acid) Grafted Scaffolds

104

6.3.2

Hematopoietic Stem Cell Culture and Assays

104

6.3.2.1

Ex Vivo Hematopoietic Expansion Culture

105

6.3.2.2

Flow Cytometry

105

6.3.2.3

Preparation for Scanning Electron Microscopy

106

6.3.2.4

Colony-Forming Cell Assay106

6.3.2.5

Long-Term Culture-Initiating Cell Assay106

6.3.2.6

Mouse Engraftment Assay

106

6.3.3

Statistical Analysis

107
viTable of Contents

6.4

Experimental Results

107

6.4.1

Surface Characterization of Aminated Nanofiber Scaffolds

107

6.4.2

Ex Vivo HSPC Expansion on Aminated Nanofiber Scaffolds

110

6.4.3

Morphology of Adherent Cells on Aminated Scaffolds

112

6.4.4

HSPC Clonogenic Potential on Various Scaffolds

116

6.4.5

HSPC NOD/SCID Repopulation Potential on Various Scaffolds 118

6.5

Discussion

119

6.6

Concluding Remarks

122

Chapter 7

Adhesive Cell-Scaffold Interaction through Aminated Nanofiber
Scaffold Promotes Hematopoietic Stem/Progenitor Cell
Maintenance and Lineage Commitment

7.1

Summary123

7.2

Introduction

124

7.3

Experimental Methods

125

7.3.1

Fabrication of PES Nanofiber Scaffolds

7.3.1.1

Surface Amination of PES Nanofiber Scaffolds

125
125

7.3.2

Hematopoietic Stem Cell Culture and Assays

126

7.3.2.1

Ex Vivo Hematopoietic Expansion Culture

126

7.3.2.2

Cell Harvest

127

7.3.2.3

Flow Cytometry

127

7.3.2.4

Colony-Forming Cell Assay

128

7.3.3
7.4

Statistical Analysis

128

Experimental Results

128

7.4.1

Lineage Analysis of Adherent and Non-Adherent HSPCs

128

7.4.2

Clonogenic Differences of Adherent and Non-Adherent HSPCs

131

7.5

Discussion

133

7.6

Concluding Remarks

135

Conclusions

136

Chapter 8
Appendix

140

References

144

vii

Summary

SUMMARY
This thesis presents the studies of bio-functionalization of electrospun nanofibers,
which can serve as cell culture scaffolds that can promote cell-substrate interactions
and are bioactive in soliciting favorable cellular responses like cell adhesion, cellmorphological reorganization, cell differentiated functions or cell proliferation.
The general strategy of scaffold development involves nanofiber scaffold
fabrication via the electrospinning technique, followed by nanofiber biofunctionalization.

Thebio-functionalization

process

involves

theinitialfunctionalization of the nanofiber surface with carboxylic acid groups using UVinitiated poly(acrylic acid) grafting method. This is followed by conjugation of
bioactive molecules onto the functionalized nanofiber surfaces. We then tested theefficacy of this nanofiber bio-functionalization strategy on hepatocyte scaffold
cultures and hematopoietic stem cell expansion culture systems.
Through galactose bio-functionalization, we have developed galactosylated
nanofiber scaffolds that can support the hepatic functions (albumin secretion,ammonia removal and cytochrome P450 activity) of cultured primary hepatocytes.
Interestingly, the nanofiber topography and the surface-immobilized galactose ligand
synergistically enhance the hepatocyte-nanofiber interaction, and the galactosylated
nanofiber scaffolds exhibit the unique property of promoting hepatocyte aggregatesand cell infiltration within the mesh and around the fibers, forming an integrated
spheroid-nanofiber construct. Subsequently, we have also demonstrated that
hepatocyte cytochrome P450 activity enhancement can be brought about throughfurther 3-Mc bio-functionalization of this galactosylated nanofiber scaffold.
viii

Summary

Through amine molecule bio-functionalization, we have developed aminatednanofiber scaffolds that can support ex vivo hematopoietic stem / progenitor cell
(HSPC) expansion. We have shown that aminated nanofiber meshes supported a highdegree of cell adhesion, percentage of CD34+CD45+ cells and expansion of CFUGEMM forming progenitor cells. SEM imaging also revealed discrete colonies ofcells proliferating and interacting with the aminated nanofibers. In addition, we haveshown that nanofiber scaffolds immobilized with amine functional groups of different
carbon spacer chain lengths could further modulate HSPC proliferation and phenotype
maintenance, resulting in different HSPC proliferation kinetics, cell population
phenotypic expression, mouse engraftment potential and also colony-forming ability.
The adherent hematopoietic cell populations on the aminated nanofiber scaffolds also
showed enrichment of CD34+CD45+ cells compared with the non-adherent cell
population, and indicated significant commitment towards the myeloblast / monoblast
lineage, while the non-adherent population showed skewed commitment towards theerythroid lineage. These observations suggested the importance of nanofiber
topography and amino functional group mediated cell-scaffold interactions inregulating HSPC proliferation and self-renewal. In addition, they also highlight theimportance of cell-scaffold interactions as a new approach in modulating HSPC
multipotency maintenance and lineage commitment.
In conclusion, this thesis has:
(1) Presented a nanofiber bio-functionalized strategy to develop polymericnanofiber constructs that can serve as cell culture scaffolds.
(2) Demonstrated through primary hepatocyte cultures and HSPC expansion
cultures that these scaffolds can promote cell-substrate interactions and areix

Summary

bioactive in regulating cellular responses like cell adhesion, cellmorphological reorganization, cell differentiated functions, cell proliferationor cell phenotype maintenance.
(3) Demonstrated the synergistic effects that both nanofiber topography andsurface immobilized biochemical cues play in enhancing these cell-scaffold
interactions and regulation of cellular functions.

xList of Publications

LIST OF PUBLICATIONS
1.

Chua KN, Lim WS, Zhang PC, Lu HF, Wen J, Ramakrishna S, Leong KW, Mao
HQ. Stable Immobilization of Rat Hepatocyte Spheroids on Galactosylated
Nanofiber Scaffold. Biomaterials 2005; 26(15):2537-2547.

2.

Lu HF, Chua KN, Zhang PC, Lim WS, Ramakrishna S, Leong KW, Mao HQ.
Three-Dimensional Co-Culture of Rat Hepatocyte Spheroids and NIH/3T3
Fibroblasts Enhances Hepatocyte Functional Maintenance. Acta Biomaterialia
2005; 1(4):399-410.

3.

Luong-Van E, Grondahl L, Chua KN, Leong KW, Nurcombe V, Cool SM.
Controlled Release of Heparin from Poly(epsilon-caprolactone) Electrospun
Fibers. Biomaterials 2006; 27(9):2042-2050.

4.

Chua KN, Chai C, Lee PC, Tang YN, Ramakrishna S, Leong KW, Mao HQ.
Surface-Aminated Electrospun Nanofibers Enhance Adhesion and Expansion of
Human

Umbilical

Cord

Blood

Hematopoietic

Stem/Progenitor

Cells.

Biomaterials 2006; 27(36):6043-6051.

5.

Chua KN, Chai C, Lee PC, Ramakrishna S, Leong KW, Mao HQ. Functional
Nanofiber Scaffolds with Different Spacers Modulate Adhesion and Expansionof Cryopreserved Umbilical Cord Blood Hematopoietic Stem/Progenitor Cells.
Experimental Hematology (In Press).

xi

List of Figures

LIST OF FIGURES
Figure 1.1:

The cellular microenvironment.

Figure 2.1:

SEM images of fibers prepared by electrospinning of non-degradable2

and degradable polymers.

6

Figure 2.2:

Schematic illustration of an electrospinning setup.

7

Figure 2.3:

SEM images of electrospun PES with increasing concentrations in
dimethylsulfoxide solvent (w/w).

Figure 2.4:

9

SEM images of PCLEEP fibers co-electrospun with increasing
concentrations of R18 in PCLEEP (w/w).

10

Figure 2.5:

Interior modification of electrospun nanofiber scaffolds.

17

Figure 2.6:

Exterior modification of electrospun nanofiber scaffolds.

18

Figure 2.7:

Approaches to cellular therapies for the treatment of liver disease.

19

Figure 2.8:

Morphology of hepatocyte spheroids.

21

Figure 2.9:

Control of hematopoiesis in a bone marrow microenvironment.

27

Figure 2.10: Generic HSPC cultivation variants.
Figure 3.1:

30

Surface modification scheme for galactose conjugation to PCLEEP
nanofiber mesh.

39

Figure 3.2:

SEM characterization of PCLEEP nanofiber mesh.

46

Figure 3.3:

Effect of acrylic acid monomer concentration used for UV-initiatedgraft polymerization on the surface concentration of the graftedcarboxyl groups on the PCLEEP nanofiber mesh and spin-coated film
surface.

Figure 3.4:

Hepatocyte attachment on galactosylated and unmodified nanofiber
meshes and spin-coated films 3 h after seeding.

Figure 3.5:

48

49

Albumin secretion level of hepatocytes at various time points
normalized against the total number of attached cells.

50

xii

List of Figures

Figure 3.6:

Urea synthesis function of hepatocytes at various time points
normalized against the total number of attached cells.

Figure 3.7:

3-Mc induced P450 function of hepatocytes at various time points
normalized against the total number of attached cells.

Figure 3.8:

51

Hepatocyte spheroid adhesion on galactosylated scaffolds after 5 daysof culture.

Figure 3.9:5152

Morphology of hepatocytes at 3-h, 1-day and 3-days after seedingwhen cultured on different substrates.

54

Figure 3.10: SEM images of hepatocytes after 8 days of culture.

55

Figure 3.11: SEM images of freeze-fractured hepatocytes on Gal-nanomesh after 8days of culture.
Figure 4.1:

56

Electrospun galactosylated, 3-Mc loaded PCLEEP nanofiber scaffold
fabrication scheme.

64

Figure 4.2:

Scaffold condition illustration for transwell experiment.

68

Figure 4.3:

SEM images of electrospun PCLEEP nanofiber mesh layers.

70

Figure 4.4:

Hepatocyte attachment on various galactosylated 3-Mc loaded
composite nanofiber scaffolds (gnPCLEEP (0.0-8.0)% 3-Mc), single
layer galactosylated scaffolds (gnPCLEEP control) and TCPS control
3 h after cell seeding.

Figure 4.5:

Cytochrome P450 function of hepatocytes at various time points
normalized against the total number of attached cells.

Figure 4.6:

72

Albumin synthesis function of hepatocytes at various time points
normalized against the total number of attached cells.

Figure 4.7:

7173

Cytochrome P450 function of hepatocytes at various time points
normalized against the total number of attached cells. For gnPCLEEP
8% 3-Mc transwell condition, hepatocytes were cultured without
physical contact with the 8.0% 3-Mc mesh.

75

Figure 5.1:

PES scaffold surface modification scheme.

80

Figure 5.2:

SEM images of electrospun PES nanofiber mesh.

86
xiii

List of Figures

Figure 5.3:

Fold expansion of total nucleated cells and CD34+ cells following a
10-day culture of 600 human cord blood HSPCs on different
substrates.

Figure 5.4:

88

CFU counts generated after 14 days of culture, using the cells fromthe 10-day expansion cultures on various substrates and from theunexpanded HSPCs.

Figure 5.5:

Surface antigen expression on cells after 10-day ex vivo expansion ondifferent substrates.

Figure 5.6:

90

91

SEM images of human cord blood HSPCs after a 10-day expansion
culture on aminated PES nanofiber mesh and on aminated PES film at
various magnifications.

Figure 5.7:

Confocal laser microscopy images of human cord blood HSPCs aftera 10-day expansion culture on aminated PES nanofiber mesh.

Figure 6.1:

94

95

PES nanofiber scaffold amination scheme with different spacer chain
length amines.

103

Figure 6.2:

The XPS spectra of various modified PES nanofiber surfaces.

109

Figure 6.3:

Fold expansion of total nucleated cells and CD34+ cells following a
10-day culture of 600 human cord blood HSPCs on different
substrates.

Figure 6.4:

111

Representative FACS profiles and surface marker expression
summary of cells after 10-day ex vivo expansion on TCPS and EtDA,
BuDA and HeDA nanofiber scaffolds.

Figure 6.5:

SEM images of HSPCs after 3-day and 8-day cultures on PES BuDA
nanofiber mesh at various magnifications.

Figure 6.6:

112

114

SEM images of adherent cell colonies after 10 days of expansion on
PES EtDA, BuDA and HeDA nanofiber mesh at various
magnifications.

Figure 6.7:

115

CFU counts after 14 days of culture, using the cells from the 10-day
expansion cultures on various substrates and unexpanded HSPCs,
normalized to CFU per 100 initial unexpanded HSPCs.

117

xiv

List of Figures

Figure 6.8:

LTC-IC counts after 7 weeks of culture, using the cells from the 10day expansion cultures on various substrates and unexpanded HSPCs,
normalized to LTC-IC per 100 initial unexpanded HSPCs.

Figure 6.9:

117

Engraftment efficiency of human CD45+ cells in bone marrow of sublethally irradiated NOD/SCID mice transplanted with unexpanded
HSPCs, cells from the 10-day expansion cultures on various
substrates, and irradiated carrier cells alone.

Figure 7.1:

Image of a representative adherent cell colony formed on aminated(BuDA) nanofiber scaffold 10 days after ex vivo HSPC expansion.

Figure 7.2:

118

125

Representative FACS profiles of cells after 10-day ex vivo expansion
on TCPS, PES-BuDA, and non-adherent and adherent fractions fromPES-BuDA conditions.

Figure 7.3:

130

Surface marker expression summary of cells after 10-day ex vivo
expansion on TCPS, PES-BuDA, and non-adherent and adherent
fractions from PES-BuDA conditions.

Figure 7.4:

131

Specific CFU fractions after 14 days of culture, using the cells from
10-day expansion cultures on TCPS, PES-BuDA, and non-adherent
and adherent fractions from PES-BuDA conditions, normalized to

Figure 8.1:

CFU type per total CFU generated.

132

Synthesis scheme for AHG.

140

xv

List of Tables

LIST OF TABLES
Table 5.1:

Characterization of surfaces modified with various functional groups. 87

Table 6.1:

XPS elemental analysis of PES nanofiber surfaces modified withdifferent functional groups.

108

xvi

Chapter 1

CHAPTER ONE
General Overview
1.1

Background
Biomaterials play central roles in modern strategies in cell culture as designablebiophysical and biochemical milieus that direct cellular behavior and function [1,2].
In most approaches, the Biomaterial is engineered into a scaffold which provides aniche for cells to proliferate and differentiate.
The intended uses for scaffold-based cell cultures are vast: In some applications,the cells develop into tissues that are suitable for implantation, or applied as part of acell-based artificial organ [2-6] (e.g. bioartificial pancreas, bioartificial liver). In otherapplications, the cells are harvested after being expanded on proliferation-inductive
scaffolds, for use in cell-based therapies [2-6] (e.g. stem cell therapy). In all cases, ascaffold that can interact and influence the cellular behavior is the crucial component.
The success of scaffold-based cell cultures largely depends on the optimal eventsof attachment, proliferation, differentiation, and phenotypic maintenance, which inturn are governed by a host of signals provided by the cell-scaffold microenvironment.
These signals include: (1) homotypic / heterotypic cell-cell interaction; (2) soluble
signaling molecules; and (3) cell-substrate interaction signals which consists ofsubstrate-bound signaling molecules, scaffold topographical cues and scaffold
biomechanical properties (Fig. 1.1). Therefore, an ideal scaffold culture system should
include all these interactive components [1,2,7,8].

1

Chapter 1

Figure 1.1: The cellular microenvironment. The behavior of individual cells and the
dynamic state of multicellular tissues is regulated by intricate reciprocal molecularinteractions between cells and their surroundings.

In recent years, scaffolds based on electrospun nanofibers have been investigatedintensively [9-27]. This is largely due to the unique nano-topographical cues that thenanofiber scaffold provides as compared to 2-dimensional substrates, micro-porousand micro-fiber scaffolds and hydrogels traditionally used in cell cultures. Indeed,morphological and cytoskeletal reorganization of cells induced by the nanofiber
topographical cues has been clearly demonstrated in many literatures [14-27].
Though several nanofiber scaffolds of unique topographical textures (aligned
fiber scaffold, multilayered fiber scaffold, etc.) have been designed throughmanipulation of the electrospinning process [9-13,28-29], the nanofiber scaffolds usedin current literature are mainly pristine and lack of substrate-bound signaling
molecules [14-27]. In contrast, abundant research on the traditional film, micro-fiberor gel scaffolds have shown that scaffold functionalization (surface immobilization or
entrapment) with bioactive molecules (e.g. proteins, peptides, drugs, simple chemicalgroups, etc.) are necessary in soliciting favorable cellular responses like cell adhesion
2Chapter 1

and proliferation responses [30-36]; and electrospun nanofiber scaffolds should not be
the exception. Therefore, the design of a nanofiber scaffold modified with at least one
bioactive molecule would be important in enhancing cell-substrate interaction, withthe eventual goal of mimicking the cell’s native microenvironment.
Due to the similarities in the materials used, the common modification methods
for bio-functionalizing the traditional scaffolds can also be directly imported tomodify the nanofibers. In this thesis, we present a comprehensive approach tosystematically incorporate various types of biochemical cues into nanofiber scaffolds
that are critical for hepatocyte functional maintenance as well as for hematopoietic
stem cell proliferation and primitive maintenance.
1.2

Thesis Objectives
The overall objective of this thesis is to develop polymeric nanofiber constructs

that can serve as cell culture scaffolds, which can promote cell-substrate interactions
and are bioactive in soliciting favorable cellular responses. We believe that although
the topographical cues on a pristine nanofiber scaffolds are able to induce
morphological and cytoskeletal reorganization in cells [14-27], they are insufficient inproviding optimal regulation of cell behavior.
We therefore hypothesize that the development of nanofiber scaffolds that
present bioactive molecules is important in mimicking the native cellular
microenvironment, as these bioactive scaffolds can actively engage with cells andconsequently regulate their cellular activities.
We also hypothesize that a combination of nanofiber topographical cues andsurface biochemical cues will synergistically enhance the cell-substrate interactions

3

Chapter 1

and consequently induce further favorable cellular responses like cell adhesion, cellmorphological reorganization, cell differentiated functions and/or cell proliferation.
Through the systematic testing of unmodified and bioactive molecule-conjugatedfilms and nanofiber scaffolds in different primary cell culture models, we will be able
to demonstrate these synergistic cell-substrate interactions. In addition, we hope todemonstrate the versatility of our nanofiber bio-functionalization strategy for cell
culture applications through applying it in different cell culture models.
1.3

Thesis Scope
The general strategy of scaffold development involves nanofiber scaffold

fabrication via the electrospinning technique, followed by nanofiber biofunctionalization.

Thebio-functionalization

process

involves

theinitialfunctionalization of the nanofiber surface with carboxylic acid groups using UVinitiated poly(acrylic acid) grafting method. This is followed by conjugation of
bioactive molecules onto the functionalized nanofiber surfaces.
In this thesis, we will test the efficacy of this nanofiber bio-functionalization
strategy on two cell culture systems: (1) hepatocyte scaffold cultures and, (2)hematopoietic stem cell expansion cultures. The effect of immobilized bioactive
molecules in promoting cell-substrate interactions will be investigated. In addition, we
will also be focusing on the effect of nanofiber topography in synergisticallyenhancing these cell-substrate interactions, as outlined in the thesis objectives.
We

first

describethegalactose

bio-functionalizationofelectrospunpoly(caprolactone-co-ethyl ethylene phosphate) nanofibers for liver cell culture. Prior
to this study, nanofiber bio-functionalization strategies have never been demonstrated
in literature before. Using the bio-functionalization strategy described above, we have4

Chapter 1

developed a nanofiber scaffold culture that can sustain primary hepatocyte viability as
well as maintain the differentiated functions of the hepatocytes. The importance ofscaffold topographical cues and immobilized galactose biochemical cues on
hepatocyte morphological reorganization and function maintenance are investigated.
In addition, efforts to further enhance the hepatocyte functions through additional
nanofiber scaffold modification (3-methylcholanthrene incorporation) are presented.
Subsequently, we describe the amine bio-functionalization of electrospun
polyethersulfone nanofibers for ex vivo hematopoietic stem / progenitor cells (HSPCs)
expansion. HSPC expansion is commonly performed in a suspension culture format
where the importance of cell-substrate interactions has been undermined. Throughout
the course of this thesis research, we have discovered the significant roles that surface
immobilized amine molecules play in providing cell-substrate interactions to the
HSPCs. Using the same bio-functionalization strategy as described above, we havedeveloped an aminated nanofiber scaffold culture that can promote HSPC growth
while preserving the primitive HSPC multipotency. The importance of scaffold
topographical cues, immobilized amine biochemical cues and amine spacer lengths onregulating cellular responses like HSPC adhesion, proliferation and primitive
maintenance are also systematically investigated and presented.

5Chapter 2

CHAPTER TWO
Literature Review
2.1

Electrospun Nanofibers
Over the past decade, several techniques have been developed to fabricate

polymeric nanofibers. These techniques include electrospinning, drawing, phaseseparation, self-assembly, and template synthesis [9-13,28-29]. Among them,electrospinning, a technique that can produce continuous fibers with diameters
ranging from tens of nanometers to a few microns, is by far the most popular
technique because of its relative simplicity and scalability for industrial level
manufacturing and applications [9-13,28-29].

Figure 2.1: SEM images of fibers prepared by electrospinning of non-degradable (A-C)
and degradable polymers (D-F): (A) polyethersulfone (PES); (B) polyvinyl alcohol; (C)poly(bisphenol A carbonate); (D) polyhydroxybutyrate; (E) polycaprolactone; and (F)poly(caprolactone-co-ethyl ethylene phosphate) (PCLEEP).

6

Chapter 2

Electrospinning can be applied to spinning of a wide range of polymers (someexamples shown in Fig. 2.1), and the list of synthetic and natural polymers (bothbiodegradable and non-degradable) that can be electrospun into nanofibers has been
expanding rapidly [10-13]. Due to its simplicity and versatility in nanofiber
fabrication, the electrospinning technique has generated great interest in manypotential applications like nano-sensors, military protective clothing, media filtrationand life science applications [9-11].
2.1.1

Principles and Mechanisms
A typical laboratory electrospinning setup is schematically shown in Fig. 2.2.

The major components include: (1) a polymer solution feed unit (e.g. syringe pump);(2) a spinneret unit (e.g. syringe needle); (3) a high voltage power generator; and (4) agrounded collector.

Figure 2.2: Schematic illustration of an electrospinning setup. The inserts show adrawing of the electrified Taylor cone and a typical SEM image of nanofibers deposited
onto the collector.

The process of electrospinning is driven by electrical forces on free charges on
the surface or inside a polymeric liquid. In a typical electrospinning process, when alarge electric potential is applied between the collector and the spinneret, an electrical
7

Chapter 2

field is simultaneously induced. The polymer solution ball-shape drop pendent on thenozzle exit is then deformed, as a consequence of the force interactions between the
coulombic force (exerted by the external electric field) and the surface tension of thepolymer solution, into a conical shaped Taylor Cone [28,29,37-39]. At sufficiently
high electric potentials (typically 6 – 30 kV, depending on the surface tension ofpolymeric solutions), the electric field strength reaches a threshold value, and theelectrostatic force overcomes the surface tension, resulting in an ejection of a polymer
liquid jet. This jet is then subjected to an extremely high ratio of stretching throughwhipping1 [28,29,37-39] and rapid evaporation of solvent, leading to the formation of
sub-micron sized nanofibers, which were then attracted and gathered into a mesh at
the collector.

2.1.2

Parameters that Control the Electrospinning Process
Although electrospinning is said to be a relatively simple fiber fabrication

technique, there are surprisingly many parameters that govern this process, and it isthrough control variations of these parameters that result in generation of many
interesting nanofiber morphologies and structures as briefly discussed in the following
subsections.

1

The formation of nanofibers by electrospinning was previously attributed to the splitting or splayingof the electrified jet as a result of repulsion between surface charges. It appears that the cone shaped,
instability region is composed of multiple jets [29].
However, recent experimental observations demonstrated that the thinning of a jet during
electrospinning is mainly caused by the bending instability associated with the electrified jet. It appears
that the conical envelop contains only a single, rapidly bending or whipping thread. The frequency ofwhipping is so high that conventional photography cannot properly resolve it, giving the impressionthat the original liquid jet splits into multiple branches as it moves towards the collector [29].
In some cases, splaying of the electrified jet might also be observed, though it was never a dominant
process during electrospinning.

8Chapter 2

Electrospinning parameters in general can be classified under 2 categories:(1) Parameters which control the resultant fiber morphology (e.g. shape, size,
uniformity, defects, etc.); and(2) Parameters which control the resultant fiber mesh morphology (e.g. random,aligned, composite structures, etc.).
Among them, the four parameters presented in the following subsections arefound to be the more dominant control factors, as reported frequently in literature.
2.1.2.1

Effect of Polymer Concentration in Electrospinning Solution

The polymer solution concentration is an important parameter that affects thediameter, shape and the uniformity of the resultant fiber. The solution concentration
decides the limiting boundaries for the formation of electrospun nanofibers due tovariations in the viscosity and surface tension [12,40].

Figure 2.3: SEM images of electrospun PES with increasing concentrations in
dimethylsulfoxide solvent (w/w). (A) 5%; (B) 10%; (C) 15%; (D) 18%; (E) 20%; and(F) 25%. The polymer solutions are fed at a rate of 0.3 mL/h, electrospun at 13 kV, andfibers or beads are collected onto a grounded surface 160 mm away from the spinneret.
9Chapter 2

In general, low concentration solution forms droplets due to the influence of
surface tension, while higher concentration prohibits fiber formation due to higher
viscosity [40]: When the solution concentration increases, the resultant polymer
morphology shifts from polymer droplets, to beaded nanofibers, to uniform nanofibersof increasing diameters; until the solution becomes too viscous for fiber formation, as
shown in Fig. 2.3.

Figure 2.4: SEM images of PCLEEP fibers co-electrospun with increasing
concentrations of R18 in PCLEEP (w/w). (A) 0%; (B) 0.02%; (C) 0.1%; (D) 0.5%; (E)
1.0%; and (F) PCLEEP, R18 loading − PCLEEP fiber diameter relationship. PCLEEP
and R18 are dissolved in 8:2 dichloromethane / methanol solvent mixture The polymer
solutions are fed at a rate of 0.3 mL/h, electrospun at 12 kV, and the fibers or beads arecollected onto a grounded surface 60 mm away from the spinneret.

2.1.2.2

Effect of Ionic Additives in Electrospinning Solution

Fiber diameter can also be controlled via the doping of ionic additives into thepolymer solution. Charged ions in the polymer solution are highly influential in jet
formation. The ions increase the charge carrying capacity (electro-conductivity) of thejet, thereby subjecting it to higher tension with the applied electric field [12,41]. Also,

10

Chapter 2

the polymer solution jet radius has been demonstrated to vary inversely to the cube
root of the electrical conductivity of the solution [12]. The resultant effect is reduction
in bead formation or significant reduction in fiber diameters. To date, several reports
have successfully employed ionic additives like sodium chloride [41,42], heparin [43],octadecyl rhodamine B chloride (R18, Fig 2.4), pyridine [44], ammonium acetate, etc.,
to control nanofiber diameter and morphology.
2.1.2.3

Collector Design

Electrospun nanofibers are usually deposited on the surface of the collector(often a flat piece of conductive substrate) as randomly oriented nonwoven mesh,because of the bending instability associated with the spinning jet (Fig. 2.2). However,in recent years, new collector designs have been developed that were able to collectelectrospun nanofibers as uniaxially aligned arrays. The collector designs work
mainly by modifying the polymer jet movement via controlling the distribution ofelectric field between the spinneret and the collector [45-49], aligning the fibers
towards the sharp edges or corners of the collectors. Some of these designs include
the use of a pair of split electrodes [45-47] or a rotating drum, frame or wheel [48,49]
as the collector and they have all successfully demonstrated aligned nanofiber meshcollection.
2.1.2.4

Spinneret Design

The most recent addition to electrospinning process control that can significantly
influence both the fiber and fiber mesh morphology is spinneret design. In particular,
the fabrication of core-sheath nanofibers is a hallmark of the spinneret design
parameter [50-52]. Core-sheath nanofibers are fabricated by co-electrospinning two
different polymer solutions through a spinneret comprising of two coaxial capillaries.
As the electrospinning process took place very quickly, there would not be enough11

Chapter 2

time for the polymer chains from the two different polymer solutions to be mixed
before solidification. The resultant nanofiber core will have a material composition
that is different from its outer shell. Though the coaxial electrospinning technique isstill at its early development stages, recent papers have demonstrated that thenanofiber core can be used as a storage reservoir for proteins and drugs and that thisfiber system has potential in drug / protein delivery applications [50-52].
Another unrelated spinneret design is electrospinning using multiple spinnerets[53,54]. In this design, different polymer solutions are fed into two or more separate
spinnerets. Electrospinning using these spinnerets are then performed either
sequentially or simultaneously over the same collector, and thus multilayering
electrospinning or mixing electrospinning can be performed respectively. This design
has demonstrated the fabrication of multilayered nanofiber mesh as well as nanofiber
mesh with different polymer fibers that are intertwined or woven together.
2.1.2.5

Other Parameters

Other

processing

parameters

includespinneret−collector

gap

distance,

temperature, humidity, air-flow, applied electric field strength, solution feed rate,
solvent characteristics and composition2, etc. These parameters generally function as“fine-tuning” factors, affecting the fiber uniformity and reproducibility of theelectrospinning process. Although their roles have been discussed in literature [913,28-29], their influence in determining the fiber and fiber mesh morphology is not
as drastic as the four parameters previously discussed. However, we stress that future2

The intrinsic conductivity of the solvent will also contribute to the charge carrying capacity of thepolymer solution and will therefore determine the resultant fiber diameter range during electrospinning.
However, since the range of solvents with different conductivities is very limited for any given
polymer, the control of fiber diameters via different solvents compositions generally does not yieldsignificant differences.

12

Chapter 2

industrial electrospinning applications may still need to precisely control theseparameters in order to achieve high quality standards and reliability.
2.1.3

Electrospun Nanofibers in Cell Culture Applications
As discussed earlier, the relative versatility and simplicity of electrospinning in

fabricating nanofibers of various morphologies and structures has led to keen interestin various research fields [9-13,28-29]. In particular, the potential applications of
nanofibers as viable cell culture scaffolds have been intensely investigated in recent
years.
The key interest has been mainly the unique fibrous, surface nano-topographicalfeatures that a typical nanofiber mesh presents, compared with the smooth, featurelesssurfaces of tissue-culture plastics commonly used as cell-substrates for ex vivo cellprocessing 3 , and several researchers have even compared the topographicalmorphology of nanofiber mesh to resemble those of extracellular matrix (ECM)4 in
the native cell microenvironment. Indeed, abundant literature exists indicating that avariety of cell types, including fibroblasts, endothelial cells, muscle cells and stem
cells responded differently to the nano-featured surface topography as compared totheir smooth film counterparts, with or without the influence of additional physical orbiochemical cues [55-59].
It has long been recognized that the in vivo extracellular matrix, which provides arich context to the residing cells, includes topographical cue at the nanoscale [60-62].

3

Examples of tissue-culture plastics include polystyrene for culture flasks and plates, andpolytetrafluoroethylene for culture bags. These cultures surfaces are usually gas plasma treated, toprovide an optimal growth surface for the matrix-dependent tissue cultures.
4

Tissues are assemblies of one or more types of cells and their associated intercellular materials called
the extracellular matrix. For vertebrate animals, the ECM is made of a complex mixture of proteins andcarbohydrates, which are produced and maintained by the cells embedded in the network.

13

Chapter 2

A typical example is the basal lamina (basement membrane) that can be found inmany tissues. Inspired by the hypothesis that such a nanoscale feature may exert
unique interaction with cells, several groups have been investigating the role ofnanostructures on cell adhesion, proliferation, differentiation and migration [1427,55-59]. For example, the Curtis et al. has shown that nano-featured substrates
mediate different responses in epithelial fibroblasts, endothelial cells, smooth muscle
cells, and peripheral blood mononuclear cells compared to smooth film surfaces [5759]. The nano-featured substrates induce faster cytoskeleton organization, celladhesion and spreading in cells, accompanied by clearer and smaller focal adhesion
plaques, and a larger number of filopodia interactions with growth substrate.
Several groups investigating on cellular responses to nanofiber substrates havealso shown that these nanofiber substrates generally lead to differences inmorphological organization, gene expression, proliferation and differentiation
responses in fibroblasts, smooth muscle cells, endothelial cells, chondrocytes,
cardiomyocytes, bone marrow stromal cells, keratinocytes, mesenchymal stem cell,etc. [14-27]. For example, Li et al. demonstrated that seeding mesenchymal stem cells
on nanofiber scaffolds facilitated their differentiation into adipogenic, chondrogenic
or osteogenic lineages, with corresponding increases in the expression of lineagespecific genes [16,17]. Xu et al. showed that smooth muscle cells cultured on aligned
nanofiber scaffolds attached and migrated along the axis of the aligned nanofibers,
expressed spindle-like contractile phenotype, and exhibited actin and myosin
cytoskeleton organization that are parallel to the direction of the nanofibers [24].
Yang et al. also demonstrated in aligned and nonwoven nanofiber scaffolds thatnanometer diameter fibers enhances neurite growth in cerebellum stem cells better
than micron-sized fibers [25].
14Chapter 2

2.1.4

Nanofiber Modification for Cell Culture Applications
At present, the majority of these electrospun nanofiber studies have only

examined the effect of pristine nanofiber surface on cell behavior [14-27]. However,
we believe that optimal regulation of cell behavior requires more than an “inert”
scaffold that only provides topographical cues; and the electrospun nanofiber
scaffolds should also present specific binding domains for cells and growth factors
and serve many other functions (e.g. modulate growth factor responsiveness) that are
critical to the regulation of cell activities. The systematic design and modification of ananofiber scaffold containing these functional entities (bioactive molecules) would be
important in mimicking the cellular microenvironment.
A few groups have suggested electrospinning of pure ECM components or ECM/ synthetic polymer blends into nanofiber scaffolds as the alternative to synthetic
polymeric nanofiber scaffolds [63-66]. However, this strategy is only limited to fibrilforming proteins like fibrinogen, collagen, gelatin and elastin, and someglycosaminoglycans like hyaluronan. In addition, the fiber morphology is inherently
unstable in aqueous medium (the fibers degrade immediately) and additionalcrosslinking steps (e.g. treatment with glutaraldehyde, 1,6-diisocyanatohexane, poly
(ethylene glycol)-diacrylate, etc.) are usually taken to stabilize these scaffolds for cell
culture. Therefore, this strategy is not feasible for the presentation and delivery of the
majority of other bioactive molecules to cells. Nonetheless, several of these ECM
components have been successfully electrospun and stabilized as nanofiber scaffolds,
and cells (keratinocytes, fibroblasts, endothelial cells, etc.) cultured on these scaffolds
have showed enhancement in cell adhesion and proliferation compared with synthetic
polymer scaffolds [63-66].

15

Chapter 2

In general, nanofiber modification methods can be categorized into two different
approaches: either modifying the interior or bulk of the fiber, and/or modifying theexterior or surface of the synthetic polymeric nanofiber with bioactive molecules toprovide the desired cell responsive properties.
2.1.4.1

Doping of Bioactive Molecules

In this strategy, the nanofiber core is modified through the incorporation of
bioactive molecules like drugs or proteins into the polymer fibers, as illustrated in Fig.
2.5. The bioactive molecules are first added into the polymer solution. The doped
polymer solution is then electrospun into a nanofiber mesh. The bioactive moleculesin the nanofiber mesh are subsequently released and absorbed by cells during culture.
Various bioactive molecules like heparin, nerve growth factor, DNA
nanoparticles, drugs and bone morphogenetic protein have been incorporated into thenanofiber [43,67-69]. These doped nanofiber scaffolds were able to provide sustained
release of bioactive molecules to the target cells for extended periods of 1 week to 2
months, and the release kinetics of these molecules is dependent on both the bioactive
molecule solubility characteristics, as well as the degradation characteristics of thenanofiber scaffold. In general, for non-degradable and slow-degrading scaffolds, thebioactive molecule release kinetics is a function of the molecule diffusibility andsolubility [43,67,68], while for fast-degradable scaffolds the release kinetics is also
coupled with the scaffold degradation [69].
Luong-Van et al. demonstrated that sustained release of heparin from doped
polycaprolactone nanofiber scaffolds prevented the proliferation of vascular smoothmuscle cells in culture [43]. Liang et al. showed that controlled release of DNA
nanoparticles released from doped poly(D,L-lactic-co-glycolic acid) nanofiber

16

Chapter 2

scaffolds are effective in transfecting 3T3 cells in vitro [69]. We also demonstrated in
Chapter 4 that galactosylated nanofiber scaffolds doped with 3-methylcholanthrene isalso able to induce and regulate cytochrome P450 activity of hepatocytes in vitro.

Figure 2.5: Interior modification of electrospun nanofiber scaffolds.

2.1.4.2

Nanofiber Surface Modification

In this strategy, bioactive molecules are chemically immobilized onto thenanofiber surfaces, as illustrated in Fig. 2.6. These immobilized bioactive moleculesthen serve as ligands which will induce cell responses like adhesion, morphologicalorganization, proliferation or differentiation upon interaction with cells.
Numerous surface modification protocols are available in literature, which
describe conjugation of bioactive molecules onto film surfaces [30-36]. Nanofiber
surface modification strategies [70-73] have also imported these methods that haveworked well with film modification. In general, plasma or UV-initiated grafting
treatments, or chemical hydrolysis methods like aminolysis are first employed tofunctionalize the nanofiber surface with simple functional groups like carboxylic acid,amine, or aldehyde groups. Peptides, proteins, glycosaminoglycans and other ligands
are subsequently conjugated onto the functionalized surfaces via chemical
crosslinkers (e.g. glutaraldehyde, carbodiimide, etc.) [30-36,70-73].
17

Chapter 2

Figure 2.6: Exterior modification of electrospun nanofiber scaffolds.

Kim et al has demonstrated that cell attachment, spreading, and proliferation of
3T3 cells were greatly enhanced in RGD peptide immobilized electrospun poly(D,Llactic-co-glycolic acid) nanofibers, compared with unmodified nanofibers [73]. Ma et
al showed endothelial cells cultured on gelatin immobilized polycaprolactone
nanofibers exhibit enhanced spreading, proliferation, and expression of endothelial
cell markers [71].

2.2

Biomaterials Design for Primary Hepatocyte Culture
Liver failure has been the cause of death for thousands of people worldwide eachyear. When liver failure suddenly occurs in healthy individuals with normal livers, it
is termed acute liver failure (ALF), while the loss of liver function that complicateschronic liver disease is termed acute-on-chronic liver failure. Both ALF and acute-onchronic liver failure are curative via immediate liver transplantation [74,75]. Thoughpatient survival after transplantation has improved with advances in both patientmanagement and surgical techniques in recent years, the procedure however, is notalways available in a timely fashion due to the problems of organ availability [76].

18Chapter 2

To alleviate this problem, alternatives to whole liver transplantation organ are
currently under active investigation. Some of these methods include extracorporeal
bioartificial liver devices (BALs), transgenic xeno-transplantation, isolated celltransplantation, and tissue engineering of implantable constructs (Fig. 2.7) [77-81]. In
particular, research on BALs has been widespread as it is seen as a viable form of
supportive treatment to liver transplantation. BALs are generally developed as
temporary systems to attempt to expedite recovery from acute decompensation,
facilitate regeneration in ALF, and serve as a bridge to liver transplantation [77-81].

Figure 2.7: Approaches to cellular therapies for the treatment of liver disease.
Extracorporeal devices perfuse patient’s blood or plasma through bioreactors
containing hepatocytes. Hepatocytes are transplanted directly or implanted on
scaffolds. Transgenic animals are being raised to harvest a humanized liver.

BALs typically incorporate isolated cells (primary hepatocytes) into bioreactors
to simultaneously promote cell survival and function as well as provide for a level oftransport seen in vivo. The optimal design of a BAL generally spans across several
research disciplines. To cell biologists, the design and choice of the BAL cellularcomponent has been a primary focus: Optimization of medium formulations thatenhance primary hepatocyte functions and viability [82,83] as well as the design of19Chapter 2

immortalized cell-lines (e.g. NKNT-3, HepLiu, etc.) that express hepatic functions
have been their key research areas [79,80,84,85].
To BAL engineers, their research focus have been challenges in bioreactor scaleup for effective clinical therapy, as well as challenges in bioreactor designs thatprovide optimal bi-directional mass transport of oxygen, nutrients, patient’s plasma,etc., that is needed to sustain cell viability and allow export of therapeutic cell
products [78,79,86].
Lastly, to biomaterials scientists, their key interests have been the design andoptimization of biomaterial scaffolds that promote hepatocyte phenotype stabilizationin vitro. This is because although primary hepatocytes represent the most direct
approach to replacing liver function in hepatic failure, they are anchorage-dependent
cells and notoriously difficult to maintain in vitro: When enzymatically isolated fromthe liver and cultured in monolayer, scaffold or suspension cultures, the primary
hepatocytes rapidly lose adult liver morphology and differentiated functions [77-81].
One approach of hepatocyte phenotype stabilization includes the use of
extracellular matrix (ECM) components, which included both variations incomposition and topology. For example, surfaces coated with various ECM proteins,
such as laminin, fibronectin, and collagen [87-91], or conjugated with cell adhesion
peptides, such as RGD and YIGSR [92], have been used for hepatocyte culture.
Hepatocytes have been shown to attach well to these substrates [87-92]. An
improvement on the ECM culture system is sandwich cultures [89-91] or
microencapsulation

cultures[93,94]

which

were

designed

to

mimic

the

microenvironment of the adult hepatocyte where cells are sandwiched by extracellularmatrix in the space of Disse [78-80]. These sandwich or microencapsulation cultures

20

Chapter 2

typically packed hepatocytes closer together at higher densities, thereby promoting
homotypic cell-cell interactions. Hepatocytes cultured in this configuration have beenshown to stably express many liver-specific functions [89-91,93,94]. However, these“ECM scaffolds” face the same problems as electrospun ECM nanofibers in that they
are inherently unstable and attempts to scale-up these culture methods have met withlimited success so far.
Nevertheless, the importance of high density cell-packing in promoting
homotypic or heterotypic (in the case of hepatocyte cocultures with non-parenchymal
cells [95-97]) cell-cell interactions, which in turn stabilizes and maintains hepatocyte
liver-specific functions has been well documented [77-81,89-91,93-120], and this has
been the basis of culture systems involving hepatocyte spheroid formation [98-120].
2.2.1

Hepatocyte Function Maintenance through Spheroid Formation
Primary hepatocytes, when cultured on certain substrates conditions, will

physiologically undergo a series of morphological and functional changes, andeventually self-assemble into spheroids [33,34,98-120].

Figure 2.8: Morphology of hepatocyte spheroids. (A) Light microscope image of
spheroids after 4 days culture (bar represents 100 µm). (B) SEM imaging shows that thesurface of a mature spheroid is relatively smooth and cell-cell contacts are tight.

21

Chapter 2

Hepatocyte spheroids are three-dimensional, compacted multicellular sphericalaggregates that exhibit high degrees of cell-cell contacts (Fig. 2.8) [90,98-101]. Theyshow several structural similarities to native liver tissue such as gap junctional
complexes and bile canaliculi-like channels [98-101]. Hepatocyte spheroids exhibit
prolonged viability and express high levels of liver-specific functions including
albumin production, urea synthesis, and cytochrome P450 activity, in contrast to cells
cultured as monolayers [102-104]. At present, several different protocols havedemonstrated successful in assembling spheroids in vitro, they include:(1) Encapsulating [93,94], sandwich [89-91], or other packing (e.g. polyurethane
foam [105-107]) cultures where hepatocytes are physically packed close
together to facilitate cell-cell interactions and spheroid assembly;(2) Positively-charged polystyrene surfaces (PrimariaTM, BD Biosciences) [108]or negatively-charged proteoglycan-coated surfaces [109], which induces theformation of non-surface-adherent spheroids;(3) Rotary suspension cultures, where the swirling motion facilitates cellclustering [110,111]; and,(4) Hepatocyte cultures on galactose-immobilized substrates, where thehepatocyte-specific galactose ligand attaches hepatocytes and induces
spheroid formation along the substrate surface [33,34,112-120]. We shall beusing the scaffold galactosylation strategy to bio-functionalize our scaffolds
for hepatocyte culture.

22

Chapter 2

2.2.2

Hepatocyte Cultures on Galactosylated Scaffolds
Galactose-conjugated substrates have been proposed as alternatives forhepatocyte culture [33,34,112-120]. These substrates mediate hepatocyte adhesion
through the galactose−asialoglycoprotein receptor (ASGPR) interaction, andminimize the involvement of the integrin-mediated signaling pathway, which has
been shown to induce the loss of hepatocyte phenotypes [103]. The characteristic
attribute of these galactosylated substrates is also the propensity of hepatocytes toform aggregates or spheroids on them, in concomitance with maintaining higher
hepatocyte synthetic functions.
At present, several studies have shown that polymeric biomaterial surfaces
conjugated with galactose ligands can improve hepatocyte attachment and sustain
most of the cellular functions. This has been demonstrated in poly-N-p-vinylbenzylD-lactonamide-coated polystyrene surfaces or foam [113,114] and in galactosylated
polyethylene oxide hydrogel or polyacrylamide gel [115,116]. In addition,
galactosylated biodegradable polymeric scaffolds, such as alginate/galactosylated
chitosan sponge, galactosylated microcapsules, and polylactide-co-glycolide foam
[117-120], have also been designed for hepatocytes culture.
Recently, galactosylated PET films have also been developed for hepatocyte
spheroid culture [33,34]. A galactose ligand called 1-O-(6’-Aminohexyl)-Dgalactopyranoside (AHG) was designed for this culture system. This AHG ligand
consist of: (1) the galactosyl group; (2) a 6-carbon spacer (~ 0.7 nm) between thegalactosyl group and the surface conjugating point to facilitate the conjugation
reaction and to increase the accessibility of the ligand to cell surface receptors(ASGPR); and (3) a terminal primary amine group that allowed AHG conjugation toother surfaces via cross-linking chemistry. Details of AHG ligand synthesis are23

Chapter 2

attached in Appendix I. The AHG ligand is conjugated onto poly(acrylic acid)-grafted
PET surface through carbodiimide cross-linking chemistry [33,34]. We shall be using
this AHG ligand to bio-functionalize our hepatocyte culture scaffolds.
2.2.3

Bio-functional Nanofiber Scaffolds for Hepatocyte Cultures
Besides the ligand–receptor interaction, the substrate topography in micro-andnanometer ranges has been shown to influence cellular behavior and functions
including adhesion, migration, proliferation and gene expression [14-27,55-59].
Hepatocytes cultured on silicon scaffolds with micro-channels or in polyurethane
foams [105-107] have also exhibit aggregation behavior and functional maintenancethat are dependent upon the pore size of the scaffold. Electrospinning has been
increasingly investigated as an interesting technique to produce polymeric fibrous
scaffolds for cell culture applications. Several studies have shown that these nanofiber
scaffolds effect favorable cellular responses [14-27,70-73]. In this thesis, we would
like to extend the investigation to primary rat hepatocytes cultured on nanofiber mesh.
We would investigate how nanofiber topography and fiber bio-functionalization canbe employed to synergistically enhance cell-substrate interactions and hepaticfunctions for primary hepatocyte culture systems.

2.3

Biomaterials Design for Ex Vivo HSPC Expansion
Ex vivo hematopoietic stem/progenitor cells (HSPCs) expansion is one of themost challenging fields in cell culture. This is a rapidly growing area of tissue
engineering with widespread potential applications like gene therapy, immunotherapy,
bone marrow transplantation, and the production of mature blood cells for transfusion
medicine [132-140].

24

Chapter 2

The importance for HSPC ex vivo expansion in medicine is clearly demonstrated
in cell transplantation therapy treatments for cancer patients undergoing
chemotherapy and radiation therapy. Both chemotherapy and radiation therapy
treatment procedures target cells in mitosis, based on the fact that tumor cells arecontinuously proliferating. As a consequence, all other cells that undergo rapid
proliferation are also damaged and this applies especially to the hematopoietic system.
In most cases, this leads to total loss of all hematopoietic stem cells, making
allogeneic or autologous stem cell transplantation obligatory. Therefore, ex vivo
HSPC expansion can either increase the number of umbilical cord blood derived cells,
making this source also available for adult patients, or reduce the number ofleukapheresis procedures necessary for the collection of autologous cells [132-140].
Transplantation of ex vivo expanded lineage-restricted progenitor cells are also
viable cell therapy treatments [132-140]. For example, expansion of neutrophil
granulocytes and megakaryocytes can help to reduce the periods of neutropenia andthrombopenia respectively after chemotherapy. In addition, the expansion of naturalkiller cells or T-lymphocytes gives access to cell-mediated immunotherapy, while thegeneration and specific loading of dendritic cells offers possibilities for the in vivo
induction of antigen-specific immunity. The generation of erythrocytes or
thrombocytes can also lead to blood transfusions without a risk of viral contaminants.
Finally, hematopoietic stem cells can also serve as a target for somatic gene therapy
as they can offer a chance of a lifelong cure from genetic disorders due to their selfrenewing capacity.
As hematopoiesis in vivo is a highly regulated and complex process, thechallenge of ex vivo HSPC expansion has always been the optimization of cell
cultivation techniques that can control and regulate every step of differentiation of and25

Chapter 2

maturation of the hematopoietic cell, mimicking that of the cell’s native
microenvironment. Also, with much progress made in the understanding of thehematopoietic system over the past decades, the cultivation techniques and concepts
have also been evolving continuously, in order to expand HSPCs efficiently andeffectively.
In the following subsections, we will present and discuss the various knowledgeof the hematopoietic system, cell characterization techniques and cell cultivation
strategies that have progressively shaped the ex vivo HSPC expansion field.
2.3.1

The Hematopoietic System
Hematopoiesis is the process of generating mature blood cells. In the human

body, hematopoiesis mainly occurs in the bone marrow, predominantly in the femur,
sternum and pelvic bones [137-139]. In the marrow, the hematopoietic cells areembedded in the stromal tissue, which consists of several different cells types(adipocytes, fibroblasts, marcophages, endothelial cells, etc.) responsible for the
production of ECM (collagen, laminin, fibronectin, glycosaminoglycans, etc.) as wellas providing membrane-bound and soluble growth factors [137-140]. The interactions
between hematopoietic cells, stromal cells and ECM controls and regulates the entire
hematopoiesis process (Fig. 2.9): Everyday, almost 400 billion hematopoietic cells ofdifferent subtypes are produced in an average human to replace the natural loss ofcells [137]. Despite this tremendous proliferation, the balance between the different
lineages is very efficiently controlled by the bone marrow microenvironment toguarantee the many functions of the blood [137].
Despite enormous research progress over the past decades, the hematopoietic
system is still not completely understood. However, it is agreed that all hematopoietic

26

Chapter 2cells originate from a small population of multipotent hematopoietic stem cells thatproliferate and differentiate into the whole spectrum of mature blood cells [132-140].
The hematopoietic stem cells are the only hematopoietic cells that have the capacity
to self-renew, while for all other cells, proliferation is inevitably combined with alineage-specific differentiation and loss of immaturity.

Figure 2.9: Control of hematopoiesis in a bone marrow microenvironment.

2.3.2

Hematopoietic Stem/Progenitor Cell Sources
HSPCs can be harvested from the (1) bone marrow, (2) peripheral blood after

stem cell mobilization, or from (3) umbilical cord blood. The bone marrow is a goodsource of HSPCs and supporting stromal cells as it is the natural site forhematopoiesis. However, the harvesting of cells from bone marrow is an invasive
procedure that requires manual extraction under spinal or general anesthesia of theallogeneic donor, so alternative sources are preferred whenever possible [135-137].
HSPCs can also be harvested from peripheral blood after administration of G-CSF or
GM-CSF, which mobilizes HSPCs from the bone marrow. Though this is the standard
procedure to collect HSPCs from patients for autologous transplantation after high-27

Chapter 2

dose chemotherapy, it is not applicable to allogeneic transplantation, due to the healthrisk involved in using HSPC mobilization cytokines [135-137].
Umbilical cord blood is a promising alternative source of HSPCs for bothallogeneic and autologous hematopoietic stem cell transplantation, as it contains asignificant amount of progenitor cells with high proliferative potential, and the HSPCs
are harvested through procedures that are non-invasive for the mother or the neonate[135,139,141,144]. Cord blood sources also contains a lower burden of common viralcontaminations like EBV and CMV compared with the other two sources, and as thelymphocytes are more naive, the risk of a graft-versus-host disease is also reduced[135]. However, the main disadvantage of cord blood is the low number of HSPCs
obtainable due to the small volume of blood collected from the umbilical cords, sotransplantation is limited to pediatric patients of body weight less than 40 kg [141145]. Nevertheless, it is hoped that through development of efficient ex vivo
expansion strategies, sufficient quantities of HSPCs can be produced to engraft andsustain long-term hematopoiesis for adult patients.
2.3.3

Hematopoietic Stem/Progenitor Cell Characterization Techniques
HSPC characterization assays are important to ex vivo expansion cultures in thatthey evaluate and identify the primitive HSPC population in the cultures before orafter cell expansion have been carried out. They also serve as a means of assessing theeffectiveness of an expansion culture technique in producing HSPCs. There are atpresent a handful of assays used frequently in literature that have demonstrated merits
in identifying and evaluating HSPCs, and we will be using most of these assays
throughout this thesis to assess our scaffold expansion culture system. The detailed
protocols to these characterization methods are described in Chapter 5.3.2 & 6.3.2.

28

Chapter 2

HSPC characterization using flow cytometry techniques has been popular since
the discovery of the CD34 cell surface marker [146-149]. The CD34 antigen isfrequently used as a marker of stem cells as the CD34+ cell population contains a highamount of CFC, LTC-IC and mouse-repopulating capacity [135-138,143-156]. CD34+
is expressed most strongly on the most primitive cells and is progressively lost as cells
differentiate [146-150]. Although recent studies have discovered other stem cell
associated markers (e.g. CD133, CXCR-4) [138,157,158] and have also described
stem cell subpopulations that are CD34− [158,159], the CD34 antigen remains a
popular marker in identifying HSPCs clinically and in research.
In addition to flow cytometry, a series of in vitro assays like colony-forming cell
(CFC) assay, cobblestone area-forming cell (CAFC) assay or long term cultureinitiating cell (LTC-IC) assay are also used in progenitor cells characterization. In the
CFC assay, a small number of cells are cultivated for 2 weeks in cytokine containing
semi-solid medium. Colony-forming cells generate distinct colonies of moredifferentiated cells depending on their lineage specificity. For the detection of moreprimitive progenitor cells, CAFC or LTC-IC assays are used. Here a small number ofcells are placed on a stromal cell layer for 5-8 weeks. During this period the earlyprogenitor CAFC and LTC-IC differentiate and, in the case of CAFC, cobblestonearea-like cells are formed that can be easily identified. In the case of LTC-IC the earlyprogenitor cells differentiate into CFC, which are detected using a subsequent CFC
assay [160,161].
Finally, to evaluate the engraftment ability of the expanded HSPCs, in vivo
assays are used, which measures the potential of these cells in repopulating thehematopoietic system of sub-lethally irradiated immunodeficient mice (NOD/SCID),
monkeys or sheep [162,163].
29

Chapter 2

2.3.4

Hematopoietic Stem/Progenitor Cell Expansion Cultures
HSPC expansion culture systems have been evolving continuously (Fig. 2.10),

progressing with current knowledge of the hematopoietic system and also with
development of new technologies. The first HSPC expansion culture system wasdescribed by Dexter et al. in 1977 [164] using stromal cells to support HSPC growth[160,164-172]. The development of human recombinant cytokines in the 1990s has
contributed to the widespread popularity of cytokine supplemented HSPC suspension
cultures [173-177]. Finally, advancement in micro-fabrication technologies and theprogressive understanding and appreciation cell-cell and cell-substrate interactions of
the HSPC microenvironment has encouraged development of scaffolds that can
support HSPC expansion [178-185]. We will briefly discuss each of these HSPC
culture variants in the following subsections.

Figure 2.10: Generic HSPC cultivation variants. (1) Stromal cultures; (2) Cytokine
supplemented suspension cultures; and (3) HSPC-immobilized scaffold cultures thatcombine different aspects of the previous two culture variants.

2.3.4.1

HSPC Cultures with Stromal Cells or Conditioned Medium

Before the development of recombinant human cytokines, a feeder layer ofstromal cells was essential for HSPC cultivation. Stromal culture of hematopoietic
cells is a generic term that covers a variety of cultivation concepts, which includes (1)30Chapter 2

direct HSPC − stromal cells co-culture, (2) HSPC culture in stromal-conditioned
medium, and (3) partitioned non-contact co-culture.
The first direct co-culture system was demonstrated by Dexter et al. [164]. Since
then, several other stromal cells lines have been developed for the HSPC cultivation,
though most stable lines are of murine origin. Some examples are HESS-5, M2-10B4,
L88/5, hu-ST, etc. [160,164-168]. The main reason that stromal culture provides asuperior HSPC cultivation environment is that they appear to mimic the in vivo bonemarrow environment. The stromal cells provide direct cell-cell interactions as well ascell-matrix interactions through secreting ECM. The stromal cells also secrete
numerous different growth factors necessary for the maintenance and expansion ofstem and progenitor cells and many of these substances are still unknown [164-169].
However, harvesting and complete separation of expanded HSPCs from the stromalcells is very difficult in such systems.
A variation of direct co-culture is the application of exogenous stromalconditioned medium [168,169]. This concept enables the application of all bioreactor
systems developed for the cultivation of isolated HSPCs and allows the culture to besupplied with (unidentified) growth factors secreted from the stromal cells. The
conditioned medium can be produced in large amounts exogenously and stored until
application. An improvement of this approach is the in-situ generation of stromalconditioned medium in a partitioned non-contact co-culture, where HSPCs andstromal cells are physically separated in different compartments using micro-porous
membranes [140,170]. This permits active interaction between HSPCs and stromalcells, mediated by secreted molecules, which has been demonstrated to support the
expansion of the early progenitor cells more efficiently than the use of exogenous

31

Chapter 2

conditioned medium [140,170]. However, this technique suffers from difficult
technical requirements in set-up and process control.
Although the main attraction of stromal cultures is their superior ability for
HSPC expansion, clinical realization of these systems has been difficult, if not
impossible. The major drawback has been the origin of the stromal support. While theuse of autologous stromal cells would be a feasible possibility [171,172], in manycases this is not feasible. Furthermore, the use of cell lines is problematic as all
stromal cells have to be removed completely prior to transplantation, a demand which
is difficult to fulfill. In the case of murine cell lines a transfer of residue cells into apatient would be a xeno-transplantation, which is faced with extensive regulatory
hurdles.
2.3.4.2

HSPC Cultures with Human Recombinant Cytokines

The development of human recombinant cytokines has allowed cultivation andsignificant expansion of isolated hematopoietic stem and progenitor cells without theuse of supporting feeder-layer of stromal cells. The simplicity of these culture systems
is their main attraction: The cells are cultivated in chemically defined culture medium
containing defined combinations of cytokines [132-139,173-177].
Different kinds of culture set-ups have been devised for cytokine supplemented
cultures, but they are generally classified as suspension cultures. The various set-ups
include well plates, tissue-culture flasks, gas-permeable culture bags, spinner flasks
and stirred vessel bioreactors [132-139]. All these systems have the advantage ofbeing easy to handle, single-use devices, which enable an uncomplicated cell harvest.
For the larger culture bag and spinner flask set-ups, cell cultivation on a clinical scaleis possible, and these systems have been used clinically [132-139].

32

Chapter 2

The critical element of cytokine supplemented suspension cultures is thecytokine formulation. However, after almost two decades of intensive research of
cytokine effects on HSPCs [173-177], the optimal choices and combinations ofcytokines for the ex vivo expansion of HSPCs has not yet been determined [132-139].
This problem is mainly due to variations in culture conditions (e.g. use of serum orserum-free medium, cell source purity, HSPC sources, etc.) or assays (e.g. LTC-IC,
CAFC, mouse-engraftment assay, etc.) employed by the different studies.
Nevertheless, there is consensus among the various research studies, whichshows a list of cytokines that can positively regulate HSPC kinetics [173-177]. These
cytokines include stem cell factor (SCF), flt3/flt2 ligand (FL), interleukin (IL)-1, IL-3,
IL-6, IL-11, IL-12, leukemia inhibitory factor (LIF), granulocyte-macrophage colonystimulating factor (GM-CSF), granulocyte colony-stimulating factor (G-CSF),
thrombopoietin (TPO), and erythropoietin (EPO). In addition, supplementation of two
or more cytokines has been repeatedly demonstrated to positively enhance regulationof HSPC kinetics, as compared to single cytokine supplementation [132-139,173-177].
For example, the addition of SCF to a combination of IL-3 and GM-CSF resulted insignificant fold increase in cell colony formation [174]. In another study, FL in
combination with TPO resulted in extensive expansion with little differentiation of
CD34+ cord blood cells cultured under stromal-free conditions [175]. In this thesis,
we will be using a cytokine formulation adapted from Miller et al. [173], which
consist of SCF, FL, TPO and IL-3. Details of the cytokine formulation and culture
conditions are described in Chapter 5.3.2, 6.3.2 & 7.3.2.
Although cytokine supplemented suspension cultures enable better control of theculture conditions, cell processing, and significantly less regulatory hurdles ascompared to stromal cultures, important elements in the in vivo regulation of33

Chapter 2

hematopoiesis are missing in this approach: the cell-cell interactions between
hematopoietic and stromal cells, and biochemical and topographical cues provided bycell-matrix interactions between hematopoietic cells and the ECM substrate [137-140].
Therefore, it is likely that the cultivation of isolated hematopoietic cells in suspension
results in suboptimal HSPC expansion [137,140].
2.3.4.3

HSPC Cultures on Scaffolds

The immobilization of HSPCs into scaffolds is an attempt to imitate the complex
topographical microenvironment of the bone marrow. Scaffold topography itself
cannot support HSPC survival, so ex vivo expansion of HSPCs on scaffolds must
always be adapted with addition of other factors like stromal cells, ECM components
and/or cytokines [178-185]. The designs of HSPC scaffold culture systems are usuallydevised to improve the existing stromal culture and cytokine culture systems.
For scaffold cultures with stromal cells, the scaffolds incorporate the 3-D
perspective into the culture system, as conventional co-cultures are usually 2-D
monolayer cultures. HSPCs and stromal cells have been co-cultured in nonwoven
fabrics disks, large pore-size cubes and nylon screens [178-180]. For example, Li et al.
have shown that culture in 3-D non-woven polyester matrices enhanced cell–cell andcell–matrix interactions and expansion of stromal and hematopoietic cells [179].
Although high local cell densities can be achieved in these systems, they neverthelessstill retain the drawbacks related to the use of stromal cells.
For scaffold cultures with cytokine supplementation, the scaffold topography
provides cell anchorage and the possibility of HSPC growth modulation via cellsubstrate interactions to the otherwise substrate-less suspension cultures. A growing
body of evidence suggests the importance of surface chemistry as well as

34

Chapter 2

topographical features on the rate of HSPC proliferation and CD34+ cell expansion on
cytokine supplemented cultures [181-185]. For example, LaIuppa et al. have shownthat the type of substrate used in culture, ranging from polymers (polystyrene,
polysulfone, polytetrafluoroethylene, cellulose acetate) to metals (titanium, stainlesssteel) and glasses, can significantly affect the outcome of ex vivo expansion of HSPCs[181]. Leong et al. have also demonstrated that covalent surface immobilization of
extracellular matrix proteins (fibronectin) [185] and adhesion peptides (CS-1 and
RGD) [184] mediate HSPC adhesion to the scaffold and HSPC expansion.
These results suggest cytokine soluble cues as well as immobilized biochemical
and topographical cues could be synergistically involved in dictating the proliferationand differentiation of cultured HSPCs. These observations also prompted us toinvestigate whether the nano-topographical cues and immobilized biochemical cueson electrospun nanofiber scaffolds could also synergistically influence HSPC
adhesion, proliferation and multipotency phenotype maintenance, and we shalldemonstrate these effects in this thesis.

2.4

Concluding Remarks
Over the past decade, the understanding of the electrospinning technique has

been very progressive: It is possible to control this process and to electrospin almost
any polymer material into nanofibers of virtually any size and mesh configuration.
The simplicity and versatility of nanofiber fabrication has attracted interests in many
research disciplines. In particular, biotechnologists and medical researchers haverecently demonstrated the possibilities of electrospun nanofibers as viable cell culture
scaffolds. Though the mechanisms in which cells interact with nanofibers remains

35Chapter 2

largely to be elucidated, we hoped that through nanofiber bio-functionalization, we
would be able to understand and control this cell-nanofiber interaction better. Thus, inthe following chapters, we will demonstrate how bio-functionalized nanofiber
scaffolds can solicit various cellular responses through the synergistic effects ofnanofiber topography and immobilized biochemical cues.
We will demonstrate in Chapters 3 & 4 that galactosylated nanofibers can induce
hepatocyte morphological reorganization, and enhance attachment, albumin, andcytochrome P450 activity functions. We will also demonstrate in Chapters 5, 6 & 7
that hematopoietic stem / progenitor cell proliferation and phenotype maintenance isenhanced via ex vivo cultures on aminated nanofiber scaffolds.

36

Chapter 3

CHAPTER THREE
Stable Immobilization of Hepatocyte Spheroids on Galactosylated
Nanofiber Scaffolds for Liver Cell Cultures
3.1

Summary
Primary rat hepatocytes self-assemble into multi-cellular spheroids and maintaindifferentiated functions when cultured on a two-dimensional (2-D) substrate
conjugated with galactose ligand. The aim of this study is to investigate how afunctional nanofiber scaffold with surface-galactose ligand influences the attachment,
spheroid formation and functional maintenance of rat hepatocytes in culture, ascompared with the functional 2-D substrate.
Highly porous nanofiber scaffolds comprising of fibers with an average diameterof 760 nm were prepared by electrospinning of poly(ε-caprolactone-co-ethyl ethylene
phosphate) (PCLEEP), a novel biodegradable copolymer. Galactose ligand with adensity of 66 nmol/cm2 was achieved on the nanofiber scaffold via covalent
conjugation to a poly(acrylic acid) spacer UV- grafted onto the fiber surface.
Hepatocytes cultured on the galactosylated PCLEEP nanofiber scaffold exhibited
similar functional profiles in terms of cell attachment, ammonia metabolism, albumin
secretion and cytochrome P450 enzymatic activity as those on the functional 2-D
substrate, although their morphologies are different. Hepatocytes cultured on
galactosylated PCLEEP film formed 50-300 µm spheroids that easily detached fromsurface upon agitation, whereas hepatocytes cultured on galactosylated nanofiber
scaffold formed smaller aggregates of 20-100 µm that engulfed the functional
nanofibers, resulting in an integrated spheroid-nanofiber construct.
37

Chapter 3

3.2

Introduction
Primary rat hepatocytes, when cultured on certain culture configurations will

self-assemble into spheroids [33,34,98-120]. Hepatocyte spheroids are threedimensional, compacted multi-cellular spherical aggregates that exhibit high degrees
of cell-cell contacts [90,98-101]. In turn, these spheroids sustain viability for extended
culture periods and maintain high levels of liver-specific functions including albumin
production, urea synthesis, and cytochrome P450 activity, in contrast to cells cultured
as monolayers [102-104]. Galactose-conjugated substrates have been proposed asalternatives for hepatocyte culture. These substrates mediate hepatocyte adhesion
through the galactose-asialoglycoprotein receptor (ASGPR) interaction, and minimize
the involvement of the integrin-mediated signaling pathway, which has been shown to
induce the loss of hepatocyte phenotypes. In addition, hepatocytes cultured on
galactosylated PET films exhibit spheroid-forming capabilities: hepatocytes attach,then migrate and aggregate on the galactosylated surface, eventually forming surface
bound spheroids [33,34].
Besides the ligand-receptor interaction, the substrate topography in micro- andnanometer ranges has been shown to influence cellular behavior and functions
including adhesion, migration, proliferation and gene expression [14-27,55-59,105107]. Electrospinning has been increasingly investigated as an interesting technique to
produce polymeric fibrous scaffolds for cell culture applications. Nanofibers ranging
from 100 nm to 5 µm can be electrospun into a non-woven or an aligned nanofiber
mesh. Several studies have shown that these nanofiber scaffolds effect favorable
cellular responses [14-27,70-73]. We would like to extend the investigation toprimary rat hepatocytes cultured on nanofiber mesh composed of poly(εcaprolactone-co-ethyl ethylene phosphate) copolymer (PCLEEP). PCLEEP is a38

Chapter 3

recently developed biodegradable polymer [121]. With ester and phosphate linkagesin the backbone, PCLEEP shows an intermediate degradation rate between poly(εcaprolactone) and poly(ethyl ethylene phosphate) with a relatively linear degradation
profile. PCLEEP is soluble in several solvents and therefore confers good
processibility for the electrospinning process. It has also shown good tissuecompatibility and low cytotoxicity [67,121-123]. Initial studies also shows that
unmodified PCLEEP confers better hepatocyte adhesion properties compared to otherpolymers such as poly(ε-caprolactone).

Figure 3.1: Surface modification scheme for galactose conjugation to PCLEEP
nanofiber mesh. Insert: PCLEEP copolymer chemical structure.

In this study, we present the development of a biofunctional PCLEEP nanofiber
construct for hepatocyte culture (Fig. 3.1). This was achieved by conjugating
hepatocyte-specific galactose ligands onto the nanofiber surface. We investigated theeffect of nanofiber mesh on hepatocyte attachment, migration, spheroid formation andmaintenance of the differentiated functions, as compared with smooth 2-D substrates
39

Chapter 3

with or without galactose ligand. It is hypothesized that such functional nanofiber
scaffolds can be incorporated into a bioartificial liver assist device design, andtogether with their textured and porous nature, may promote hepatocyte-scaffold
interaction that will improve the stability of the attached cells.
3.3

Experimental Methods
Acrylic acid was purchased from Merck. All other chemicals were purchasedfrom Sigma-Aldrich unless otherwise stated.
3.3.1

Fabrication of PCLEEP Nanofiber Scaffolds
PCLEEP copolymer (Mw: 70,760) was synthesized according to a proceduredescribed by Wen et al. [121]. The PCLEEP copolymer was dissolved in acetone(21.5 wt% concentration) and placed in a plastic syringe fitted with a 27G needle. A
syringe pump (KD Scientific) was used to feed the polymer solution into the needle
tip. The feed rate of the syringe pump was fixed at 0.3 mL/h. PCLEEP nanofiber
meshes were fabricated by electrospinning at 17 kV using a high voltage power
supply (Gamma High Voltage Research). The nanofibers were collected onto
grounded 15 mm diameter coverslips located at a fixed distance of 80 mm from theneedle tip. Fiber diameters and mesh thickness were determined from images obtainedusing a field emission scanning electron microscope (FESEM) (SL30 FEG, FEI
Company). Representative nanofiber images were analyzed using NIH ImageJsoftware (http://rsb.info.nih.gov/ij/). At least 250 measurements were recorded foreach analysis.
As a control, PCLEEP film substrate was prepared by spin-coating 5 wt%
PCLEEP solution in acetone at 2000 rpm onto 15 mm diameter coverslips using a
spin-coater machine (Brewer Science Inc.).

40

Chapter 3

3.3.1.1

Surface Grafting of Scaffolds with Poly(acrylic acid)

Acrylic acid (AAc) was freshly distilled prior to use. The PCLEEP fibrous
scaffolds were cleaned with 70% ethanol in an ultrasonic water bath for 10 min prior
to surface grafting. Poly(acrylic acid) (PAAc) was grafted onto the scaffold surface byphoto-polymerization, as described by Uchida et al. [32]. Briefly, samples were
immersed in aqueous solution containing 2%-7% AAc solution and 0.5 mM NaIO4 ina flat-bottom glass container. The temperature of the solution was maintained at 10oC
by cooling the container in an ice-water bath. The solution was then exposed to UVfrom a 400 W mercury lamp (5000-EC, Dymax) for 15 min. The distance between thesamples and the lamp was 25 cm. The PAAc-grafted scaffolds were then thoroughly
washed with deionized water at 37oC for more than 36 h to remove any ungrafted
PAAc from the surface of the scaffold. Scaffolds modified using 6% AAc solution
was used for cell culture.
The amount of PAAc grafting on scaffolds was determined by Toluidine Blue O(TBO) colorimetric staining method [33,34]. Briefly, samples of known areas were
immersed in 0.5 mM TBO solution at pH10 and 20oC for 4 h. The samples were thenthoroughly washed with water at pH10 and 20oC to remove any non-complexed dyeadhering to the surface. The dye was then desorbed from samples in a 50% acetic acidsolution, and the concentration determined by spectroscopy at 633 nm (SpectraMax
190, Molecular Devices). The surface carboxyl (COOH) group density (the amount of
COOH groups per coverslip area) was calculated from a calibration plot of the OD
versus TBO dye concentration and assuming 1:1 stoichiometry of the binding
between TBO and surface carboxylic group.

41

Chapter 3

3.3.1.2

Galactosylation of Poly(acrylic acid) Grafted Scaffolds

1-O-(6’-Aminohexyl)-D-galactopyranoside (AHG) was synthesized according toprocedures described by Yin et al. (Appendix I) [33]. AHG conjugation onto PAAcgrafted scaffolds was achieved by immersing each scaffold into sodium phosphatebuffer (0.1 M, pH 8.0, 1.0 mL) with 1 mg AHG, 1 mg N-hydroxysulfosuccinimide(sulfo-NHS)

and

10

mg

1-ethyl-3-(3-dimethylaminopropyl)

carbodiimidehydrochloride (EDC). Ten mg of EDC each was added to the reaction mixture at 12 h
and 24 h later. After 36 h, the conjugation reaction was stopped. Scaffolds were thenthoroughly washed with deionized water and subsequently sterilized and stored in
70% ethanol until use in cell culture.
3.3.2

Hepatocyte Culture and Assays

3.3.2.1

Hepatocytes Isolation

Hepatocytes were harvested from 250 − 300 g male Wister rats by a two-step insitu collagenase perfusion method. Rats were maintained on ad libitum rodent feed
and water at room temperature and 40% humidity. All animal procedures wereapproved by the National University of Singapore’s Institutional Animal Care and
Use Committee. Hepatocyte viability was determined to be >85% using Trypan Blue.
3.3.2.2

Hepatocyte Attachment Assay

Freshly isolated rat hepatocytes (3.5×105) were seeded onto each 15 mm
diameter scaffold disks (2×105 cells/cm2), and cultured in William’s E medium
supplemented with 0.5 mg/mL BSA, 10 ng/mL EGF, 500 ng/mL insulin, 5 nM
dexamethasone, 50 ng/mL linoleic acid, 100 mg/mL penicillin, 100 U/mL
streptomycin, 50 pM ZnSO4 and 100 nM CuSO4. Cells were maintained in ahumidified incubator fed with 5% CO2. After 3 h of incubation, all unattached cells
42Chapter 3

were washed off by gentle washing using culture medium, and collected bycentrifugation (2,000 rpm). The cell pellet was washed once with PBS and then lysedwith cell lysis buffer (Promega).
The protein concentration in the lysate was determined using a BCA protein
assay kit (Pierce). The number of unattached cells was determined by comparing theprotein concentration in lysate with a standard curve generated with known numbersof cells. Subsequently, the attachment efficiency of hepatocytes on different scaffolds
can be calculated based on the cell number of the unattached cells for each scaffold.
3.3.2.3

Hepatocyte Culture Maintenance

The attached hepatocytes on different scaffolds were cultured in supplemented
William’s E medium as described above. Triplicate samples were maintained for eachhepatocyte function assay for each scaffold condition. Fresh medium was replenisheddaily, and old medium was collected for albumin determination. The collected
medium was centrifuged at 4,000 rpm for 10 min and the supernatant was stored at−20oC. At various time points of the culture, morphology of hepatocytes on thesescaffolds was viewed under an inverted phase contrast microscope (Carl Zeiss) andimaged on a digital camera (Nikon).
3.3.2.4

Albumin Secretion Assay

The albumin concentration in the culture medium collected at various time points
was determined by competitive ELISA method [124]. Briefly, samples were seriallydiluted, and rabbit peroxidase-conjugated rat albumin antibody (ICN) was added to a
final concentration of 0.6 µg/mL. After incubation at 37oC for 2 h, 100 µL aliquots ofeach sample were transferred to 96-well Maxisorp plates (Nunc), which were precoated with 100 µL/well of rat albumin at 0.2 µg/mL in PBS overnight at 4oC, and43

Chapter 3

washed three times with 0.05% Tween-20 in PBS before use. The samples wereincubated at 20oC for 2 h in a humidified chamber. After incubation, the plates were
washed three times with 0.05% Tween-20 in PBS and subsequently filled with 100µL/well 1-Step Turbo TMB-substrate (Pierce). Plates were incubated at 20oC in ahumidified chamber for another 30 min and reaction was immediately stopped byadding 100 µL of 1.3 N H2SO4. Optical density of the solution in each well wasmeasured at 450 nm using a spectrophotometer (SpectraMax 190, Molecular Devices).
3.3.2.5

Urea Synthesis Assay

On Days 1, 3 and 5, hepatocyte cultures were refreshed with culture medium
containing 1 mM NH4Cl, and incubated for 90 min. The NH4Cl-containing medium
was then collected and the cultures were refreshed with normal culture medium. The
medium samples collected were stored at −20oC and subsequently quantified for urea
contents using a urea nitrogen assay kit (Sigma).
3.3.2.6

Cytochrome P450 Activity Assay

Cultures were tested for the P450 enzymatic activity (P450-dependent 7ethoxycoumarin O-deethylase activity) using protocols adapted from Jauregui et al.
and others [97,125-127]. Briefly, on Days 1, 4 and 7, hepatocytes were pre-incubatedin culture medium containing 50 µM 3-methylcholanthrene (3-Mc) for 24 h to induce
cytochrome P450 production. The medium was then replaced with culture medium
containing 0.26 mM 7-ethoxycoumarin substrate. After 5 h incubation, the 7ethoxycoumarin-containing medium was collected and the cultures were refreshedwith normal culture medium. The medium samples collected were stored at −20oC
and subsequently assayed for 7-hydroxycoumarin (HCOD) using HPLC.

44

Chapter 3

For HPLC analysis, 200 µL aliquots of each sample were pre-incubated in 1.0 M
acetate buffer (pH 5.0) with 4.744 units/mL β-glucuronidase, 0.524 units/mL sulfatase,and 4-methylumbelliferone internal standard at 37oC. After 48 h, samples weretransferred to glass tubes (13×100mm, Iwaki). Half milliliter of saturated sodium
borate buffer (pH 9.0) and 4 mL chloroform was then added. After shaking for 3 min,the aqueous layer in the tubes was aspirated off and discarded. The remaining
chloroform layer was dried using a vacuum concentrator (Savant). The dried residue
was reconstituted with 200 µL of HPLC mobile phase (1:1 methanol/water). The
metabolites, HCOD, were analyzed by reverse-phase HPLC using a Symmetry C18
stainless steel column (Waters) at 1.0 mL/min mobile phase flow rate and a PDA 996
detector (Waters) at 326 nm.
3.3.2.7

Preparation for Scanning Electron Microscopy

Hepatocyte cultures were gently rinsed with PBS and fixed with 3%
glutaraldehyde for 30 min at 20oC and postfixed with 1% osmium tetraoxide for 15
min at 20oC. The hepatocytes were then dehydrated using a graded series of ethanol(25%, 50%, 70%, 80%, 90%, 95%, and 100%) followed by CO2 critical point drying(Samdri 780A). The samples were mounted onto aluminum stubs and gold sputtercoated before viewing under FESEM.
To prepare samples for freeze fracture, hepatocyte cultures were gently rinsed
with water after 3% glutaraldehyde fixation. Excess water was removed and samples
were cooled in liquid nitrogen for 10 min. Samples were then shattered using a razor
blade and immediately placed into a freeze dryer (Labconco) for 24 h. The samples
were mounted onto aluminum stubs and gold sputter-coated before viewing under
FESEM.

45

Chapter 3

3.3.3

Statistical Analysis
All values were expressed as mean ± standard deviation (SD). Statisticaldifferences were determined by two-tailed student’s t-test.
3.4

Experimental Results

3.4.1

Optimization of PCLEEP Electrospinning
PCLEEP fibers were generated by the electrospinning process. The PCLEEP

electrospinning process is optimized by gradually increasing the polymer solution
concentration until a suitable concentration is found that will produce unbeaded,uniform fibers. We have found the optimal concentration to be 21.5 wt% PCLEEP in
acetone. The other parameters for operating the electrospinning process were
described in Chapter 3.3.1.

Figure 3.2: SEM characterization of PCLEEP nanofiber mesh. (A) SEM image showsthat the mesh is highly porous and the fibers are randomly orientated. (B) Fibers have adiameter distribution between 0.13 µm and 2.3 µm, and an average diameter of 0.76 µm.
(C) Mesh thickness is approximately 15 µm.

46Chapter 3

SEM images showed that nanofibers arranged randomly in the mesh (Fig. 3.2)
and their diameters ranged from 300 nm to 1.5 µm (with frequency higher than 5%,
Fig. 3.2B). The average diameter was 760 nm. This highly porous mesh contained
pores of mainly sub-cellular sizes (<10 µm, Fig. 3.2A). The thickness of the meshvaried with the deposition time. The meshes used for this study had a thickness ofapproximately 15 µm (Fig. 3.2C).
3.4.2

Optimization of Scaffold Galactosylation Process
Surface conjugation of galactose ligand onto PAAc grafted scaffolds wasachieved using the same protocol as Yin et al. and the effectiveness of the conjugation
scheme was successfully demonstrated through XPS analysis [33,34]. Since thegalactose conjugation efficiency using this scheme was shown to be above 90%, thesurface galactose density can therefore be controlled by varying the PAAc grafting
density on the scaffold surface [33,34].
The PAAc grafting density of PCLEEP nanofiber mesh was lower than that of
PCLEEP spin-coated film when 2-7% of AAc solution was used in the grafting
reaction (Fig. 3.3). This result was expected because UV-initiated polymerization
required direct exposure of the surface to UV light for surface grafting to occur. Dueto its porous nature, the PCLEEP nanofiber mesh had a lower projected flat-area ascompared to PCLEEP film. During UV-irradiation, the nanofiber mesh area exposed
to UV was less than that of the film on the same 15 mm coverslip, which resulted in
the lower grafting density.
Scaffolds modified using 6% AAc were used for cell culture experiments. Under
these conditions, nanofiber meshes had an average surface COOH density of 66
nmol/cm2, whereas PCLEEP films yielded an average surface COOH density of 127
47

Chapter 3

nmol/cm2. Previous studies had shown that both the attachment and spheroid forming
ability of the primary rat hepatocytes cultured on galactosylated PET surfaces were
almost independent of the surface graft density of COOH group within the range of 20
to 560 nmol/cm2 [33,34]. Thus, the difference in COOH densities between 66 and 127
nmol/cm2 was not expected to lead to significant differences in cell morphology andfunction, if comparison is made on the same type of substrate.

Figure 3.3: Effect of acrylic acid monomer concentration used for UV-initiated graftpolymerization on the surface concentration of the grafted carboxyl groups on the
PCLEEP nanofiber mesh and spin-coated film surface. Data are means ± SD of 2
independent experiments, each conducted in duplicates.

3.4.3

Hepatocyte Functional MaintenanceFor convenience, galactosylated nanofiber mesh and film will be referred to as

Gal-nanomesh and Gal-film respectively, while unmodified nanofiber mesh and film
will be referred to as Un-nanomesh and Un-film respectively. Hepatocyte attachment
efficiency measured at 3 h after cell seeding was shown in Fig. 3.4. Hepatocyte
function normalized against the total number of attached cells was evaluated byalbumin secretion level (Fig. 3.5), urea synthesis activity (Fig. 3.6) and 3-MC induced48Chapter 3

P450 activity (Fig. 3.7) as a function of time. The results were determined fromalbumin, urea, or HCOD concentrations in the medium respectively.
Hepatocyte attachment was enhanced when seeded on Gal-nanomesh and Galfilm (83% and 90%, respectively) attachment after 3 h (Fig. 3.4), compared with 5054% attachment for the unmodified surfaces (p < 0.05). The cell attachments on Galnanomesh and Gal-film conditions were similar, suggesting that the geometry of thesubstrata does not influence cell adhesion in this system. The improved attachment
efficiency on the galactosylated substrates was attributed to cell-scaffold interactions
between immobilized galactose ligand on substrates and ASGPR on the surface ofhepatocytes.

Figure 3.4: Hepatocyte attachment on galactosylated and unmodified nanofiber meshes
and spin-coated films 3 h after seeding. *p < 0.05; **p < 0.05. Data are means ± SD of 2
independent experiments, each conducted where n = 6.

Hepatocytes cultured on galactosylated scaffolds showed higher albumin
synthesis function than unmodified substrates, respectively (Fig. 3.5), leading to a
bigger difference (> 2 fold) on days 3 − 5. Albumin synthesis function declined
gradually for the two galactosylated-substrates during the first 2 − 3 days, and49

Chapter 3

maintained the same level thereafter. Comparing the two galactosylated-substrates,
albumin secretion level of Gal-nanomesh group was about 22% higher than that ofGal-film group for the first two days, but was slightly lower (5% - 9%) than that ofGal-film group from day 3 onwards.

Figure 3.5: Albumin secretion level of hepatocytes at various time points normalizedagainst the total number of attached cells. Data are means ± SD of 2 independent
experiments, each conducted in duplicates.

However, the maintenance of urea synthesis function was less pronounced thanthat of albumin secretion function (Fig. 3.6). Urea synthesis (or ammonium removal)
levels for the two galactosylated-substrates were similar throughout the assay period –they declined gradually over time. Interestingly, hepatocytes cultured on unmodified
nanofiber substrate maintained urea synthesis function at the similar level as that
cultured on galactosylated substrates. They showed about 1.6 and 2.5 fold higher urea
synthesis activity than those on Un-film on days 3 and 5, respectively.
Hepatocytes cultured on the two galactosylated substrates responded well to 3Mc induction and expressed higher levels of induced cytochrome P450 function thanthose cultured on unmodified substrates (Fig. 3.7), peaking on day 5 (> 4.4 fold). The
50

Chapter 3

nanofiber mesh substrates resulted in higher HCOD synthesis activity than film
substrates on day 1, but the difference became less apparent on days 3 and 5.

Figure 3.6: Urea synthesis function of hepatocytes at various time points normalizedagainst the total number of attached cells. Data are means ± SD of 2 independent
experiments, each conducted in duplicates.

Figure 3.7: 3-Mc induced P450 function of hepatocytes at various time points
normalized against the total number of attached cells. Data are means ± SD of 2
independent experiments, each conducted in duplicates.

51

Chapter 3

In addition, a significant difference between the Gal-nanomesh and Gal-filmgroups was the long-term adherence of attached hepatocytes, as illustrated in Fig. 3.8.
No significant cell detachment was observed for Gal-nanomesh throughout the assay
period, whereas mature spheroids gradually detached from the Gal-film surface fromday 3 onwards. Despite the special care taken during medium change for the Gal-filmgroup, spheroid detachment is inevitable, though these detached spheroids remained
viable and functional in the culture well. Only a small number of spheroids remained
attached to the Gal-film surface by day 8. The Un-nanomesh and Un-film substrates
did not show noticeable amount of cell detachment throughout the culture period.

Figure 3.8: Hepatocyte spheroid adhesion on galactosylated scaffolds after 5 days ofculture. (A) Hepatocytes cultured on Gal-nanomesh formed undetachable aggregates
throughout the scaffold, whereas; (B) Hepatocytes cultured on Gal-film formed
spheroids that can be easily detached from the surface, forming a spheroid suspension.
Scale bar is applicable to both images as they are taken at the same magnification.

3.4.4

Hepatocyte Morphological Changes
Hepatocytes exhibited very different morphologies when cultured on different

substrates. Optical microscope images at 3 h, 1 day and 3 days after cell seeding arepresented in Fig. 3.9. Three hours after cell seeding, attached hepatocytes remained52

Chapter 3

rounded and singular, similar to the morphologies exhibited before cell seeding. Both
Gal-nanomesh and Gal-film groups showed a closer packing of cells on the substrates
as compared to Un-nanomesh and Un-film groups. This is indicative of the higher cellattachment efficiency on the galactosylated substrates over unmodified substrates.
Hepatocytes cultured on both Gal-nanomesh and Gal-film began to cluster andform aggregates after day 1. Cell movement and aggregation appear to be slightly
restricted by the uneven nanofiber surface topography as compared to film surface.
By day 3, aggregate formation on Gal-nanomesh was complete, resulting in smaller,
more uniform spheroid-like structures of sizes between 20 and 100 µm in diameter, incomparison with the larger spheroids with wider distribution of the spheroid diameter(50−300 µm) on Gal-film. Single cell was rarely seen on Gal-nanomesh, but present
on Gal-film substrate (Fig. 3.8 − 3.10). Most single cells lost viability on day 3.
For hepatocytes cultured on unmodified scaffolds, the morphologies displayed
were irregular and not as uniform as those on galactosylated substrates. The absenceof ligands on substrates to direct or initiate cell migration may be the primary cause of
the morphological irregularities. For example, RGD groups on collagen-treatedsurfaces are known to induce hepatocyte spreading through RGD−integrininteractions [87-92], whereas galactose-modified surfaces direct hepatocytes to form
multi-cellular spheroids through a combination of galactose-ASGPR and cell-cellinteractions [33,34,112-120]. For unmodified substrates, non-specific hydrophobic
interactions and some ionic interactions between the cell membrane and polymer
surface probably allowed hepatocyte adhesion to take place. The morphology adopted
by the hepatocytes may be due to the variations of these local non-specificinteractions. Hepatocytes cultured on Un-nanomesh formed irregular spreading cell

53

Chapter 3

clumps. Similarly, for hepatocytes cultured on Un-film, the cells exhibited bothspreading and aggregating morphologies.

Figure 3.9: Morphology of hepatocytes at 3-h, 1-day and 3-days after seeding when
cultured on different substrates. Scale bar represents 100 µm, applicable to all images as
they are taken at the same magnification.

SEM images of remaining adherent spheroids on Gal-film showed typicalspheroid morphology with a relatively smooth surface (Fig. 3.10A−C). Numerous
microvilli also dotted around spheroid surface. These spheroids closely resembled thefunctional spheroids reported in literature, which exhibited tight cell−cell contact[90,98-101], in contrast with those rough and bumpy spheroids that are damaged by
toxicants [128]. SEM images of the spheroid-like aggregates cultured on Gal54

Chapter 3

nanomesh also showed similar surface features as compared with the spheroids on
Gal-film (Fig. 3.10D−F). However, these aggregates were distinctively flattened onto
the nanofiber mesh. The aggregates appeared to engulf the nanofibers, forming an
integrated spheroid−nanofiber construct with the nanofiber mesh. This explains thegood cell immobilization efficiency of the Gal-mesh substrate observed throughout
the culture period, as depicted in Fig. 3.8.

Figure 3.10: SEM images of hepatocytes after 8 days of culture: (A−C) Hepatocytes
cultured on Gal-film formed rounded spheroids that did not integrate with the scaffold.
In contrast; (D−F) Hepatocytes cultured on Gal-nanomesh showed that the aggregates
engulfed the functional nanofibers.

55

Chapter 3

Figure 3.11: SEM images of freeze-fractured hepatocytes on Gal-nanomesh after 8 daysof culture: (A & B) PCLEEP nanofibers (arrow) can be found within the hepatocyte
aggregate; (C & D) Aggregates exhibit no fibers within, which maybe attributed todegradation of the biodegradable PCLEEP nanofibers.

The freeze-fractured spheroid-like aggregates allow the direct observation of theinterior structure of the spheroids (Fig. 3.11). SEM images of the freeze-fractured
aggregates showed many voids reminiscent of bile canaliculi channels previously
described in literature [90,98-101]. In addition, SEM images also clearly showed that
nanofibers could be found within the aggregate (Fig. 3.11A & B). However, in many
instances, there were fewer or no nanofibers present inside the aggregates while fibers
were found along the immediate exterior of the spheroids (Fig. 3.11C & D). We
speculate that the biodegradable PCLEEP nanofiber within the aggregates, in thisinstance, might have been degraded and absorbed by the hepatocytes.

56

Chapter 3

3.5

Discussion
Previous studies on polyurethane foam scaffolds or collagen-treated siliconscaffolds with micro-channels of super-cellular pore sizes and partition barriers haveshown that hepatocyte aggregates would conform to the shape of the pore space[93,94,89-91,105-107]. The partition barriers and pores of the scaffold do influence
the way hepatocytes move and organize themselves into multi-cellular aggregates.
However, little information is available to suggest how hepatocytes would respondwhen they are cultured on a micro- or nanofiber substrate with sub-cellular-sizedpores, where such a substrate would present itself as a ‘‘rough’’ or ‘‘textured’’ surface
to a rounded hepatocyte.
Scaffold micro-architecture and topography have been shown in literature to
influence cell migration, adhesion, proliferation and various other cell behaviors,
especially for smooth muscle cell, fibroblast or endothelial cell types [14-27,55-59].
However, scaffold topography alone does not promote favorable cell morphology andfunctional maintenance of primary rat hepatocytes, as shown in the results presented.
This is echoed by a number of studies aiming to immobilize hepatocytes in porous
scaffolds or on substrates through surface conjugation of functional ligands (e.g.
galactosylation [33,34,112-120], collagen treatment [87-92]) or high-density cell
entrapment [93,94,105-107].
In this study, our goal was to examine whether the introduction of a sub-cellular
sized, highly textured topography, together with the surface-conjugated galactose
ligand, would further influence spheroid formation and functional maintenance ofhepatocytes. Electrospinning was used because it could lead to polymeric nanofiber
mesh with sub-micron diameter fibers (average of ~760 nm), which randomly
overlaid one another to form a highly textured, sub-cellular-sized micro-porous
57

Chapter 3

scaffold. This study represents the first one where cell-adhesive molecules arecovalently attached to an electrospun scaffold for cell-substrate interaction studies.
Galactose ligands (AHG) were covalently conjugated onto the PAAc-grafted
nanofiber mesh, because several studies have shown that biomaterial surfaces
conjugated with galactose ligands can improve hepatocyte attachment and retain mostof the cellular functions [33,34,112-120]. The characteristic attributes of thesegalactosylated substrates are that:(1) Hepatocytes specifically adhere to these surfaces through galactose-ASGPR
interactions; and(2) The propensity of hepatocytes to form aggregates or spheroids on them, inconcomitance with maintaining higher hepatocyte synthetic functions.
Hepatocytes cultured on galactosylated scaffolds (Gal-nanomesh and Gal-filmgroups) clearly showed superior functions in terms of cell attachment, albumin
synthesis and 3-Mc-induced cytochrome P450 function, and to a less extent, the urea
synthesis activity, as compared to hepatocytes cultured on unmodified scaffolds (Unnanomesh and Un-film control groups). Consistent with previous reports on
galactosylated membrane hepatocyte cultures [33,34], the albumin synthesis function
and urea synthesis function decreased over time, whereas cytochrome P450 function
peaked at day 5 followed by a decrease. This again suggested the complexity ofhepatocyte functional maintenance: different set of synthetic functions could be
influenced by biochemical cues or topological cues to a different extent.
The initial albumin secretion level and cytochrome P450 activity of hepatocytes
on Gal-nanomesh were higher than Gal-film on day 1–2, but the functional profiles atlater time points for Gal-nanomesh group and Gal-film group were surprisingly
58Chapter 3

similar, even though the spheroid morphologies displayed by hepatocytes in these twogroups were different: spheroids on the Gal-film appeared mostly spherical, with asmall and flattened interface with PCLEEP film; on the other hand, spheroids on Galnanomesh were flattened and integrated into the nanofiber mesh scaffold. It appears
that so long as individual hepatocytes were able to aggregate and resume appropriate
homotypic cell−cell contacts, hepatocyte viability and functions could be maintained
at a higher level compared to unmodified substrates, regardless of the resulting shapeof the spheroids. It suggests that the biochemical cue (galactose group on the substrate)in this case has a higher impact on cell functional maintenance compared with
topological cue (nanofiber substrate vs. 2-D film).
Hepatocytes cultured on galactosylated nanofiber mesh presented an interesting
morphology: hepatocytes formed spheroids that engulfed the galactosylated
nanofibers. As a consequence of this, spheroids were immobilized on the scaffold andwould not detach from the substrate upon agitation, unlike those attached on
galactosylated film. It appears that galactose-ASGPR ligand-receptor interaction
allowed hepatocytes to aggregate around and within the galactosylated nanofiber
mesh. This observation indicates that the nanofiber topography enhances the overallcell-substrate interaction, comparing with the galactosylated 2-D film. The increased
cell-substrate interaction is significant enough to alter the spheroid morphology(somewhat flattened vs. spheroidal).
This hepatocyte spheroid immobilization and stabilization strategy through theuse of galactosylated nanofiber scaffolds would be advantageous in the design of abioartificial liver device, where the hepatocytes could attach to a substrate/scaffold
with high surface area immobilized with a cell-specific ligand, maintain their59

Chapter 3

differentiated functions, and remain stable against the perfusion and shear forces inthe bioreactor.
3.6

Concluding Remarks
We have shown in this chapter that hepatocyte functions are maintained on

galactosylated nanofiber scaffolds, similar to a galactosylated-film substrateconfiguration. Interestingly, galactosylated nanofiber scaffolds exhibit the uniqueproperty of promoting hepatocyte aggregates and cell infiltration within the mesh andaround the fibers, forming an integrated spheroid-nanofiber construct.
This mechanically stable hepatocyte–nanofiber construct suggests the potential
application of galactosylated nanofiber scaffold in liver cell culture. For example, thissystem will have advantages in a bioartificial liver (BAL) setting, where highdensities of stable, immobilized spheroids can be maintained on galactosylated
nanofiber scaffolds within the BAL bioreactor.
These results also demonstrated that the nanofiber topography together withsurface-immobilized

galactose

ligandsynergisticallyenhancecell-substrateinteraction as indicated by hepatocyte adhesion and infiltration, even though thisenhanced cell-substrate interaction did not translate into significantly higher
functional enhancement as compared to galactosylated film condition.
Therefore, we shall demonstrate in the following chapter how higher hepatocyte
functional enhancement (cytochrome P450 activity) can be brought about throughfurther bio-functionalization of the galactosylated nanofiber scaffold.

60

Chapter 4

CHAPTER FOUR
Hepatocyte Cytochrome P450 Inducing Dual-Functional Nanofiber
Scaffolds for Hepatocyte Culture
4.1

Summary
In nature, cell survival, proliferation, differentiation and functions are regulated

by a set of complex, spatially and temporally controlled milieu of biochemical andtopographical cues emanated from the extracellular microenvironment. In order tocreate a cell culture scaffold that can deliver multiple microenvironmental cues that
can influence cell fate and functions, we had designed a novel dual-functional
electrospun nanofiber scaffold comprised of two nanofiber mesh layers that weremodified differently to induce two separate biological responses from hepatocytes.
The first nanofiber layer was galactosylated to mediate hepatocyte attachment andinduce aggregation formation, while the second layer was loaded with 3methylcholanthrene (3-Mc) to enhance cytochrome P450 activity of hepatocytes.
Primary rat hepatocytes cultured on the galactosylated nanofiber scaffolds loadedwith different concentrations of 3-Mc were compared for their cell attachment
efficiency, albumin secretion activity and their cytochrome P450-dependent 7ethoxycoumarin O-deethylase activity. This dual functional nanofiber scaffold
mediated hepatocyte attachment with slightly lower efficiency as compared to single
layer galactosylated nanofiber scaffold. More importantly, cytochrome P450 activityof the hepatocytes cultured on the multi-functional scaffold correlated well with 3-Mc
loading level. Transwell experiments showed that transfer of 3-Mc to hepatocytes
through cell-fiber direct contact was the dominant transport route. This study
61

Chapter 4

demonstrated the feasibility of creating multi-functional nanofiber scaffolds thatserves both as an adhesive substrate and as a delivery vehicle for bioactive molecules.

4.2

Introduction
Electrospun polymeric nanofibers have demonstrated its potential in many

biomedical applications, including the production of scaffolds for tissue engineering[14-27] and bioactive molecules delivery [43,67-69]. In the tissue engineering context,
the high porosity and surface area to volume ratio achievable in a typical electrospun
nanofiber scaffold provided large areas and spaces for cell attachment and the
topographical features provided by electrospun fibers play a significant role inregulating cell responses [14-27]. The versatility of the electrospinning process also
produced many different fibrous scaffolds with distinct geometry. For example, on
axially aligned nanofiber scaffolds, neuronal cells and cardiomyocytes have beenshown to grow axially along the fiber’s orientation [20,25]. Neuronal cells have also
been shown to elongate at different rates on aligned nanofiber scaffolds with different
fiber diameters [25].
Recently, nanofiber modification techniques were also explored in order tocontrol specific cell responses through the use of ligands or drugs incorporated into
the scaffold. Nanofiber modification techniques can be broadly categorized under 2methods: The first method involves the doping of bioactive molecules or protein
factors into the polymer solution prior to electrospinning (Chapter 2.5.1). This method
results in a nanofiber scaffold that can steadily release these molecules or proteins tothe cells in culture [43,67-69]. The second method involves chemical modification of
the polymer itself that results in covalently attached ligands on the surface of thenanofiber (Chapter 2.5.2) [70-73]. This method resulted in a bioactive nanofiber
62

Chapter 4

surface that can interact with specific ligand receptors on cell membranes and thus
inducing specific responses from the cells.
In the previous chapter, we had developed a surface bio-functionalized nanofiber
scaffold for primary hepatocyte culture. Poly(acrylic acid) was first grafted onto thefiber surface by photo-polymerization to introduce surface carboxylic acid groups.
Subsequently, amine-functionalized galactose ligands were conjugated onto thenanofiber surface using carbodiimide cross-linking chemistry. The galactosylated
nanofiber scaffold was able to mediate primary hepatocyte adhesion through thegalactose – asialoglycoprotein receptor (ASGPR) interaction. The galactosylated
nanofiber scaffold induced the formation of spheroids that enveloped the functional
nanofibers, resulting in an integrated spheroid-nanofiber construct.
This unique cell-fiber interaction could maintain hepatocyte viability and certain
functions like albumin production and urea synthesis. However, one of the crucial
functions, cytochrome P450 enzymatic activity was low, even though it can be
induced through stimulation by 3-methylcholanthrene (3-Mc) in the culture media. 3Mc, a polycyclic aromatic hydrocarbon, is a potent inducer for the induction of
CYP1-dependent xenobiotic oxidation, and is used extensively in various hepatocyte
studies [97,125-127,129]. However, hepatocyte cytochrome P450 induction required
repeated 3-Mc doping in culture medium, as 3-Mc solubility in aqueous solution is
low [130,131]. Therefore, we had fabricated a 3-Mc loaded nanofiber scaffold withthe assumption that hepatocyte cultured on this scaffold will have enhanced
cytochrome P450 activity through the absorption of 3-Mc released by the scaffold.
In this present study, we have designed a dual biofunctional electrospun
nanofiber scaffold through the layering of two nanofiber meshes that are modified

63

Chapter 4

differently to induce two separate biological responses from hepatocytes (Fig. 4.1).
The first nanofiber layer is galactosylated to mediate hepatocyte attachment andaggregation formation, while the second layer is 3-Mc loaded to enhance cytochrome
P450 activity in hepatocytes. Primary hepatocytes cultured on galactosylated
nanofiber scaffolds loaded with different concentrations of 3-Mc will be compared fortheir P450-dependent 7-ethoxycoumarin O-deethylase activity. In addition, the modeof release of 3-Mc to the hepatocytes will also be investigated using transwell cultures.

Figure 4.1: (A) Electrospun galactosylated, 3-Mc loaded PCLEEP nanofiber scaffold
fabrication scheme. (B) Chemical structures of 3-methylcholanthrene (3-Mc) and 1-O(6’-aminohexyl)-D-galactopyranoside (AHG), used in the functionalization of the
PCLEEP nanofiber scaffold.

64

Chapter 4

4.3

Experimental Methods
All chemicals were purchased from Sigma-Aldrich unless otherwise stated.

4.3.1

Fabrication of Dual-Functional Nanofiber Scaffolds
As illustrated in Fig. 4.1, the dual biofunctional electrospun nanofiber scaffold

was systematically assembled through the layering of two nanofiber meshes that aremodified differently with different bioactive molecules. The following subsections
describe in detail the nanofiber scaffold modification and assembly process.
4.3.1.1

Electrospinning of Undoped Nanofiber Mesh

Detailed protocols for undoped PCLEEP nanofiber layer fabrication can befound in Chapter 3.3.1. The collection time for each mesh was 3 min.
4.3.1.2

Poly(acrylic acid) Grafting of Undoped Nanofiber Mesh

The undoped PCLEEP nanofiber layer was subsequently grafted withPoly(acrylic acid) (PAAc). Detailed protocols for PCLEEP nanofiber PAAc grafting
can be found in Chapter 3.3.1.1. 5% AAc solution was used for the grafting reaction.
The carboxylic acid group density on the meshes was tested to be between 50-65
nmol/cm2, using Toluidine Blue O staining method [33].
4.3.1.3

Electrospinning of 3-Mc Loaded Nanofiber Mesh

PCLEEP copolymer was dissolved in 7:3 dichloromethane / methanol solvent
mixture at 8.0 wt% concentrations in 2-mL centrifuge tubes. 3-Mc was then added tothe polymer solution at 0, 0.1, 1.0, 5.0 and 8.0 wt% of the amount of PCLEEP
copolymer in each tube. After 3-Mc had dissolved and thoroughly mixed, the 3-Mc
loaded polymer solutions were then electrospun into nanofiber meshes using the same
electrospinning parameters as the fabrication of undoped PCLEEP nanofibers (Fig.
4.1). However, the collection time was reduced to 1.5 min to produce a thinner mesh
65

Chapter 4compared to the undoped PCLEEP nanofiber mesh. The 3-Mc loaded nanofiber meshwere subsequently cut and immobilized over each PAAc-grafted undoped PCLEEP
nanofiber mesh. Thus, a carboxylic acid functionalized, 3-Mc loaded 2-layercomposite nanofiber scaffold was formed.
4.3.1.4

Galactosylation of Composite Nanofiber Scaffold

To make the composite nanofiber scaffold specific for hepatocyte attachment andaggregation, we further galactosylate the scaffold by conjugating 1-O-(6’Aminohexyl)-D-galactopyranoside (AHG) onto the carboxylic acid functionalized
layer using carbodiimide cross-linking chemistry. Detailed protocols for AHG bioconjugation onto PAAc-grafted PCLEEP nanofiber mesh can be found in Chapter
3.3.1.2. The morphologies of 3-Mc loaded fibers and galactosylated fibers wereimaged using a field emission scanning electron microscope (FESEM, FEI Company)
after gold sputter-coating. Representative images were analyzed using ImageJ image
processing software (http://rsb.info.nih.gov/ij/) to extract fiber diameter, pore size(area), mesh thickness and Feret’s diameter5 information. At least 250 measurements
were recorded for each analysis.
4.3.2

Hepatocyte Culture and Assays

4.3.2.1

Hepatocytes Isolation

Detailed hepatocyte isolation protocol can be found in Chapter 3.3.2.1.
4.3.2.2

Hepatocyte Attachment Assay

Freshly isolated rat hepatocytes (3.5×105) were seeded onto each 15 mm
diameter composite scaffold disks (2×105 cells/cm2), and cultured in William’s E

5Feret's Diameter is defined as the longest distance between any two points along the selection
boundary, also known as the caliper length.

66

Chapter 4

medium supplemented with 0.5 mg/mL BSA, 10 ng/mL EGF, 500 ng/mL insulin, 15
nM dexamethasone, 50 ng/mL linoleic acid, 2 mM L-glutamine, 100 units/mL
penicillin, 0.1 mg/mL streptomycin, 50 pM ZnSO4 and 100 nM CuSO4. For controls,
hepatocytes were also cultured on single layer galactosylated nanofiber scaffolds
without 3-Mc loaded, as well as on tissue culture polystyrene surface (TCPS). Cells
were cultured in a 37oC incubator fed with 5% CO2. After 3 h of incubation, allunattached cells were washed off by gentle washing using culture medium, andcollected by centrifugation (2,000 rpm). The cell pellet was washed once with PBS
and then lysed with cell lysis buffer (Promega).
The protein concentration in the lysate was determined using a BCA protein
assay kit (Pierce). The number of unattached cells was determined by comparing theprotein concentration in lysate with a standard curve generated with known numbersof cells. Subsequently, the attachment efficiency of hepatocytes on different scaffolds
was calculated based on the cell number of the unattached cells for each scaffold.
4.3.2.3

Hepatocyte Culture Maintenance

The attached hepatocytes on different scaffolds were cultured in supplemented
William’s E medium as described above. Fresh medium was replenished daily, andold medium was collected for albumin determination. Collected medium wascentrifuged at 4,000 rpm for 10 min and the supernatant was stored at −20oC.
4.3.2.4

Cytochrome P450 Activity Assay

Cultures were tested for the P450 enzymatic activity (P450-dependent 7ethoxycoumarin O-deethylase activity) as described previously [97,125-127], withmodifications. 3-Mc and 7-ethoxycoumarin first were dissolved in dimethylsulfoxide(DMSO) and then diluted to their working concentrations with culture medium. The

67

Chapter 4

final concentration of DMSO was below 0.3%. On Days 1, 3, and 5, the control
conditions (hepatocytes cultured on single layer galactosylated nanofiber scaffolds
without 3-Mc loaded, and on TCPS control) were pre-incubated in culture medium
containing 0.05 mM 3-Mc for 24 h to induce cytochrome P450 production. Then on
Days 2, 4, and 6, all culture conditions were refreshed with culture medium
containing 0.26 mM 7-ethoxycoumarin substrate. After 5 h incubation, the 7ethoxycoumarin-containing medium was collected and the cultures were refreshedwith normal culture medium. The medium samples collected were stored at −20oC
andsubsequently

assayedfor

7-hydroxycoumarin(HCOD)

using

HPLC.

Subsequently, the samples are processed for HPLC analysis as described in Chapter
3.3.2.6.
4.3.2.5

Albumin Secretion Assay

Detailed protocols for the albumin secretion assay can be found in Chapter
3.3.2.4.
4.3.2.6

Transwell Cultures

Figure 4.2: Scaffold condition illustration for transwell experiment. (1) Galactosylated
nanofiber scaffold, (2) 8.0wt% 3-Mc loaded galactosylated nanofiber scaffold, and (3)Transwell scaffold configuration.

Three different scaffold conditions were tested as illustrated in Fig. 4.2: (1)Undoped galactosylated nanofiber scaffold, (2) 8.0 wt% 3-Mc loaded galactosylated
68Chapter 4

nanofiber scaffold, and (3) a transwell configuration where the bottom partition(containing an undoped galactosylated nanofiber scaffold) is separated from the top
partition (containing a 8.0 wt% 3-Mc loaded nanofiber mesh) through a 400 nm poresize polyester membrane transwell insert (Corning). For the third condition,
hepatocytes were seeded onto the bottom partition only, so the cells have no physicalcontact with the 3-Mc loaded nanofiber mesh in the upper partition. The hepatocytes
were cultured in supplemented William’s E medium and cultures on days 2, 4, and 6
were tested for the P450 enzymatic activity as described above.
4.3.3

Statistical Analysis
All values were expressed as mean ± standard deviation (SD). Statisticaldifferences were determined by two-tailed student’s t-test.

4.4

Experimental Results and Discussion

4.4.1

Dual-Functional Nanofiber Scaffold Characterization
A schematic of scaffold fabrication process was illustrated in Fig. 4.1. Briefly, athicker undoped PCLEEP nanofiber mesh and a thinner 3-Mc loaded PCLEEP
nanofiber mesh were separately fabricated through electrospinning. After the undoped
mesh was grafted with poly(acrylic acid), the 2 meshes were then stacked together,with the 3-Mc loaded mesh forming the top layer. Subsequently, a galactose ligand
was conjugated onto the poly(acrylic acid) grafted layer, thus creating agalactosylated, 3-Mc loaded composite nanofiber scaffold.
Image analysis of the thicker undoped mesh showed a fiber diameter distributionof 730 ± 270 nm (Fig. 4.3A), while similar analysis of the thinner 3-Mc loaded meshshowed comparable fiber diameter distribution of 760 ± 280 nm (Fig. 4.3B). 3-Mc
69

Chapter 4

loading at all concentrations did not affect the resultant fiber diameter (p > 0.05
compared with undoped fibers). This result was expected as 3-Mc is an uncharged
molecule, and it was established extensively in literature that only charged molecules
can significantly affect the electrospun fiber diameters [12,41-44]. Image analysis of
the thinner 3-Mc loaded mesh also showed a mesh thickness of 1−3 fiber diameters
thick, with a wide through-pore size distribution of 7.4 ± 7.2 µm2 and through-pore
Feret’s diameter of 2.7 ± 1.3 µm (Fig. 4.3B).

Figure 4.3: SEM images of electrospun PCLEEP nanofiber mesh layers: (A) Denser
bottom layer that was functionalized with AHG ligand; (B) 3-Mc loaded (5 wt% of
PCLEEP) nanofiber top layer that will be stacked over the denser bottom layer. Fibers in
the top layer were less dense and numerous through-pores were present to facilitate
hepatocyte interaction with the galactosylated bottom layer.

It is worth noting that our initial attempt was to create a single galactosylated 3Mc loaded electrospun nanofiber mesh layer instead of a dual layer. However, wewere unable to achieve this because the UV-initiated poly(acrylic acid) grafting postelectrospinning step will also modify the 3-Mc loaded in the fiber, rendering it nonbioactive. Thus a two layer stacking fabrication approach was necessary in this case.
An obvious advantage of the two-layered nanofiber scaffold was that it allowed thetitration of two different types of cues independently.

70Chapter 4

4.4.2

Hepatocyte Attachment Efficiency
Hepatocytes cultured on various galactosylated 3-Mc loaded composite

nanofiber scaffolds (gnPCLEEP with 0 to 8% 3-Mc) exhibited similarly highhepatocyte attachment efficiency (76% ± 2.3%) 3 h after cell seeding (Fig. 4.4). The
3-Mc concentration in nanofiber scaffold did not significantly influence cellattachment efficiency (p > 0.05 comparing the cell attachment efficiencies of allcomposite scaffolds conditions). Without surface galactosylation (TCPS control), theattachment efficiency was poor (30%). Cell attachment was highest (84%) for single
layer galactosylated scaffolds (gnPCLEEP control), suggesting that for the composite
scaffolds, the upper non-galactosylated layer could slightly hinder cell attachment (p< 0.05 compared to cell attachment efficiencies of composite scaffold conditions).
Obviously, this effect will depend on the thickness and pore size of the upper layer. In
this study, the upper layer was thin and highly porous (Fig. 4.3B) and the hepatocytes
could still interact with the lower galactosylated nanofiber layer.

Figure 4.4: Hepatocyte attachment on various galactosylated 3-Mc loaded composite
nanofiber scaffolds (gnPCLEEP (0.0-8.0)% 3-Mc), single layer galactosylated scaffolds(gnPCLEEP control) and TCPS control 3 h after cell seeding. Dotted line represents
combined mean of gnPCLEEP (0.0-8.0)% 3-Mc conditions. Data are means ± SD, n = 6.
* indicates p < 0.05.
71Chapter 4

4.4.3

Cytochrome P450 Function

Figure 4.5: Cytochrome P450 function of hepatocytes at various time points normalizedagainst the total number of attached cells. Hepatocytes were cultured on galactosylated
3-Mc loaded composite nanofiber scaffolds, on single layer galactosylated scaffolds in
3-Mc containing medium (gnPCLEEP 3-Mc med. control) and on TCPS controls. Data
are means ± SD, n = 3. * indicates p < 0.05.

On galactosylated 3-Mc loaded composite scaffolds, the hepatocyte cytochrome
P450-dependent 7-ethoxycoumarin O-deethylase activity was strongly correlated withthe 3-Mc concentration loaded into the scaffolds (Fig. 4.5). The hepatocyte
cytochrome P450 activity increased with 3-Mc concentration, with the highest HCOD
production rate recorded at 7.0 pg/cell/h for gnPCLEEP 8.0% 3-Mc scaffold condition
on day 2. This rate is significantly higher (p < 0.05) than the gnPCLEEP 3-Mc
medium control condition on day 2 (4.2 pg/cell/h), where the 3-Mc concentration used
(0.05 mM) was the highest concentration reported in literature that will not interferewith other hepatocyte functions [97,126,127]. This result showed that drug-loaded72

Chapter 4

nanofiber was a more efficient lipophilic drug delivery vehicle as compared tosupplementation into the medium, especially at early culture time points.
Hepatocytes cultured on the galactosylated 3-Mc loaded composite scaffolds
showed decreased cytochrome P450 activity with time. This can be attributed to thegradual loss of 3-Mc from the nanofiber, which was depleted through gradual
diffusion and metabolization [129] by the attached hepatocytes. In contrast, P450
function of hepatocytes cultured on single layer galactosylated scaffold controls withdaily supplementation of 3-Mc in the medium peaked at day 4 (7.9 pg/cell/h). We
have attributed this observation to hepatocyte reorganization on the galactosylated
scaffold, which may have changed the cytochrome P450 inducing susceptibility(Chapter 3.4 & 3.5).
4.4.4

Albumin Synthesis Function

Figure 4.6: Albumin synthesis function of hepatocytes at various time points
normalized against the total number of attached cells. Dotted line represents combinedmean of gnPCLEEP (0.0-8.0)% 3-Mc and gnPCLEEP 3-Mc med. control conditions.
Data are means ± SD, n = 3. * indicates p > 0.05.
73Chapter 4

Hepatocytes cultured on non-galactosylated (TCPS) surfaces showed poorer
albumin synthesis function as compared to hepatocytes cultured on galactosylated
scaffolds (Fig. 4.6). In addition, hepatocytes cultured on galactosylated composite
scaffolds with no 3-Mc loaded (gnPCLEEP 0% 3-Mc), galactosylated 3-Mc loaded
composite scaffolds (gnPCLEEP (0.1-8.0%) 3-Mc), and gnPCLEEP 3-Mc medium
control all showed comparable albumin secretion profiles (p > 0.05) throughout theculture (Fig. 4.6). This highlights that the hepatocyte albumin synthesis function was
not affected by the various 3-Mc concentrations loaded in composite scaffolds.
4.4.5

Mechanism of 3-Mc Transport from Nanofiber to Cell
Two 3-Mc transport mechanisms were possible: (1) 3-Mc could be transported

by first through diffusion from fiber into the medium, and then transported from themedium into the hepatocytes; and/or (2) 3-Mc could be directly transported throughdiffusion from the 3-Mc fibers to the hepatocyte through fiber − hepatocyte
membrane contact. A transwell culture was then set up to investigate the mode of 3Mc transfer to hepatocytes (Fig. 4.7).
Results show that for hepatocytes cultured in the transwell condition(gnPCLEEP 8.0% 3-Mc transwell), the cytochrome P450 function remained low (1.31.6 pg/cell/h) but consistently higher than the non 3-Mc scaffold culture (gnPCLEEP
control, 0.4-0.8 pg/cell/h) throughout the entire culture period, indicating that theformer 3-Mc transport mechanism may only play a minor role in 3-Mc delivery. Thisobservation maybe due to 3-Mc low solubility (1.07×10-8 M) under aqueous condition[130,131], which made dissolution into the medium limited. On the other hand,
hepatocytes cultured on the dual functional gnPCLEEP 8.0% 3-Mc scaffold expressed
cytochrome P450 function (3.2-6.3 pg/cell/h) that was significantly higher (p < 0.05)
74Chapter 4

than that of gnPCLEEP 8.0% 3-Mc transwell condition throughout the entire culture
period, indicating that the latter 3-Mc transport mechanism as the more dominant 3Mc delivery route. This result suggests that due to the adhesion of hepatocytes on thegalactosylated nanofibers, the hepatocytes had direct contact with the 3-Mc loaded
fibers; this cell − 3-Mc fiber contact might have played an important role infacilitating the diffusion of 3-Mc from the fibers to the hepatocytes.

Figure 4.7: Cytochrome P450 function of hepatocytes at various time points normalizedagainst the total number of attached cells. For gnPCLEEP 8% 3-Mc transwell condition,
hepatocytes were cultured without physical contact with the 8.0% 3-Mc mesh. Data aremeans ± SD, n = 3. * indicates p < 0.05.

4.5

Concluding Remarks
By taking advantage of the porous and layer-forming properties of electrospun

nanofibers, we had designed a dual functional scaffold that induces two different
biological responses from hepatocytes. Galactose bio-functionalization on nanofibers
resulted in a scaffold that can induce hepatocyte adhesion and re-organization, while
3-Mc loading into the fiber, working together with fiber galactosylation, resulted in ahepatocyte bioactive scaffold that can also regulate the hepatocyte cytochrome P450
function. In essence, this hepatocyte-drug delivery model can be modified to deliver
75

Chapter 4

many other lipophilic bioactive molecules [130,131] that may have therapeutic effects
for liver cell cultures. By careful understanding of cell-substrate interactions andbioactive molecules effects of other cell models, we could extend this multi-functional
scaffold strategy to induce synergistic cellular responses for other cell culture
applications.
In the following chapter, we will further investigate the efficacy of this nanofiber
bio-functionalization strategy with another cell culture system: hematopoietic stem /progenitor cell expansion. This time, using amine molecule bio-functionalization, we
will explore the synergistic effects that nanofiber topography and surface immobilizedbiochemical cues play in enhancing cell-scaffold interactions and regulating cellularfunctions like cell proliferation and multipotency phenotype maintenance.

76

Chapter 5

CHAPTER FIVE
Aminated Nanofiber Scaffolds Enhance Adhesion and Expansion of
Human Umbilical Cord Blood Hematopoietic Stem/Progenitor Cells

5.1

Summary
Interaction between hematopoietic stem/progenitor cells (HSPCs) and their extra

cellular matrix components is an integral part of the signaling control for HSPC
survival, proliferation and differentiation. We hypothesized that both substrate
topographical cues and biochemical cues could act synergistically with cytokine
supplementation to improve ex vivo expansion of HSPCs.
In this study, we compared the ex vivo expansion of human umbilical cord blood
CD34+ cells on unmodified, hydroxylated, carboxylated and aminated nanofibers andfilms. Results from 10-day expansion cultures showed that aminated nanofiber meshand film were most efficient in supporting the expansion of the CD34+CD45+ cells(195-fold and 178-fold respectively), as compared to tissue culture polystyrene (50fold, p < 0.05). In particular, aminated nanofiber meshes supported a higher degree ofcell adhesion and percentage of HSPCs, as compared to aminated films. SEM imaging
revealed the discrete colonies of cells proliferating and interacting with the aminatednanofibers.
This study highlights the potential of a biomaterials approach to influence theproliferation and differentiation of HSPCs ex vivo.

77

Chapter 5

5.2

Introduction
Umbilical cord blood is a promising alternative source of hematopoietic

stem/progenitor cells (HSPCs) for allogeneic and autologous hematopoietic stem celltransplantation for the treatment of a variety of hematological disorders and as asupportive therapy for malignant diseases [132-140]. However, the low number of
HSPCs obtainable from a single donor restricts its widespread application as a viable
source of transplantable hematopoietic cells in adults [141-145]. As the success of
HSPC transplantation is dependent on both the cell dose and the pluripotency of thecells that are transplanted, an efficient and practical ex vivo expansion strategy isnecessary to produce sufficient quantity of HSPCs that can engraft and sustain longterm hematopoiesis for adult patients.
In conventional expansion culture, HSPCs are generally regarded as suspension
cells and numerous protocols implement HSPC suspension culture in the presence ofvarious combinations of early acting cytokines [132-139,173-177]. They are often
performed in flasks or bags that provide no micro-architecture for cellular interaction
with the substrate. However, it is generally accepted that the native bone marrow
microenvironment provides a complex 3-D meshwork of extracellular matrix (ECM)
that serves as a stem cell niche to regulate HSPC functions [137-140]. A growing
body of evidence suggests the importance of surface chemistry as well as
topographical features on the rate of HSPC proliferation and CD34+ cell expansion[178-185]. These results suggest that biochemical as well as topographical cues could
be actively involved in dictating the proliferation and differentiation of cultured
HSPCs.
Electrospinning has recently been employed as a versatile technique to produce
polymeric fibrous substrates for cell culture and tissue engineering applications [1478Chapter 5

27,63-66]. Several studies have shown that these fibrous scaffolds can enhancecellular responses like cell adhesion and cell phenotype maintenance [14-27,63-66].
In the previous chapters, we have demonstrated that polymeric nanofiber mesh withsurface-conjugated galactose ligands stimulate the formation of hepatocyte spheroids
that engulf the modified nanofibers, resulting in an integrated spheroid-nanofiber
construct. This suggests that the functionalized nanofiber mesh enhanced hepatocytesubstrate interaction.
This observation prompted us to investigate whether the nano-topographical cuesand various chemical cues on the nanofiber surface can synergistically influence
HSPC adhesion, proliferation and multipotency phenotype maintenance. As the firstof a series of studies, we examined HSPC expansion on surface-functionalized
polyethersulfone (PES) nanofiber meshes and PES films. PES has been widely used
as a non-degradable hollow fiber material in bioreactors for culturing mammaliancells [186,187]. It has been chosen as the substrate material because of its ease ofprocessibility: (1) it can be dissolved and electrospun into nanofibers using mild
solvents such as DMSO (Fig. 2.3); (2) PES membranes are commonly surfacemodified through UV or plasma treatments [188]; and (3) the polymer is stable insolvents like acetonitrile, in which amination reactions can be carried out efficiently.
In this study, PES nanofibers and films were surface-conjugated with thesimplest set of functional groups: carboxylic, hydroxyl and amino groups (Fig. 5.1).
These set of surface functional groups have been shown to mediate different patternsof focal adhesion by immature osteoblast-like cells and different degrees ofdifferentiation [189]. In this chapter, we have demonstrated that these surface-boundfunctional groups effected different levels of HSPC adhesion, proliferation, and79

Chapter 5

multipotency phenotype maintenance. In addition, the aminated nanofiber mesh
induced the formation of unique cell colonies.

Figure 5.1: PES scaffold surface modification scheme.

5.3

Experimental Methods
All chemicals were purchased from Sigma-Aldrich unless otherwise stated.

5.3.1

Fabrication of PES Nanofiber Scaffolds
Polyethersulfone (PES) granules (Mw: 55,000) was purchased from Goodfellow

Cambridge Limited. PES pellets were dissolved in dimethylsulfoxide (DMSO) at 20
wt% concentration and placed in a plastic syringe fitted with a 27G needle. A syringe
pump (KD Scientific) was used to feed the polymer solution into the needle tip. The
feed rate of the syringe pump was fixed at 0.3 mL/h. The PES nanofiber meshes were
fabricated by electrospinning at 13 kV using a high voltage power supply (Gamma
High Voltage Research). Nanofibers were collected directly onto grounded 15 mm
diameter glass coverslips (Paul Marienfeld) located at a fixed distance of 160 mm
from the needle tip, over a collection time of 25 min. PES films were fabricated by
dip-coating glass in 10 wt% PES in DMSO. The deposited nanofiber and film samples
were washed thoroughly in distilled water and then in ethanol to remove any residual
DMSO, and subsequently dried and stored in a desiccator.

80

Chapter 5

5.3.1.1

Surface Grafting of Scaffolds with Poly(acrylic acid)

Acrylic acid (AAc) (Merck) was distilled and stored at -20°C prior to use.
Poly(acrylic acid) (PAAc) was grafted onto the PES nanofiber mesh surface by photopolymerization, as described previously in Chapter 3.3 with slight modification on thegrafting conditions. Briefly, samples were immersed in aqueous solution containing
3% AAc solution and 0.5 mM NaIO4 in a flat-bottom glass container. The
temperature of the solution was maintained at 8°C by cooling the container in a coldwater bath. The samples were then exposed to UV from a 400 W mercury lamp
(5000-EC, Dymax) for 2 min at a distance of 25 cm. The PAAc-grafted meshes werethen thoroughly washed with deionized water at 37°C for over 36 h to remove any
ungrafted PAAc from the surface of the scaffold and dried in a storage desiccator.
5.3.1.2

Amination of Poly(acrylic acid) Grafted Scaffolds

The PAAc-grafted PES nanofiber mesh and films were further conjugated with
ethylenediamine (EtDA) using a 2-step carbodiimide cross-linking method (Fig 5.4).
Briefly, each scaffold was first gently shaken in 2 mL acetonitrile containing 50 mM
N-hydroxysuccinimide (NHS) and 50 mM dicyclohexylcarbodiimide (DCC). After 6h, the reaction solution was carefully aspirated and each scaffold was immediately
immersed into 2 mL acetonitrile containing 0.03 mmol EtDA. After 12 h, the reaction
solution was carefully aspirated and each scaffold was thoroughly washed in absoluteethanol to remove any dicyclohexyl urea (DCU), which is a by-product of the
conjugation reaction.
As for control, several PAAc-grafted PES nanofiber meshes were hydroxylated
instead, by conjugation with ethanolamine using the same modification protocol as
described above. All substrates were subsequently sterilized in 70% ethanol, thenloaded into 24-well tissue culture plates (Nunc) and stored in sterile PBS until use.
81

Chapter 5

5.3.1.3

Surface Analysis of PES Scaffolds

Surface amine density was quantified according to the method described by
Kakabakos et al. [190]. Briefly, primary amino groups on the substrates were first
converted to sulfhydryl groups through reaction with excess 2-iminothiolane (Pierce).
The surface sulfhydryl groups were then determined using a BCA assay kit (Pierce)using L-cysteine to generate a standard curve.
Surface wettability of the various substrates was characterized by measuring thewater contact angle at room temperature using a video contact angle goniometer(Advanced Surface Technology).
Samples of unmodified and aminated PES nanofiber meshes were also imaged
using a field emission scanning electron microscope (FESEM, FEI Company) for
detection of any morphology changes caused by the entire amination process. Fiber
diameters were measured by analyzing representative SEM mages of nanofibers using
NIH ImageJ software (http://rsb.info.nih.gov/ij/). At least 250 measurements wererecorded for each analysis.
5.3.2

Hematopoietic Stem Cell Culture and Assays
Frozen human umbilical cord blood CD34+ HSPCs were purchased from

AllCells, which were obtained from normal volunteers participating in an Institute
Reviewing Board (IRB) approved donor program (AllCells). The CD34+ purity in the
HSPC was determined to be 98% by flow cytometry and the viability was determined
to be more than 97% by Trypan blue. Purified recombinant human stem cell factor(SCF), Flt-3 ligand (Flt3), thrombopoietin (TPO) and IL-3 was purchased from
Peprotech Inc. The StemSpanTM serum-free expansion medium and MethoCult GF+
H4435 medium were all from StemCell Technologies.

82

Chapter 5

5.3.2.1

Ex Vivo Hematopoietic Expansion Culture

For ex vivo HSPC expansion cultures, different substrates were secured at thebottoms of wells of a 24-well tissue culture plate. Six hundred HSPCs were seeded
onto each scaffold in 0.6 mL StemSpan™ serum-free expansion medium, which
consists of 1% BSA, 0.01 mg/mL recombinant human insulin, 0.2 mg/mL humantransferrin, 0.1 mM 2-mercaptoethanol and 2 mM L-glutamine in Iscove’s MDM,
supplemented with 0.04 mg/mL low density lipoprotein (Athens Research andTechnology Inc.), 100 ng/mL SCF, 100 ng/mL Flt3, 50 ng/mL TPO and 20 ng/mL IL3. Cells were cultured at 37°C in an atmosphere containing 5% CO2 for 10 days
without medium change. Similar cultures were also performed on tissue culture
polystyrene surface (TCPS), which serve as a positive control in this study. In total, 8surface conditions were tested: TCPS, PES-(unmodified, carboxylated, aminated)films and PES-(unmodified, carboxylated, hydroxylated, aminated) nanofiber meshes.
Cells were harvested after 10 days of expansion. All substrates were washedonce with non-trypsin cell dissociation solution and twice with 2% FBS Hanks' buffer
at 5-10 min intervals between each wash. The cell suspensions collected were thenconcentrated through centrifugation at 500×g for 10 min. Aliquots of the concentrated
cells were then used for cell counting by a hematocytometer, flow cytometry analysis,
as well as for colony-forming cell assays.
5.3.2.2

Flow Cytometry

Fluorescently labeled antibodies for CD34 and other cell surface markers (CD13,
CD15, CD19, CD38, CD45 and GlyA) were purchased from BD Biosciences.
Fluorescently labeled antibodies for CD41 were purchased from Dako. Cell aliquots
were incubated at 4°C for more than 30 min in 2% FBS Hanks’ buffer in the presence83Chapter 5of various antibody combinations. After antibody staining, the cells were washedtwice using Hanks’ buffer and fixed in 1% paraformaldehyde.
Cells were analyzed by triple-color flow cytometry on a FACSCalibur analyzer(BD Biosciences). Relevant isotype controls were also included to confirm specificity
and for compensation setting. At least 20,000 events were acquired. The MilanMulhouse gating method was used for cell enumeration [151], where a double gating(CD34+CD45+) strategy was used to identify the primitive hematopoietic progenitor
cell population in the ex vivo expansion cultures. The CD34 marker is generally
expressed by primitive hematopoietic progenitor cells, while CD45 marker isexpressed on all cells of hematopoietic origin with the exception of red blood cells
and their immediate precursors.
5.3.2.3

Colony-Forming Cell Assay

Aliquots of expanded cells from each scaffold condition in the expansion
cultures were suspended into 3.3 mL of MethoCult GF+ H4435 medium (StemCell
Technologies) and the cell suspension was plated onto two 35mm culture dish (1.1mL each) as instructed in the procedure manual by StemCell Technologies. Duplicate
assays are performed for each condition. The culture dishes were then incubated at
37°C, 5% CO2 for 14 days, after which the number of erythropoietic colonies[erythroid burst-forming units (BFU-E)], granulopoietic colonies [granulocytemacrophage CFU (CFU-GM)], and multilineage colonies (CFU-GEMM) were
determined by manual counting under an inverted microscope. Positive colonies arescored on the basis of an accumulation of 20 or more cells. As a control, freshly
thawed HSPCs were also evaluated for colony-forming potential.

84

Chapter 5

5.3.2.4

Preparation for Scanning Electron Microscopy

Selected cultures samples were gently rinsed with PBS, fixed with 3%
glutaraldehyde for 30 min at 20°C, and post-fixed with 1% osmium tetraoxide foranother 15 min at 20°C. Samples were then dehydrated using a graded series of
ethanol

(25%,

50%,

70%,

90%,

95%,

100%,

100%)

followedbyhexamethyldisilazane drying. The samples were mounted onto aluminum stubs andgold sputter-coated before viewing under FESEM.
5.3.2.5

Preparation for Laser Scanning Confocal Microscopy

Selected culture samples were gently rinsed with Hanks’ buffer and fixed with
1% formaldehyde for 10 min at 20°C and immediately washed with 2% FBS Hanks’buffer. Samples were then incubated with PE-labeled CD34 antibodies in 2% FBS
Hanks’ buffer for >30 min at 4°C. For nuclear staining, Syto16 (Invitrogen) was used.
Fluorescent images were taken using a laser confocal microscope (Leica).
5.3.3

Statistical Analysis
All data were presented as mean ± standard deviation (SD). The statisticalsignificance of the data obtained was analyzed by the Student’s t-test. Probability
values of p < 0.05 were interpreted as denoting statistical significance.
5.4

Experimental Results

5.4.1

Modification of PES Substrates and Surface Characterization
Nonwoven PES nanofiber meshes were prepared by electrospinning. Parametersthat significantly influence the diameter, consistency and uniformity of theelectrospun PES fibers included PES concentration in DMSO, applied voltage andneedle-collector distance. These parameters were adjusted until unbeaded and uniform
fibers could be obtained. The optimal conditions for obtaining such PES nanofiber85

Chapter 5

meshes were described in the Methods section. Under the optimized condition, fibers
with an average diameter of 529 ± 114 nm were obtained (Fig. 5.2A & 5.2C). PES
films were prepared as a 2-D control by dip-coating glass coverslips with a diameterof 15 mm. The film surface exhibited submicron bumps and the average film
thickness was 22.5 ± 3.9 µm as analyzed from SEM images of freeze-fractured PES
films.
The PES fiber meshes and films were first carboxylated by UV-initiatedpoly(acrylic acid) grafting. Amino or hydroxyl groups were subsequently introduced
to the fiber or film surfaces by reacting ethylenediamine or ethanolamine with thesurface carboxylic acid groups using carbodiimide chemistry. SEM images comparing
unmodified and aminated PES nanofiber mesh (Fig. 5.2B) did not show any
significant morphological difference, indicating that the modification steps did notcause significant degradation/ablation of PES.

Figure 5.2: SEM images of electrospun PES nanofiber mesh: (A) unmodified; (B) after
surface conjugation with ethylenediamine. (C) Fiber diameter distribution profile of PES
fibers, electrospun from a 20 wt% PES in DMSO solution (at least 250 measurements
taken).

Table 5.2 show that after PAAc grafting, the contact angle of PES film dropped
from 76º to 53º, suggesting an increased surface wettability. The wettability further
increased after amination (contact angle was 7º for aminated PES film). The contact
86

Chapter 5

angle of the unmodified nanofiber mesh (133º) was higher than PES film (76º).
However, the contact angle of PES nanofiber meshes decreased from 133º to 0º after
PAAc grafting. Similar observations on the wettability of modified nanofiber surfaces
were also reported by Fujihara et al. [191]. After carboxylation, the micropores in therelatively more hydrophilic nanofiber mesh exerted a capillary effect that imbibed thewater droplet into the scaffold [192,193]. The aminated and hydroxylated PES fiber
meshes also exhibited an un-measurable contact angle (~ 0º). The density of
conjugated primary amino groups on aminated PES nanofiber and film was between
40−60 nmol/cm2, as quantified by the Kakabakos’ method [190]. All other surfaces
showed a background amine density of < 5 nmol/cm2.
Table 5.1: Characterization of surfaces modified with various functional groups.

Surface

Water contact angle

Primary amine group density(deg.)(nmol/cm2)

TCPS

56.0 ± 1.4

0.4 ± 0.3

PES film

76.2 ± 5.1

0.9 ± 0.5

PES carboxylated film

52.9 ± 7.3

3.4 ± 0.8

PES aminated film

7.2 ± 2.7

50.1 ± 12.5

PES fiber

133.1 ± 1.8

1.3 ± 1.2

PES carboxylated fiber

N.D.*

4.0 ± 0.0

PES hydroxylated fiber

N.D.*

1.8 ± 0.8

PES aminated fiber

N.D.*

56.2 ± 12.6

Data shown are means ± SD of triplicate surfaces.
* N.D. The contact angle was too low (~0º) to be measured accurately.

5.4.2

Ex Vivo HSPC Expansion on Various PES Substrates
Efficiency of various substrates (unmodified and modified films and nanofiber

meshes) in supporting HSPC expansion was evaluated in a 10-day expansion culture.
87

Chapter 5

Fig. 5.3 shows the total nucleated cell fold expansion and CD34+CD45+ cell foldexpansion after a 10-day expansion culture on different substrates.

Figure 5.3: Fold expansion of total nucleated cells and CD34+CD45+ cells following a
10-day culture of 600 human cord blood HSPCs on different substrates. Total cellnumbers were determined by hematocytometer cell counting, while CD34+CD45+ cells
were determined by FACS analysis at the end of culture. Data shown are means ± SD of
3–8 independent experiments, each conducted in triplicates. Unmodified, carboxylated,hydroxylated and aminated conditions were designated as “unmod.”, “COOH”, “OH”
and “NH2” respectively.

Cells harvested from the expansion cultures showed greater than 95% viability inall culture conditions. Noticeable differences were observed in the cell proliferation
response of HSPCs to the different substrates. HSPCs cultured on unmodified,carboxylated and hydroxylated PES nanofiber meshes and films yielded low
proliferation (85- to 293-fold expansion) of total nucleated cells compared to that
cultured on TCPS (850-fold). The CD34+CD45+ cell fraction was between 11.3% −26.1% of total cells as analyzed by flow cytometry, which corresponded to a low (1188Chapter 5

to 58-fold) CD34+CD45+ cell expansion. Although HSPCs cultured on TCPS surface
proliferated extensively (850-fold), the fraction of CD34+CD45+ cells was only 5.8%of total expanded cells, thus resulting in only about 50-fold expansion of
CD34+CD45+ cells.
In contrast, the expansion of CD34+CD45+ cells on aminated film and nanofiber
mesh was significantly better than other test groups (p < 0.05): aminated PES
nanofiber mesh yielded 751-fold expansion of total cells and 195-fold expansion of
CD34+CD45+ cells (25.9% of total cells), whereas aminated PES film yielded 859fold expansion of total cells and 178-fold expansion of CD34+CD45+ cells (20.8% of
total cells). There was no statistically significant difference in fold expansion of
CD34+CD45+ cells cultured on aminated PES nanofiber mesh, compared with that on
PES film (p > 0.05).
5.4.3

Colony-Forming Cell Assay
CFU assays (Fig. 5.4) were conducted to evaluate the fraction of primitive

progenitor cells in the expanded cultures. Consistent with total and CD34+CD45+ cell
expansion results, cells expanded on unmodified, carboxylated and hydroxylated PES
nanofiber meshes and films yielded lower total CFU counts (1071 ± 560 to 1996 ±
213) as compared to TCPS (2890 ± 450), aminated PES film (3471 ± 371), andaminated PES nanofiber mesh (3996 ± 358).
Interestingly, for aminated PES substrates, a significant difference was observedin the number of more primitive CFU-GEMM units generated by cells expanded on
film versus nanofiber mesh with 20% (704/3471) and 28% (1124/3996) of totalcolony counts, respectively (p < 0.05). TCPS on the other hand, generated only 15%89Chapter 5(433/2890) CFU-GEMM units (p < 0.05 compared with aminated film or nanofiber
mesh groups).

Figure 5.4: CFU counts generated after 14 days of culture, using the cells from the 10day expansion cultures on various substrates and from the unexpanded HSPCs. Data arenormalized to CFU number per 100 initial unexpanded HSPCs. Data shown are themean ± SD of 3–8 experiments, each conducted in triplicates. * p < 0.05; # p < 0.05.
Unmodified, carboxylated, hydroxylated and aminated conditions were designated as“unmod.”, “COOH”, “OH” and “NH2” respectively.

In contrast, TCPS generated relatively higher percentages of CFU-GM units(63%), indicating significant differentiation commitment of the TCPS-expanded cells
towards the myeloblast / monoblast lineage, as compared to both aminated PES film
and nanofiber scaffold (55% and 49%, respectively, p < 0.05 for TCPS vs. PES film,p < 0.01 for TCPS vs. PES nanofiber, p > 0.05 for PES film vs. PES nanofiber).

90

Chapter 5

5.4.4

Expanded HSPC Surface Marker Expression

Figure 5.5: Surface antigen expression on cells after 10-day ex vivo expansion ondifferent substrates. (A) Percentage of total cells that are CDX+. (B) Percentage of total
cells that are CD34+CD45+CDX+. (C) Percentage of the CD34+CD45+ cell populationthat are CD34+CD45+CDX−. “CDX” represents CD34, CD45, CD13, CD15, CD41,
GlyA, CD19 or CD38. Data shown are mean ± SD of 3–6 experiments, each conductedin duplicates. # denotes statistical significance of p < 0.05.
91Chapter 5

The lineage marker expression of the expanded cells was analyzed by flow
cytometry (Fig. 5.5). Only cells expanded on TCPS, aminated PES film and nanofiber
mesh, and hydroxylated nanofiber mesh were analyzed because they generated
sufficiently high numbers of cells for complete lineage marker expression analysis.
Lineage marker expression of freshly thawed, unexpanded cord blood HSPCs was
also analyzed as a control. In addition to the definitive human blood progenitor
markers which include CD34, CD45 and CD38, the cells were also evaluated formarkers for myeloblast / monoblast (CD13, CD15), megakaryoblast (CD41),
erythroid (GlyA) and pro-B cell (CD19) lineages. The following observations can bemade:(1) Unexpanded HSPC stocks showed high expression of CD34 (98%), CD45(99%), CD13 (92%) and CD38 (98%) (Fig. 5.5A);(2) Cells expanded on all substrates expressed negligible levels of CD19 (<1%),
whereas cell populations expressing CD15, CD41 or GlyA increased (Fig.
5.5A);(3) Cells expanded on aminated PES nanofiber showed the highest percentage(25.9% ± 8.5%) of CD34+CD45+ cells (Fig. 5.5B);(4) The CD34+CD45+ cell population of expanded cells were primarily negative
for CD41, GlyA and CD19 but co-expression of CD13 and CD15 wassignificant (Fig. 5.5C);(5) The CD34+CD45+ fraction of expanded cells displayed lower CD38
co-expression compared to unexpanded cells (Fig. 5.5C), an effect that had
been attributed to serum-free culture condition [184,194];92

Chapter 5(6) The CD34+CD45+ cell population expanded on aminated and hydroxylated
PES nanofiber meshes showed lower CD13 expression compared to cells
expanded on TCPS and aminated PES film (Fig. 5.5C).

5.4.5

Imaging of Adherent Cells on Aminated Substrates
After 10 days of expansion culture, some samples were processed for SEM andconfocal laser microscopy to detect the presence of any adherent cells on thesesubstrates. It was noted that expanded cells adhered weakly to TCPS, unmodified,carboxylated film and nanofiber, and hydroxylated PES nanofiber substrates; andmost of these cells could easily be detached with very gentle rinsing. As such, only
sparsely scattered cells could be seen under SEM. This observation confirmed theweak adhesion of HSPCs on these substrates. In contrast, on aminated PES nanofiber
mesh and film, cell adhesion was evident, although the arrangement of adherent cellson these two substrates appeared to differ greatly (Fig. 5.6).
On aminated PES nanofiber mesh, approximately 40% cells were adherent
following 3 gentle washes; distinct and circular cell colonies were abundant on themesh (Fig. 5.6A). Cell colonies ranged from 100 µm to 1.3 mm in diameter, with cells
numbering between 50 to a few thousand. In some of the larger colonies, cells could
be seen densely packed at the center but thinned out towards the periphery of thecolony. At high magnification, the adherent cells could be seen to be anchored via
numerous processes in intimate contact with the aminated nanofibers as well asprocesses from other cells, indicative of cell-fiber and cell-cell interactions (Fig. 5.6B
& 5.6C).

93

Chapter 5

Figure 5.6: SEM images of human cord blood HSPCs after a 10-day expansion culture
on aminated PES nanofiber mesh (A-C) and on aminated PES film (D-F) at various
magnifications. Abundant distinct, circular cell colonies are evident on the aminatednanofiber scaffold (black arrows). Filopodia extend from the cells and interact with theaminated nanofibers (white arrows). On aminated film, fewer cells are adherent withoutcolony formation; cells appear to attach along cracks.
94

Chapter 5

On aminated film substrates, however, about 25% total cells were adherent, but
only sparsely on the surface, compared to adherent cells on aminated nanofiber mesh(Fig. 5.6D). No discrete cell colony was evident but instead, the adherent cells
appeared to align along crevasses (defects generated during film processing) on thesurface of the film (Fig 5.6E). We also observed that most of the adherent cells along
the edges of the crevasses sent filopodia into the fissures (Fig. 5.6F). Cells on thesmooth surface were washed off by gentle rinsing with PBS.

Figure 5.7: Confocal laser microscopy images of human cord blood HSPCs after a 10day expansion culture on aminated PES nanofiber mesh. Green indicates Syto16 nuclear
staining and red indicates CD34-PE staining. (A) Fluorescent image of two
representative cell colonies stained with Syto16. (B, C) CD34+ cells can be found onthese cell colonies and they appear to concentrate around the periphery of the colonies.
“x” denotes the approximate center of the cell colony.

Prompted by the observation of the unique cell colonies on the surface ofaminated nanofiber mesh, we proceeded to investigate the CD34 antigen expression
among the adherent cell population. Indeed, confocal laser microscope imaging
confirmed that a fraction of the cells in the cell colonies showed positive staining with
CD34-PE antibody (Fig. 5.7). Interestingly, the CD34+ cells were located mostly
around the peripheries of the cell colonies. Cells at the center of the colonies appeared
to be predominantly CD34−. This suggests that HSPCs and the expanded cells were95

Chapter 5

proliferating in an outward, radial manner along the surface of aminated nanofiber
mesh, resulting in the formation of circular cell colonies.

5.5

Discussion
This report focuses on examining the effects of surface functional groups,

together with surface topography, on the proliferation and differentiation of HSPCs
under a typical expansion condition − using commercially available serum-free stem
cell media. The results show that under typical culture condition (StemSpan™ serumfree expansion medium and a cytokine cocktail), both substrate chemistry (amino vs.
hydroxyl vs. carboxyl groups) and topographical features affect the expansion
outcome.
HSPCs cultured on unmodified, hydroxylated or carboxylated PES substrates
exhibited low proliferation. In contrast, HSPCs cultured on aminated PES substrates
were able to proliferate as rapidly as those cultured on TCPS, with the advantage thatthe fold expansion of CD34+ cells on aminated substrates was more than 3.5 times
higher than that on commercial TCPS surface (Fig. 5.3); the expanded cells also
generated higher numbers of CFU-total and CFU-GEMM counts (Fig. 5.4). Theseobservations indicate that aminated substrate may play a role in facilitating HSPC
proliferation and/or maintenance of the HSPC phenotype.
One possible mechanism to account for the observed effects is that the aminatedsubstrate, being positively charged, could selectively enrich certain protein
components from the medium, which then contribute to the expansion outcome[189,195]. Keselowsky et al. [189] have shown that the functional presentation ofadsorbed fibronectin was different on hydroxylated, methylated, aminated and96Chapter 5

carboxylated surfaces, which consequently led to variations in cell adhesion anddifferentiation. It is possible that aminated PES surface mediated HSPC proliferation
by a similar mechanism − by binding critical cytokines and growth factors from themedium, and presenting them in a more effective immobilized form, thereby
mimicking a salient feature of the bone marrow hematopoietic stem cell niche [137140,196,197].
Another possible mechanism by which aminated surface enhanced HSPC
expansion and CD34+ phenotype maintenance is by its direct interaction with the
HSPCs through their surface CD34 antigen. CD34 antigen is a highly sialylated andnegatively charged glycophosphoprotein, and its expression decreases as HSPCs
become differentiated [146-156]. We therefore postulate that a positively charged“ligand” − in this case the surface-bound amine groups − could bind and engage thenegatively charged CD34 antigen, and the engagement of CD34 antigens on HSPCs
might activate downstream signaling pathways that subsequently influence fatechoices upon proliferation [150]. Tada J. et al. have shown that stimulation ofundifferentiated hematopoietic (myeloblastic leukemia cell line) KG1a cells with antiCD34 antibody induces homotypic cytoadhesion [150]. Binding of aggregating
antibody to CD34 antigens induced tyrosine phosphorylation, cell polarization andadhesion, and perhaps cell motility. Interestingly, being co-localized with F-actin, thecrosslinked CD34 “cap” is quite stable and persists on the cell surface for at least 2days after stimulation, whereas many other cell-surface molecules are rapidlyinternalized for degradation or recycling, upon stimulation. It is possible that theaminated PES surface serves the same role by engaging cell surface CD34 antigen.
The most interesting finding of this study is that surface topography also plays arole in HSPC adhesion and expansion. Aminated nanofiber mesh mediated the highest
97Chapter 5

degree of cell adhesion on substrate; and the expanded CD34+CD45+ cells on
aminated PES nanofiber mesh exhibited a lower monoblastic (CD13+) phenotype (Fig.
5.5C) and higher CFU potential (Fig. 5.4) compared to the same population of cells
expanded on aminated PES film. Moreover, the adherent hematopoietic cells on theaminated nanofiber displayed numerous filopodia and attachment sites on the fibers(Fig. 5.6A-5.6C), which might mediate cell migration that allows rearrangement of
the proliferated cells on the substrate surface. A consequence of this stronger adhesion
was that the cells proliferated on the nanofiber mesh surface in a radial and outward
planar fashion, resulting in distinct, circular colonies.
We do however observe that HSPCs did not survive on aminated surfaces in theabsence of cytokine supplementation, suggesting that aminated nanofiber substrateitself is not sufficient to induce the proliferation of HSPCs. The surface-bound aminogroups and topographical cue are therefore likely to play a supporting/synergistic rolefor cytokines and growth factors (supplemented in the medium) to influence HSPC
proliferation and differentiation.
Although the precise mechanisms by which amine group and nanofiber
topography mediate more efficient adhesion and expansion of CD34+CD45+ cells
remains to be elucidated, our data suggest a positive correlation between substrateadhesion of HSPCs with higher expansion efficiency of CD34+CD45+ cells. A recent
study suggests that HSPC-substrate adhesion is required for cell migration andhoming [157]; this HSPC-substrate adhesion might be part of the natural process
occurring in the hematopoietic stem cell niche that governs the proliferation anddifferentiation of HSPCs [198].

98

Chapter 5

5.6

Concluding Remarks
The ex vivo expansion of hematopoietic stem cells is one of the most challenging

fields in cell culture. This is a rapidly growing area of tissue engineering with manypotential applications in transfusion medicine, bone marrow transplantation or genetherapy. Over the last decade much progress has been made in understanding
hematopoietic differentiation, identification and isolation of HSPC subtypes,
discovery of cytokines and in the development of a variety of culture scaffold andbioreactor techniques. All this has led to a number of preliminary clinical trials thathighlighted the benefits that can be obtained from the use of expanded hematopoietic
cells in therapy. Moreover, as we understand the complexity and the regulation ofhematopoiesis, it becomes obvious that cultivation techniques and concepts must
constantly evolve and improve in order to expand HSPCs efficiently and effectively.
In this chapter, we demonstrated the effectiveness of surface modified
electrospun PES nanofiber mesh as potential scaffolds for ex vivo HSPC expansion
under serum-free conditions. The expansion profiles of human umbilical cord HSPCs
are evidently different following a 10-day culture on modified and unmodified
polymeric substrates with different functional groups and nanofiber topographical cue.
Among the carboxylated, hydroxylated, and aminated PES substrates and TCPS,
aminated PES substrates mediated the highest expansion efficiency of CD34+CD45+
cells and CFU potential. Aminated nanofiber mesh could further enhance the HSPCsubstrate adhesion and expansion of CFU-GEMM forming progenitor cells. Although
the underlying mechanisms remain to be elucidated, this study clearly underscores theimportance of culture substrate in influencing the proliferation and differentiation of
HSPCs.

99

Chapter 5

In the following chapter, we will seek to better understand the mechanism bywhich aminated nanofibers mediate these cellular responses by investigating theeffects of spacer chain lengths of the grafted amine groups. In addition, themultipotency maintenance potential, differentiation characteristics and engraftment
potential of the cells expanded from aminated nanofiber scaffolds will be further
investigated through short-term and long-term clonogenic assays and NOD/SCID
mouse engraftment assays.

100Chapter 6

CHAPTER SIX
Nanofiber Scaffolds Modified with Different Spacer-Length Amines
Modulate Hematopoietic Stem/Progenitor Cell Maintenance andProliferation Kinetics
6.1

Summary
We have shown in the previous chapter that aminated nanofiber scaffold caneffectively enhanced HSPC proliferation, while supporting the multipotency
phenotype (CFU-GEMM) of the cells expanded in HSPC ex vivo expansion cultures.
In this present study, the effects of the nanofiber scaffolds immobilized with aminefunctional groups of increasing 2-, 4- and 6-carbon spacer chain lengths (EtDA,
BuDA and HeDA conditions, respectively) on ex vivo expansion and maintenance of
HSPCs is investigated.
Results show that EtDA and BuDA nanofiber scaffold showed similar expansion
profiles (773- and 805-fold expansion, respectively) and the expanded cells
maintained 25.9% and 29.2% of the CD34+CD45+ phenotype, respectively.
Interestingly, HSPC proliferation on HeDA nanofiber scaffold was significantly lower(210-fold), although the CD34+CD45+ cell fraction was the highest at 41.1% of total
cells. This increased CD34+CD45+ percentage offsets the low cell proliferation andcells expanded from HeDA nanofiber scaffold exhibited similar CFU-GEMM and
LTC-IC maintenance as compared to cells expanded from EtDA and BuDA nanofiber
scaffolds, although the NOD/SCID mice engraftment potential was not as efficient.

101

Chapter 6

This study further suggests the importance of immobilized amino functionalgroups in influencing cell-scaffold interactions and modulating HSPC proliferation
kinetics and multipotency maintenance in aminated nanofiber scaffolds.

6.2

Introduction
In Chapter 5, we have demonstrated that surface covalent immobilization offunctional groups could mediate HSPC-substrate adhesion and proliferation. In
particular, we discovered in ethylenediamine-modified nanofiber scaffolds that
surface biochemical cues and topographical cues both played synergistic roles inenhancing HSPC-substrate adhesion and maintenance of HSPC proliferation andmultipotency. HSPCs proliferated well on aminated nanofiber scaffolds and the
expanded cells generated the highest maintenance of primitive CFU-GEMM forming
cells. Scanning electron microscopy imaging also revealed that the HSPCs extended
numerous uropodia that associated intimately with the aminated nanofibers andanchored the cells to the fibrous scaffold. In addition, the unique HSPC proliferation
pattern on these aminated nanofibers resulted in the formation of abundant distinct,
densely packed circular cell colonies on the scaffold surface. This observation was in
sharp contrast to HSPC interaction with aminated film, where there were only sparse
adherent cells on the film surface, and most of the adherent cells were found anchoredalong crevasses on the film surface.
The finding that surface amino groups and nanofiber topography could
synergistically act to promote the adherence of HSPCs and regulate their proliferation
is novel and unexpected. In this study, we further investigate the effect of amineconjugation on HSPC expansion, with a long term goal to better understand themechanism by which aminated nanofibers mediate these cellular responses (Fig. 6.1).
102

Chapter 6

Several studies had shown that spacer properties can affect the interaction between
cells and immobilized biofunctional molecules such as ligands [199,200], providing
motivation to investigate the effect of chain length of the grafted amines on the
proliferation rate and phenotype of cultured cord blood HSPCs. The differentiationand engraftment potential of the cells was assessed by clonogenic assays and
NOD/SCID mouse engraftment assays, respectively.

Figure 6.1: PES scaffold amination scheme with different spacer chain length amines.

6.3

Experimental Methods
All chemicals were purchased from Sigma-Aldrich unless otherwise stated.

6.3.1

Fabrication of PES Nanofiber Scaffolds
Detailed protocols for PES nanofiber scaffold fabrication can be found in

Chapter 5.3.1.
6.3.1.1

Surface Grafting of Scaffolds with Poly(acrylic acid)

The PES nanofibers were subsequently grafted with Poly(acrylic acid) (PAAc).
Detailed protocols for PES nanofiber PAAc grafting can be found in Chapter 5.3.1.1.

103

Chapter 6

6.3.1.2

Amination of Poly(acrylic acid) Grafted Scaffolds

The PAAc-grafted PES nanofiber meshes were further conjugated with 1,2ethanediamine (EtDA), 1,4-butanediamine (BuDA) or 1,6-hexanediamine (HeDA)
using carbodiimide cross-linking method (Fig. 6.1). Briefly, each scaffold was firstgently shaken in 2 mL acetonitrile containing 50 mM N-hydroxysuccinimide and 50
mM dicyclohexylcarbodiimide. After 6 h, the reaction solution was carefully aspirated
and each scaffold was immediately immersed into 2 mL acetonitrile containing 0.03
mmol EtDA, BuDA or HeDA. After 12 h, the reaction solution was carefully
aspirated and each scaffold was thoroughly washed in absolute ethanol to remove any
dicyclohexyl urea, which is a by-product of the conjugation reaction. All substrates
were subsequently sterilized in 70% ethanol, then loaded into 24-well tissue culture
plates (Nunc) and stored in sterile PBS until use.
Surface characterization and atomic compositions of various PES nanofiber
surfaces were determined using XPS (PHI-1800, Physical Electronics). Binding
energies were referenced to the CC/CH2 C(1s) peak at 284.6 eV.
6.3.2

Hematopoietic Stem Cell Culture and Assays
Frozen human umbilical cord blood CD34+ HSPCs were purchased from

AllCells. The CD34+ purity in the HSPC was determined to be 98% by flow
cytometry and the viability was determined to be more than 97% by Trypan blue.
Purified recombinant human stem cell factor (SCF), Flt-3 ligand (Flt3),
thrombopoietin (TPO) and IL-3 was purchased from Peprotech Inc. Low density
lipoprotein (LDL) was purchased from Athens Research & Technology Inc. The
StemSpanTM serum-free expansion medium, MethoCult GF+ H4435 and MyeloCult
H5100 were all from StemCell Technologies.

104

Chapter 6

6.3.2.1

Ex Vivo Hematopoietic Expansion Culture

Six hundred HSPCs were seeded onto each scaffold. HSPCs were cultured in 0.6
mL StemSpan™ serum-free expansion medium supplemented with 0.04 mg/mL LDL,
100 ng/mL SCF, 100 ng/mL Flt3, 50 ng/mL TPO and 20 ng/mL IL-3 at 37oC, 5%
CO2 for 10 days. Similar cultures were also performed on tissue culture polystyrene
surface (TCPS), which served as a positive control in this study. In total 6 surface
conditions were tested: TCPS, unmodified PES nanofiber mesh (Unmod.),carboxylated nanofiber mesh (AAc), and nanofiber mesh aminated with EtDA, BuDA
or HeDA.
On day 10, the expanded cells were harvested and aliquoted. Briefly, allsubstrates were washed once with non-trypsin cell dissociation solution and twicewith 2% FBS Hanks' buffer at 5-10 min intervals between each wash. The cellsuspensions collected were then concentrated through centrifugation at 500 ×g for 10
min. Aliquots of the concentrated cells were then used for cell counting by ahematocytometer, flow cytometry analysis, colony-forming cell assays, long-term
culture-initiating cell assay and mouse engraftment assay.
6.3.2.2

Flow Cytometry

Fluorescently labeled antibodies for CD13, CD34 and CD45 were purchasedfrom BD Biosciences (USA). The cell samples were incubated with antibodies at 4oC
for more than 30 min in 2% FBS Hanks' buffer. After antibody staining, the cells were
washed twice with Hanks’ buffer and fixed in 1% paraformaldehyde. Cells wereanalyzed by flow cytometry on a FACSCalibur analyzer (BD Biosciences). Relevant
isotype controls were also included to confirm specificity and for compensation
setting. At least 20,000 gated events were acquired. The Milan-Mulhouse gating
method was used for cell enumeration [151].
105

Chapter 6

6.3.2.3

Preparation for Scanning Electron Microscopy

Selected cultures samples were gently rinsed with PBS, fixed with 3%
glutaraldehyde for 30 min at 20°C, and post-fixed with 1% osmium tetraoxide foranother 15 min at 20°C. Samples were then dehydrated using a graded series of
ethanol

(25%,

50%,

70%,

90%,

95%,

100%,

100%)

followedbyhexamethyldisilazane drying. The samples were mounted onto aluminum stubs andgold sputter-coated before viewing under field emission scanning electron microscope(SEM, FEI Company).
6.3.2.4

Colony-Forming Cell Assay

Detailed protocols for CFC assay can be found in Chapter 5.3.2.3.
6.3.2.5

Long-Term Culture-Initiating Cell Assay

For LTC-IC, expanded cells from each scaffold condition in the ex vivo
hematopoietic expansion cultures and freshly thawed HSPCs, which serve as controls,
were plated onto irradiated M2-10B4 murine fibroblast feeder cells in 35mm culture
dishes and cultured in MyeloCult H5100 medium as instructed in the procedure
manual by StemCell Technologies. After 5 weeks, all the cells from each dish were
harvested, and cultured according to the CFC assay as described above. LTC-IC
numbers were then calculated and normalized according to instructions in theprocedure manual.
6.3.2.6

Mouse Engraftment Assay

Non-obese diabetic/severe combined immunodeficient (NOD/SCID) mice
(Animal Resource Center, Perth, Australia) were maintained at the Biological
Resource Center (BRC), Biopolis, Singapore. All animals were handled according to
BRC regulations. 6-8 weeks old mice were irradiated at 350 cGy. Cells harvested

106Chapter 6

from 10-day ex vivo expansion cultures were mixed with 4 × 105 irradiated (1,500cGy) CD34-depleted human bone marrow cells (carrier cells) and injected into themice via the tail vein. For positive controls, 2 mice groups of 600 or 20,000 injected
unexpanded HSPCs / 4 × 105 irradiated carrier cells mix were also examined. Finally,
un-irradiated mice, irradiated mice and irradiated mice with 4 × 105 injected irradiatedcarrier cells groups were also included to serve as negative controls in this study.
Mice were sacrificed 6 weeks after cell transplantation. After euthanasia,
bilateral femora and tibia were harvested from each animal and bone marrow cells
were flushed out with 2% FBS, 5% human serum Hanks' buffer. The cells were
subsequently stained with fluorescently labeled human CD45 antibody and the red
blood cells lysed using FACS lysing solution (BD Biosciences). The percentage ofhuman hematopoietic cells in the mouse bone marrow was quantified by flow
cytometry. At least 40,000 gated events were acquired. Successful humanhematopoietic stem cell engraftment was defined by the presence of at least 0.1% of
human CD45+ cells in the NOD/SCID mouse bone marrow cell population.
6.3.3

Statistical Analysis
All data were presented as mean ± standard deviation (SD). The statisticalsignificance of the data obtained was analyzed by non-parametric Mann-Whitney test
for mouse engraftment results and Student’s t-test for all other results. Probability
values of p < 0.05 were interpreted as denoting statistical significance.
6.4

Experimental Results

6.4.1

Surface Characterization of Aminated Nanofiber Scaffolds
Nonwoven PES nanofiber meshes with an average diameter of 529 ± 114 nm

were prepared by electrospinning process. The PES nanofiber meshes were first107

Chapter 6

carboxylated by UV-initiated PAAc grafting and subsequently conjugated withdiamines with 2, 4, or 6 alkane spacers (EtDA, BuDA or HeDA respectively) using
carbodiimide cross-linking method (Fig. 6.1). XPS analysis showed the surface
elemental concentration of nitrogen on the aminated fibers to be between 11.6% −13.4% (Table 6.1), which indicated the similar conjugation efficiencies of thedifferent diamines on the nanofiber surface. The unmodified and PAAc-grafted fiberson the other hand showed background nitrogen concentrations of <0.2%.
Table 6.1: XPS elemental analysis of PES nanofiber surfaces modified withdifferent functional groups.

PES nanofiber surface

C atomic

O atomic

N atomic

S atomicratio (%)

ratio (%)

ratio (%)

ratio (%)

Unmodified

74.2

20.2

0.2

5.4

AAc

68.6

27.7

0.1

3.6

EtDA

65.2

19.8

13.4

1.6

BuDA

67.9

17.5

13.2

1.4

HeDA

71.8

14.9

11.6

1.7

In addition, the carbon XPS spectra (C1s) showed that after PAAc grafting, the
π→π* shake-up satellite region at 291.7 eV (caused by aromatic carbon species inPES) was absent in PES AAc fiber surface, and replaced with the PAAc characteristic
O−C=O region (Fig. 6.2). Subsequently, the XPS C1s spectra also showed absence of
O−C=O species for all amine conjugated fibers, indicating the complete conversion of
the surface PAAc carboxylic acid groups to amide groups during the amine
conjugation reaction.

108

Chapter 6

Figure 6.2: The XPS spectra of various modified PES nanofiber surfaces. Left panel:
Survey spectra showing the relative abundance of O, N and C elements. Right panel:
C1s spectra showing the relative abundance of the different carbon bonds.

109

Chapter 6

6.4.2

Ex Vivo HSPC Expansion on Aminated Nanofiber Scaffolds
The efficiency of the various nanofiber scaffolds for supporting HSPC expansion

was evaluated through 10-day expansion cultures. Fig. 6.3 showed the total nucleatedcell fold expansion and CD34+CD45+ cell fold expansion after a 10-day expansion
culture on different spacer amine nanofiber and control surfaces. Cells harvested fromthe expansion cultures showed >95% viability in all culture conditions. In general,
HSPCs culture on Unmod and AAc surfaces yielded the lowest proliferation of total
nucleated cells (85- and 152-fold, respectively); the CD34+CD45+ cell fraction was
13.3% and 26.1% of total cells, respectively, as analyzed by flow cytometry, whichcorresponded to a low 11- and 40-fold CD34+CD45+ cell expansion respectively.
Although HSPCs cultured on TCPS surface proliferated extensively (895-fold), theCD34+CD45+ cell fraction was only 5.9% of total cells, corresponding to a 53-foldCD34+CD45+ cell expansion.
In contrast, the expansion of CD34+CD45+ cells on EtDA, BuDA and HeDA
nanofiber mesh was significantly better than other test groups: EtDA and BuDA
nanofiber mesh showed similar expansion profiles (p > 0.05) and yielded 773- and805-fold expansion of total cells respectively (Fig. 6.3), with 25.9% (200-fold) and29.2% (235-fold) of total cells expressing the CD34+CD45+ phenotype, respectively(Fig. 6.4). Interestingly, although HSPCs proliferation on HeDA surface wassignificantly lower as compared with EtDA and BuDA surfaces (210-fold, p < 0.05),
the CD34+CD45+ cell fraction was the highest at 41.1% of total cells (86-foldCD34+CD45+ cell expansion).

110Chapter 6

Figure 6.3: Fold expansion of total nucleated cells and CD34+ cells following a 10-day
culture of 600 human cord blood HSPCs on different substrates. Total cell and CD34+
cell numbers were determined by hematocytometer cell counting and FACS analysis
respectively at the end of culture. Data are means ± SD of 3-8 independent experiments,
each conducted in triplicates.

In addition, we also observed that the CD34+CD45+ cell population expandedfrom HeDA nanofiber scaffolds co-expressed significantly lower levels of the
myeloid CD13 marker (60.3 ± 7.3 % of expanded CD34+CD45+ cell population)compared to EtDA and BuDA nanofiber scaffolds (94.2 ± 3.5 %, p < 0.05 and 92.8 ±3.8 %, p < 0.05 of expanded CD34+CD45+ cell population respectively (Fig. 6.4D).

111

Chapter 6

Figure 6.4: Representative FACS profiles (A, B) and surface marker expression
summary (C, D) of cells after 10-day ex vivo expansion on TCPS and EtDA, BuDA andHeDA nanofiber scaffolds. (A) CD45 vs. CD34. (B) CD13 vs. CD34. (C) Percentage of
total cells expressing one or multiple CD markers. (D) Percentage of the CD34+CD45+
cell population that are CD34+CD45+CD13+. Data shown are mean ± SD of 5-8
experiments, each conducted in duplicates.

6.4.3

Morphology of Adherent Cells on Aminated Scaffolds
SEM imaging was used to monitor the proliferation kinetics of the adherent

HSPC population on the nanofiber scaffolds. At selected time points during the 10day expansion culture, samples were processed for SEM to image the presence of any
adherent cells on the nanofiber scaffolds. We noted that expanded cells adhered
weakly to TCPS, Unmod, and AAc surfaces, and most of these cells could easily be

112

Chapter 6

washed off with very gentle rinsing. As such, only sparsely scattered cells remained
adherent for SEM imaging.
In contrast, on both EtDA and BuDA modified PES nanofiber mesh, HSPC
interaction and adhesion on the nanofiber surfaces were clearly evident. When SEM
imaging was performed on day 3 cultures, small pockets of adherent HSPCs could
already be observed interacting and proliferating on the aminated nanofiber mesh (Fig.
6.5A & 6.5B). The adherent HSPCs were anchored to the aminated nanofibers via
numerous uropodia radiating from the cell surface (Fig. 6.5C). Cells undergoing
division were also evident on the nanofiber surface (Fig. 6.5D). By day 8 of
expansion culture, the adherent HSPCs proliferated to form distinct, densely
populated circular cell colonies on the aminated nanofiber mesh (Fig. 6.5E & 6.5F).
The distinct circular cell colonies most likely arose from single or small clusters of
HSPCs proliferating outwards in a radial manner along the nanofiber surface. The cellcolonies ranged from 0.1 to 1.3 mm in diameter, with cells numbering between 50 to a
few thousand.
Adherent HSPCs proliferated well on EtDA (Fig. 6.6A & 6.6B) and BuDA (Fig.
6.6C & 6.6D) nanofiber surfaces to form densely populated cell colonies after 10 daysof culture, and this was mirrored by the high mononucleated cell counts (Fig. 6.3).
Conversely, the considerably lower proliferation rate on HeDA modified nanofiber
surface (210-fold; Fig. 6.3) was reflected by smaller colony size, each containing lessthan 50 cells (Fig. 6.6E & 6.6F). Besides differences in adherent cell density andcolony size, no obvious morphological differences could be discerned among theadherent cells expanded on the EtDA, BuDA, or HeDA nanofiber surfaces.

113

Chapter 6

Figure 6.5: SEM images of HSPCs after (A-D) 3-day and (E, F) 8-day cultures on PESBuDA nanofiber mesh at various magnifications. (A, B) Pockets of adherent HSPCs
were observed (white circles) proliferating on the aminated nanofiber surface. (C) Cells
exhibited numerous filopodia which were interacting with the aminated nanofibers. (D)Cell division was also observed occurring on the nanofiber surface. (E, F) Towards day
8, HSPCs proliferated to form circular colonies (black arrows) on the nanofiber surface.
114

Chapter 6

Figure 6.6: SEM images of adherent cell colonies after 10 days of expansion on PES(A, B) EtDA, (C, D) BuDA and (E, F) HeDA nanofiber mesh at various magnifications.
Colonies of densely packed adherent cells were observed on EtDA and BuDA nanofiber
surfaces. On HeDA nanofiber surfaces, adherent cells were sparsely located.

115

Chapter 6

6.4.4

HSPC Clonogenic Potential from Various Scaffolds
CFC and LTC-IC assays were conducted to evaluate the fraction of primitive

progenitor cells in the expanded cultures. The CFC results (Fig. 6.7) showed that cells
expanded from Unmod and AAc nanofiber meshes yielded lower total CFU counts(1199 and 1609 respectively) as compared to TCPS control (2890, p < 0.05).
Conversely, cells expanded from EtDA, BuDA and HeDA nanofiber meshes yieldedsignificantly higher total CFU counts (3996, 4208 and 3742 respectively) compared to
TCPS control (p < 0.05). In addition, significant differences were also observed in thenumber of primitive CFU-GEMM units generated by cells expanded on EtDA, BuDA
and HeDA nanofiber mesh with 28.1% (1124/3995), 27.6% (1163/4207) and 28.4%(1064/3742) of total colony counts respectively, compared to cells expanded on TCPS
(15.0%, 433/2890, p < 0.05). TCPS, on the other hand, generated higher percentagesof CFU-GM units (63%), indicating differentiation commitment of the TCPSexpanded cells towards the myeloblast / monoblast lineage.
Results from LTC-IC assays (Fig. 6.8) suggested that HSPCs expanded from
EtDA-, BuDA- and HeDA-scaffolds may be more primitive than those cultured oncontrol surfaces. More importantly, the LTC-IC numbers generated from these
conditions were significantly higher than that from unexpanded cells control (p <0.05), suggesting higher degree of HSPC self-renewal on the aminated nanofiber
scaffolds.
Interestingly, cells expanded from HeDA nanofiber scaffolds generated
comparatively high numbers of colony units similar to EtDA and BuDA conditions(Fig. 6.7 & 6.8), even though the total cell expansion (Fig. 6.3) was shown to be low.
We propose that the relatively high CD34+ phenotype expression of cells expandedfrom HeDA nanofiber scaffolds may have contributed to this result.
116

Chapter 6

Figure 6.7: CFU counts after 14 days of culture, using the cells from the 10-day
expansion cultures on various substrates and unexpanded HSPCs, normalized to CFU
per 100 initial unexpanded HSPCs. Data are means ± SD of 3-8 experiments, eachconducted in triplicates.

Figure 6.8: LTC-IC counts after 7 weeks of culture, using the cells from the 10-day
expansion cultures on various substrates and unexpanded HSPCs, normalized to LTC-IC
per 100 initial unexpanded HSPCs. Data are means ± SD of 2 experiments, eachconducted in triplicates. * indicates p < 0.05.

117Chapter 6

6.4.5

HSPC NOD/SCID Repopulation Potential from Various Scaffolds
To access the effect of surface modified nanofiber scaffolds on the maintenanceof HSPCs and their engraftment potential, cells harvested from 10-day expansion
cultures were injected intravenously into sub-lethally irradiated NOD/SCID mice
together with 4 × 105 irradiated carrier cells. As positive controls, 600 and 20,000(“20k” group in Fig. 6.9) unexpanded CD34+ cells were also injected into 2 groups ofmice. The presence of > 0.1% human CD45+ cells among the murine bone marrowcells after 6 weeks was used as a criterion for successful primary engraftment in thebone marrow of NOD/SCID mice.

Figure 6.9: Engraftment efficiency of human CD45+ cells in bone marrow of sublethally irradiated NOD/SCID mice transplanted with unexpanded HSPCs, cells from the
10-day expansion cultures on various substrates, and irradiated carrier cells alone.
Numbers in parentheses indicate mice survival in the different experimental groups. *indicates p < 0.05.

Based on this criterion, only cells expanded on EtDA and BuDA scaffolds, alongwith 20,000 freshly thawed uncultured cells showed positive engraftment (Fig. 6.9).
Moreover, there was statistical significance between EtDA vs. 600 cells and BuDA vs.
118

Chapter 6

600 cells groups (p < 0.05), indicating improvement of HSPC engraftment potential
following ex vivo expansion on EtDA and BuDA nanofiber scaffolds (600 cells was
the initial cell seeding number for the expansion cultures). However, the HeDA groupfailed to show positive engraftment, even though its corresponding CFC and LTC-IC
results were comparable to that of EtDA and BuDA groups.

6.5

Discussion
The present study investigates the effects of covalently grafted primary aminefunctional groups in conjunction with spacer chain length and surface nanofiber
topography on ex vivo expansion and multipotency maintenance of human umbilicalcord HSPC in serum-free culture.
We have shown in the previous chapter that both chemical and topographicalcues can modulate HSPC-substrate interaction. On top of supporting total nucleatedcell proliferation of the cultured HSPCs at rates highly comparable to that ofcommercial TCPS surface, aminated (EtDA) nanofiber scaffolds and films also
demonstrated additional benefit of enriching CD34+CD45+ cell proportion to several
fold higher than that on TCPS. Specifically, enhanced HSPC-scaffold interaction andadhesion was observed on aminated (EtDA) nanofiber scaffolds, compared withaminated film conditions. In addition, cells expanded from aminated nanofiber
scaffolds exhibited better multipotency maintenance by supporting higher percentagesof CFU-GEMM cells as compared to aminated film conditions.
Results in this chapter further confirmed the findings from Chapter 5. We
showed that cells expanded on aminated nanofiber scaffolds generated significantly
higher numbers of total CFU, CFU-GEMM units and LTC-IC counts, in contrast to
the carboxylated nanofiber scaffold, unmodified nanofiber scaffold and TCPS
119

Chapter 6

substrate conditions, which yielded various degree of reduction of these progenitor
cells (Fig. 6.7 & 6.8).
More importantly, our results indicated that the spacer (ethylene, butylene andhexylene) linkages between amino groups and nanofiber surface influenced the
expansion of HSPCs (Fig. 6.3). HSPCs cultured on BuDA-nanofiber scaffold showedsimilar expansion efficiency to that on EtDA-nanofibers. However, increasing theamine spacer length to 6-carbon alkyl chain (HeDA-nanofibers) reduced total cell
expansion by 3.8 times but increased CD34+CD45+ cell percentage by 1.5 times(41.1% of total cells). Therefore, it appears that HeDA nanofiber scaffold was mostefficient at preserving the CD34+ phenotype, but at the expense of overall cellproliferation. The outcomes of ex vivo expansion experiments are largely determinedby the balance between self-renewal and differentiation of HSPCs in culture[138,142]: Differentiation and hence, depletion of stem and progenitor cell
populations, are often accompanied by rapid proliferation of differentiated cells.
Because of this reduced total cell expansion (4-times lower than that on BuDAnanofibers, Fig. 6.3), and hence lower total cell transplantation dose, the engraftment
efficiency of cells expanded on HeDA-nanofibers in NOD/SCID mice was lower thanthat expanded on BuDA- and EtDA-nanofiber scaffolds. This result confirmed that
HSPC transplantation dose is one of the critical parameters for successful engraftment[132-136,141-145]. In addition, we also observed that the CD34+CD45+ cell
population expanded from HeDA nanofiber scaffolds expressed significantly lower
levels of the myeloid CD13 antigen compared to EtDA and BuDA nanofiber scaffolds(Fig. 6.4D). We believe that this lower CD34+CD45+CD13+ / CD34+CD45+
expression was also a possible cause of low engraftment frequency for cells expanded
on HeDA-nanofiber scaffold, given that a recently report suggests a highly positive120

Chapter 6

correlation between myeloid marker expression and the engraftment potential of
human CD34+ HSPCs [155].
SEM imaging analysis provided the direct evidence of HSPC adhesion onaminated nanofiber substrates (Fig. 6.5 & 6.6). Despite the varying degrees ofproliferation, intimate binding of cells with nanofibers was evident for all three typesof aminated nanofibers. The strikingly distinct circular colonies most likely arosefrom single or small clusters of HSPCs proliferating outwards in a radial manneralong the nanofiber surface, suggesting that the sub-micron scale feature created byelectrospun fibers provided traction and contact guidance for the dividing cells as theymigrated away from the center of the colony. Numerous threadlike processes anduropodia emanating from the cell surface [201] apparently anchored the cells to thenanofibers and likely mediated the migration of the cells.
The distinct circular colony features that remained following several sample
preparation steps prior to SEM imaging also clearly indicates the strong cell adhesion
strength of the expanded cells on the aminated nanofiber mesh. In addition, the colony
sizes and cell densities within the colonies correlated well with the total nucleated cell
expansion data: HeDA-nanofiber scaffold yielded fewer and smaller colonies
compared with EtDA- and BuDA-nanofiber scaffolds. Together with the resultspresented in the previous chapter, these SEM data suggested the importance of HSPC
adhesion in regulating proliferation and self-renewal.
We again hypothesize that the adhesion of HSPCs on aminated nanofiber
scaffolds may likely be mediated by CD34 antigen, the highly sialylated andnegatively charged glycophosphoprotein, due to electrostatic charge-chargeinteraction. Recent evidences begin to unfold the important role of CD34 antigen in121Chapter 6

regulating cell adhesion [147-150,152-154,156]. Under normal conditions, due to its
halo of negatively charged sialic acid, CD34 antigen functions as antiadhesin,preventing cell-cell adhesion of HSPCs [147,148,150,156]. However, when CD34 isbound to antibodies [147,148,150,156] or to an extracellular ligand that is yet to be
identified [148], cell-cell adhesion is enhanced, either through recruitment and“concentration” of CD34 to a cap region [150], and/or through antibody-CD34 (orligand-CD34) mediated intracellular signaling [147,148,150,156], which may result inan up-regulation of cell adhesive molecules in HSPCs [148]. This latter suggests that
it is likely that HSPCs interact with surface amino groups, either directly mediate orindirectly facilitate HSPC adhesion to the substrate. Nevertheless, the exact
mechanism by which CD34 mediated HSPC adhesion and signaling plays on HSPC
expansion and maintenance of pluripotency remains to be elucidated.

6.6

Concluding Remarks
In this chapter, we discussed the spacer effect of the surface-grafted aminogroups. Nanofiber scaffolds amine-functionalized with different spacer-lengths
modulate HSPC proliferation and phenotype maintenance differently, resulting indifferent HSPC proliferation kinetics, cell population phenotypic (CD34 and CD13
markers) expression, and also colony-forming and mouse engraftment ability. Theseobservations further suggested the importance of nanofiber topography and amino
functional group mediated cell-scaffold interactions in regulating HSPC proliferationand self-renewal.

122Chapter 7

CHAPTER SEVEN
Adhesive Cell-Scaffold Interaction through Aminated Nanofiber
Scaffold Promotes Hematopoietic Stem/Progenitor Cell Maintenanceand Lineage Commitment
7.1

Summary
We have shown in previous chapters that a combination of nanofiber topography

and immobilized amine mediated cell-substrate interactions has enhanced HSPC
adhesion on aminated nanofiber scaffolds, and generated a sub-population of highlyadhesive HSPCs proliferating along nanofiber scaffold surface. In this present study,
we compare the surface marker phenotypic and clonogenic differences of theseadherent and non-adherent hematopoietic cell populations that arise after ex vivo
expansion culture of CD34+ human umbilical cord hematopoietic stem/progenitor
cells (HSPCs) on aminated electrospun nanofiber scaffolds.
Detailed flow cytometry analysis and in vitro assay showed that the adherent cell
population expressed significantly higher percentage of CD34+CD45+ cells (43.8%),
compared with the non-adherent cell population (21.9%, p < 0.05). In addition, theadherent cell population also expressed higher percentage of CD13 myeloid marker(68.3% vs. 49.4%, p < 0.05) and lower percentage of erythroid marker (CD71high,
14.1% vs. 46.2%, p < 0.05), compared with the non-adherent cell population. CFU
assay also indicated significant commitment of the adherent population towards the
myeloblast / monoblast lineage, while the non-adherent population showed skewed
commitment towards the erythroid lineage.

123Chapter 7This study highlights the importance of cell-scaffold interactions as a new
approach in modulating HSPC multipotency maintenance and lineage commitment,
other than cytokine modulation frequently described in literature.
7.2

Introduction
We have shown in Chapters 5 that both chemical cues and topographical cues

synergistically regulate HSPC proliferation through cell-substrate interaction.
Specifically, aminated nanofiber scaffolds are efficient in supporting total nucleatedcell proliferation of cultured HSPCs at rates highly comparable to that of commercial
TCPS surface, while concurrently enriching the CD34+CD45+ cell proportion toseveral times higher than that on TCPS.
Chapter 6 highlighted the spacer effect of the surface-grafted amino groups.
Nanofiber scaffolds amine-functionalized with different spacer-lengths modulate
HSPC proliferation and phenotype maintenance differently, resulting in different
HSPC proliferation kinetics, cell population phenotypic (CD34 and CD13 markers)
expression, and also colony-forming and mouse engraftment ability.
A direct consequence of these active cell-substrate interactions is the enhanced
HSPC adhesion to aminated nanofiber scaffolds, compared with aminated films orother non-aminated surfaces (e.g. TCPS). The enhanced HSPC adhesion also resulted
in sub-populations of highly adhesive HSPCs proliferating along nanofiber scaffold
surface, forming unique circular cell colonies (Fig. 7.1). This is a unique growth
pattern that is observed for the first time.
In this chapter, we will further investigate the phenotypic differences of these
non-adherent and adherent hematopoietic cell populations expanded on aminated124Chapter 7(BuDA) nanofiber scaffolds, in an effort to further understand this HSPC-aminatednanofiber interaction.

Figure 7.1: Image of a representative adherent cell colony formed on aminated (BuDA)
nanofiber scaffold 10 days after ex vivo HSPC expansion.

7.3

Experimental Methods
All chemicals were purchased from Sigma-Aldrich unless otherwise stated.

7.3.1

Fabrication of PES Nanofiber Scaffolds
Detailed protocols for PES nanofiber scaffold fabrication can be found in

Chapter 5.3.1.
7.3.1.1

Surface Amination of PES Nanofiber Scaffolds

The PES nanofibers were subsequently grafted with Poly(acrylic acid) (PAAc).
Detailed protocols for PES nanofiber PAAc grafting can be found in Chapter 5.3.1.1.
The PAAc-grafted PES nanofiber meshes were further conjugated with 1,4butanediamine (BuDA) using carbodiimide cross-linking method. Briefly, each125

Chapter 7

scaffold was first gently shaken in 2 mL acetonitrile containing 50 mM
dicyclohexylcarbodiimide and 50 mM N-hydroxysuccinimide. After 6 h, the reaction
solution was carefully aspirated and each scaffold was immediately immersed into 2
mL acetonitrile containing 0.03 mmol BuDA. After 12 h, the reaction solution was
carefully aspirated and each scaffold was thoroughly washed in absolute ethanol toremove any dicyclohexyl urea, which is a by-product of the conjugation reaction. All
substrates were subsequently sterilized in 70% ethanol, then loaded into 24-well tissue
culture plates (Nunc) and stored in sterile PBS until use.
7.3.2

Hematopoietic Stem Cell Culture and Assays
Frozen human umbilical cord blood CD34+ HSPCs were purchased from

AllCells. The CD34+ purity in the HSPC was determined to be 98% by flow
cytometry and the viability was determined to be more than 97% by Trypan blue.
Purified recombinant human stem cell factor (SCF), Flt-3 ligand (Flt3),
thrombopoietin (TPO) and IL-3 was purchased from Peprotech Inc. Low density
lipoprotein (LDL) was purchased from Athens Research & Technology Inc. The
StemSpanTM serum-free expansion medium and MethoCult GF+ H4435 medium were
all purchased from StemCell Technologies.
7.3.2.1

Ex Vivo Hematopoietic Expansion Culture

Six hundred HSPCs were seeded onto each BuDA-conjugated nanofiber scaffold.
HSPCs were cultured in 0.6 mL StemSpanTM serum-free expansion medium
supplemented with 0.04 mg/mL LDL, 100 ng/mL SCF, 100 ng/mL Flt3, 50 ng/mL
TPO and 20 ng/mL IL-3 at 37oC, 5% CO2 for 10 days. Similar cultures were also
performed on tissue culture polystyrene surface (TCPS), which serve as a HSPC nonadhesive surface control in this study.

126

Chapter 7

7.3.2.2

Cell Harvest

Cells were harvested after 10 days of expansion culture. Briefly, TCPS substrates
and a portion of the BuDA-conjugated nanofiber scaffolds were washed once with
non-trypsin cell dissociation solution and twice with 2% FBS Hanks' buffer at 5 minintervals between each wash; and the cell suspensions collected were designated as“TCPS’ and “PES-BuDA” conditions respectively.
For the remainder of the BuDA-conjugated nanofiber scaffolds, the cells were
harvested into 2 fractions. Briefly, the scaffolds were first gently washed twice with
2% FBS Hanks' buffer at 5 min intervals between each wash to harvest the nonadherent cells; and the cell suspensions collected were designated as “PES-BuDA‘sus’ fraction”. Subsequently, the scaffolds were washed once with non-trypsin celldissociation solution and twice with 2% FBS Hanks' buffer at 5 min intervals between
each wash to harvest the adherent cells; and the cell suspensions collected were
designated as “PES-BuDA ‘adh’ fraction”.
The cell suspensions collected were then concentrated through centrifugation at
500 ×g for 10 min. Aliquots of the concentrated cells were then used for cell counting
by a hematocytometer, flow cytometry analysis and colony-forming cell assays.
7.3.2.3

Flow Cytometry

Fluorescently labeled antibodies for CD13, CD15, CD34, CD38, CD45, CD71
and GlyA cell surface markers were purchased from BD Biosciences. The cellsamples were incubated with various antibody combinations at 4oC for more than 30
min in 2% FBS Hanks' buffer. After antibody staining, the cells were washed twice
using Hanks’ buffer and fixed in 1% paraformaldehyde. Cells were analyzed bytriple-color flow cytometry on a FACSCalibur analyzer (BD Biosciences). Relevant

127Chapter 7

isotype controls were also included to confirm specificity and for compensation
setting. At least 20,000 gated events were acquired. The Milan-Mulhouse gating
method was used for cell enumeration [151].
7.3.2.4

Colony-Forming Cell Assay

Detailed protocols for CFC assay can be found in Chapter 5.3.2.3.
7.3.3

Statistical Analysis
All data were presented as mean ± standard deviation (SD). The statisticalsignificance of the data obtained was analyzed by the Student’s t-test. Probability
values of p < 0.05 were interpreted as denoting statistical significance.

7.4

Experimental Results

7.4.1

Lineage Analysis of Adherent and Non-Adherent HSPCs
After 10-day expansion culture using 600 initial CD34+ HSPCs, cell proliferationon HSPC non-adhesive TCPS surface was 835 ± 196 folds, while cell proliferation on
BuDA nanofiber scaffolds was comparatively similar at 702 ± 152 folds (p > 0.05).
For BuDA nanofiber scaffold, the non-adherent cell population made up 54 ± 10 % of
the total cell population, while the adherent cell population accounted for theremaining 46 ± 10 %.
Fig. 7.2 & 7.3 show the representative flow cytometry plots and surface marker
expression summary of the ex vivo expanded cells. Consistent with previous
observations in Chapters 5 & 6, expanded cells on BuDA condition expressedsignificantly higher percentages of CD34+CD45+ cells (33.1 ± 9.1 %) as compared to
TCPS condition (7.1 ± 3.4 %, p < 0.05, Fig. 7.2A & 7.3A). Cells from BuDA

128

Chapter 7

condition also expressed closer percentages of myeloid / erythroid (CD71 [61.2 ± 3.9%]), erythroid (GlyA [21.1 ± 4.3 %]) and myeloid (CD13 [64.7 ± 8.9 %], CD15 [40.2± 5.7 %]) markers, as compared with TCPS condition (CD71 [52.6 ± 3.3 %], GlyA
[10.7 ± 2.5 %], CD13 [90.2 ± 4.1 %], CD15 [51.8 ± 8.3 %]), suggesting balanced
differentiation of HSPCs into both the erythroid and myeloid lineages on aminatednanofiber scaffolds (Fig. 7.2B, 7.2C & 7.3A). As discussed previously, cells fromTCPS condition were more committed towards the myeloblast / monoblast lineage(Chapters 5 & 6). However, comparing between the adherent and non-adherent cell
populations of BuDA scaffold condition, distinct differences were observed in surface
marker distributions:(1) The adherent cell population expressed higher percentage of theCD34+CD45+ primitive phenotype (43.8 ± 11.1 %), as compared to the nonadherent cell population (21.9 ± 6.5 %, p < 0.05, Fig. 7.2A & 7.3A).
(2) The adherent cell population expressed higher percentage of CD13 myeloid
marker (68.3 ± 4.6 %), as compared to the non-adherent cell population(49.4 ± 5.7 %, p < 0.05, Fig. 7.2B & 7.3A).
(3) The non-adherent cell population expressed higher percentage of CD71
antigen (81.0 ± 3.1 %), as compared to adherent cell population(39.2 ± 8.2 %, p < 0.05, Fig. 7.2C & 7.3A). In particular, CD71high
expression was significant in the non-adherent population (46.2 ± 3.2 %, p <0.05), as compared to adherent population (14.1 ± 4.7 %, Fig. 7.2C). This
indicated that cells in the non-adherent cell population were skewed towards
erythroid lineage.

129

Chapter 7(4) The non-adherent cell population expressed higher percentage of late CD15
myeloid marker (44.4 ± 1.2 %, p < 0.05) and late GlyA erythroid marker(35.7 ± 7.4 % p < 0.05) as compared to adherent cell population (CD15[33.5 ± 1.2 %], GlyA [12.4 ± 2.5 %], Fig. 7.3A). This indicates that cells in
the non-adherent cell population were more differentiated.
(5) The CD34+CD45+ fraction of the non-adherent cell population expressedhigher co-expression of CD71 (57.0 ± 9.4 % of CD34+CD45+ cells, p < 0.05)
and CD71high (8.9 ± 2.3 % of CD34+CD45+ cells, p < 0.05) as compared tothe corresponding adherent cell population (26.9 ± 6.5 % and 3.6 ± 2.1 % of
CD34+CD45+ cells, respectively, Fig. 7.3B).

Figure 7.2: Representative FACS profiles of cells after 10-day ex vivo expansion on
TCPS, PES-BuDA, and non-adherent (‘sus’) and adherent (‘adh’) fractions from PESBuDA conditions. (A) CD45 vs. CD34. (B) CD13 vs. CD34. (C) CD45 vs. CD71.

130

Chapter 7

Figure 7.3: Surface marker expression summary of cells after 10-day ex vivo expansion
on TCPS, PES-BuDA, and non-adherent (‘sus’) and adherent (‘adh’) fractions fromPES-BuDA conditions. (A) Percentage of total cells expressing one or multiple CDmarkers. (B) Percentage of the CD34+CD45+ cell population that are also CDX+, where
CDX represents CD13, CD15, CD38, CD71 or GlyA. Data shown are mean ± SD of 3
experiments, each conducted in duplicates.

7.4.2

Clonogenic Differences of Adherent and Non-Adherent HSPCs
CFC assays were conducted to evaluate the fraction of primitive progenitor cells

in the expanded cultures (Fig. 7.4). Cells expanded from BuDA-conjugated nanofiber
scaffolds generated significantly higher percentages of the more primitive CFU131

Chapter 7

GEMM units (23.8 ± 2.2 %, p < 0.05), as compared to cells expanded from TCPS
condition (13.0 ± 3.6 %). TCPS condition, on the other hand, generated higher
percentages of CFU-GM units (66.5 ± 9.1 %, p < 0.05), compared with BuDA
condition (52.3 ± 6.8 %), indicating differentiation commitment of the TCPSexpanded cells towards the myeloblast / monoblast lineage.

Figure 7.4: Specific CFU fractions after 14 days of culture, using the cells from 10-day
expansion cultures on TCPS, PES-BuDA, and non-adherent (‘sus’) and adherent (‘adh’)fractions from PES-BuDA conditions, normalized to CFU type per total CFU generated.
Data are means ± SD of 2 experiments, each conducted in triplicates. * indicates p <0.05.

Comparing between the adherent and non-adherent cell populations of BuDA
scaffold condition, we see that the adherent cell population yielded significantly
higher percentages of CFU-GM units (57.4 ± 4.4 %, p < 0.05) and lower percentagesof BFU-E units (19.6 ± 9.2 %, p < 0.05), as compared to the non-adherent cell
population (CFU-GM [39.8 ± 3.2 %] and BFU-E [35.3 ± 4.6 %]). This result indicatesdifferentiation commitment of the adherent population towards the myeloblast /132

Chapter 7

monoblast lineage, while for the non-adherent population differentiation commitment
towards the erythroid lineage is significant.
7.5

Discussion
The phenomenon that aminated nanofibers can actively interact and modulate

HSPC proliferation and phenotype maintenance underscores the importance of culture
substrate properties in current stromal-free, cytokine-supplemented ex vivo HSPC
expansion systems. A consequence of these interactions is the identification of aproliferative HSPC population subset that is highly adhesive to the aminatednanofiber surface. This study represents the first to identify such an adhesive
population among HSPCs. Using BuDA-conjugated nanofiber scaffolds, we sought toinvestigate the phenotypic properties of this adherent hematopoietic cell population,
as compared to the non-adherent hematopoietic cell population in the ex vivo HSPC
expansion cultures.
The flow cytometry surface marker analysis results show that cells expressing
CD34+ were significantly enriched in the adherent cell population, compared with the
non-adherent cell population, on aminated nanofiber scaffold cultures. The adherent
cell population also exhibits a lesser degree of late differentiation markers (CD15 andGlyA), compared with the non-adherent cell population. This evidence suggests theimportant roles that cell-scaffold interactions play in maintaining HSPC proliferative
capacity, as well as primitive phenotype maintenance, in cytokine-supplemented
expansion cultures.
We have shown here and in Chapters 5 & 6 that in pure suspension cultures lackof any cell-substrate interaction signals (e.g. TCPS condition), the CD34 primitive
phenotype maintenance of the cultured HSPCs is inefficient. In addition, we have also

133

Chapter 7

shown that for scaffold cultures without the correct substrate signals (e.g. nonaminated scaffolds), the HSPC proliferative capacity in these scaffold cultures is poor.
These observations suggest that the surface-immobilized amine molecules may be
acting as a kind of “HSPC ligand” that mediates HSPC-scaffold interaction, and thenanofiber textured topography further enhances this cell-scaffold interaction [57-59].
As proposed earlier in Chapters 5 & 6, the results presented here further suggestthat the CD34 surface marker likely serves as a receptor for the “amine-ligand”. In thenormal state, the highly sialylated and negatively-charged hematopoietic CD34
antigen acts as anti-adhesive molecule, preventing cellular adhesion in hematopoietic
cells strongly expressing this marker (i.e. HSPCs) [147-150,152-154,156].
Furthermore, our observations have shown that cell-scaffold adhesion correlates wellwith CD34+ expression, substantiating the hypothesis that the negatively-charged
CD34 antigen could interact with the positively-charged immobilized amino groups.
An interesting finding that arises from comparing the surface marker phenotypes(Fig. 7.2 & 7.3) and clonogenic differences (Fig. 7.4) of the adherent and nonadherent hematopoietic cell populations is that the predominant lineage commitmentsof HSPCs within the 2 cell populations are different: The adherent HSPC population
is prevalently committed towards the myeloblast / monoblast lineage, while the nonadherent HSPC population is significantly committed towards the erythroid lineage.
Currently, HSPC lineage commitment and differentiation is frequently
modulated using soluble signals from recombinant cytokines in stromal-free ex vivo
hematopoietic expansion cultures. Recombinant cytokines like IL, TPO, SCF, FL, etc.
are used alone or in combination to regulate hematopoiesis in these suspension
cultures [173-177,202,203]. For example, TPO, in combination with IL-1, IL-6, IL-11

134

Chapter 7

and SCF has been shown to promote proliferation and differentiation of
megakaryocytes from CD34+ cells in vitro [202]; while FL, in combination with IL-7
and IL-12 has been shown to promote expansion of T- and B-lymphocyte progenitors[203]. FL and SCF are also important in regulating early hematopoiesis and it isfrequently used in combination with GM-CSF, TPO or other cytokines to stimulate
the proliferation of early hematopoietic progenitor cells [173-177].
However, the role of cell-substrate interaction signals has rarely been
investigated in HSPC expansion cultures, maybe because of the lack of a suitable
HSPC-adhesive substrate to investigate its effects. In this study, through the use of
HSPC-adhesive aminated nanofiber scaffolds, we demonstrated the cell-scaffold
interaction, as a new and convenient approach to regulate HSPC (CD34+)
multipotency maintenance as well as lineage commitment. Although the specificcontrol of hematopoiesis is currently limited in our present scaffold system, we
envision that by combining with potential factors like recombinant cytokines andother cell adhesion molecules [173-177,202-205], modulating HSPC-substrateinteraction through nanofiber scaffold will become a highly useful method to regulate
HSPC expansion.
7.6

Concluding Remarks
Aminated nanofibers enhance the interaction with HSPCs in ex vivo expansion

culture; and the adherent hematopoietic cell population is enriched with CD34+CD45+
cells. In addition, different lineage commitment patterns are observed between theadherent and non-adherent cell populations. This study highlights the importance ofcell-scaffold interactions in modulating HSPC maintenance and lineage commitment.

135Chapter 8

CHAPTER EIGHT
Conclusions
The success of scaffold-based cell cultures largely depends on the optimal eventsof attachment, proliferation, differentiation, and phenotypic maintenance, which inturn are governed by a host of signals provided by the cell-scaffold microenvironment.
These signals include: (1) homotypic / heterotypic cell-cell interaction; (2) soluble
signaling molecules; and (3) cell-substrate interaction signals. An ideal scaffold
culture system should include all these interactive components. In this thesis, we havefocused on cell-substrate interaction signals mediated by both substrate-boundsignaling molecules and nanofiber scaffold topographical cues.
The general strategy of developing a scaffold that can provide both topographicaland immobilized biochemical cues involves first nanofiber scaffold fabrication via theelectrospinning technique, followed by nanofiber bio-functionalization. The biofunctionalization process involves the initial functionalization of the nanofiber surface
with carboxylic acid groups using UV-initiated poly(acrylic acid) grafting method.
This is followed by conjugation of bioactive molecules onto the functionalized
nanofiber surfaces. In this thesis, we have presented the efficacy of this nanofiber biofunctionalization strategy on various cell culture systems like hepatocyte scaffold
cultures and hematopoietic stem cell expansion cultures.
In Chapter 3, we have described the galactose bio-functionalization ofelectrospun PCLEEP nanofibers for liver cell cultures. Both galactosylated nanofiber
scaffolds and galactosylated films supported the hepatic functions (albumin secretion,ammonia removal and cytochrome P450 activity) of the cultured primary hepatocytes
136

Chapter 8

better than unmodified nanofiber scaffolds and films. Interestingly, galactosylated
nanofiber scaffolds exhibit the unique property of promoting hepatocyte aggregatesand cell infiltration within the mesh and around the fibers, forming an integrated
spheroid-nanofiber construct. These results demonstrated that the nanofiber
topography together with surface-immobilized galactose ligand synergisticallyenhance cell-substrate interaction as indicated by hepatocyte adhesion and infiltration,even though this enhanced cell-substrate interaction did not translate into significantly
higher functional enhancement as compared to galactosylated film condition.
We subsequently demonstrated in Chapter 4 that hepatocyte cytochrome P450
activity enhancement can be brought about through further 3-Mc bio-functionalizationof the galactosylated nanofiber scaffold. By taking advantage of the porous and layerforming properties of electrospun nanofibers, we had designed a dual functional
scaffold that induces two different biological responses from hepatocytes. Galactose
bio-functionalization on nanofibers resulted in a scaffold that can induce hepatocyte
adhesion and re-organization, while 3-Mc loading into the fiber, working together
with fiber galactosylation, resulted in a hepatocyte bioactive scaffold that can also
regulate the hepatocyte cytochrome P450 function.
In Chapter 5, we further presented the efficacy of this nanofiber biofunctionalization strategy on hematopoietic stem / progenitor cell expansion. Using
amine molecule bio-functionalization, we have demonstrated the effectiveness ofaminated electrospun PES nanofiber mesh as potential scaffolds for ex vivo HSPC
expansion under serum-free conditions. Among the carboxylated, hydroxylated, andaminated PES substrates and TCPS, aminated PES substrates mediated the highest
expansion efficiency of CD34+CD45+ cells and CFU potential. Aminated nanofiber
mesh could further enhance the HSPC-substrate adhesion. In particular, aminated137Chapter 8

nanofiber meshes supported a higher degree of cell adhesion, percentage of
CD34+CD45+ cells and expansion of CFU-GEMM forming progenitor cells, ascompared to aminated films. SEM imaging also revealed the discrete colonies of cells
proliferating and interacting with the aminated nanofibers.
In Chapter 6, we showed that nanofiber scaffolds immobilized with aminefunctional groups of increasing 2-, 4-, 6-carbon spacer chain lengths could further
modulate HSPC proliferation and phenotype maintenance, resulting in different HSPC
proliferation kinetics, cell population phenotypic (CD34 and CD13 markers)
expression, mouse engraftment potential and also short-term and long-term colonyforming ability. These observations further suggested the importance of nanofiber
topography and amino functional group mediated cell-scaffold interactions inregulating HSPC proliferation and self-renewal.
Finally, in Chapter 7, we compared the surface marker phenotypic andclonogenic differences of the adherent and non-adherent hematopoietic cell
populations that arise after ex vivo expansion culture of CD34+ HSPCs on aminatedelectrospun nanofiber scaffolds. Detailed flow cytometry analysis showed that theadherent cell population expressed significantly higher percentage of CD34+CD45+
cells, compared with the non-adherent cell population. In addition, flow cytometry
analysis and CFU assay also indicated significant commitment of the adherent
population towards the myeloblast / monoblast lineage, while the non-adherent
population showed skewed commitment towards the erythroid lineage. This study
also highlights the importance of cell-scaffold interactions as a new approach inmodulating HSPC multipotency maintenance and lineage commitment, other thancytokine modulation frequently described in literature.

138

Chapter 8

In summary, this research has:
(1) Presented a nanofiber bio-functionalization strategy to develop polymericnanofiber constructs that can serve as cell culture scaffolds.
(2) Demonstrated through primary hepatocyte cultures and hematopoietic stem /progenitor cell expansion cultures that these scaffolds can promote cellsubstrate interactions and are bioactive in regulating cellular responses likecell adhesion, cell morphological reorganization, cell differentiated functions,cell proliferation, and cell phenotype maintenance.
(3) Demonstrated the synergistic effects that nanofiber topography and surface
immobilized biochemical cues play in enhancing these cell-scaffold
interactions and regulation of cellular functions.

139

Appendix

APPENDIX[1]

Synthesis of 1-O-(6’-Aminohexyl)-D-galactopyranoside (AHG)
The galactose ligand AHG was synthesized according to procedures reported by

Yin et al. [33]. The scheme is shown in Fig. 8.1.

Figure 8.1: Synthesis scheme for AHG.

Benzyl N-(6-hydroxyhexyl) carbamate
Carbobenzoxy chloride solution (50% in toluene, 40 mL) and K2CO3 solution(8.3 g in 30 mL of H2O) were added dropwise to an ice-cooled solution of 6-amino-1hexanol (11.7 g, 0.1 mol) in 400 mL of ethyl acetate from two addition funnels
simultaneously. After the addition, the mixture was further stirred at room
temperature for 3 h, followed by washing with 1 N HCl (3×200 mL) and water(3×200 mL). The solution was dried over anhydrous MgSO4 and evaporated todryness. The residue was recrystallized from ethylacetate to yield a white powder(13.4 g, 53.4%), m.p. 80-82°C.
140Appendix

1,2,3,4,6-Penta-O-acetyl-D-galactopyranose (2)β-D-galactose (1) (18.0 g, 0.10 mol) was dissolved in a mixture of dry pyridine(150 mL, 1.86 mol) and acetic anhydride (150 mL, 1.60 mol) and stirred at room
temperature for 3 days. The mixture was concentrated by vacuum rotary evaporation
to yield a yellow syrupy residue. The residue was dissolved in 200 mL of CHCl3,
extracted with 200 mL of cold 2 N H2SO4, and washed with saturated NaHCO3
solution (200 mL) and water (2×200 mL). The organic phase was dried overanhydrous MgSO4. The solution was filtered, concentrated, and vacuum dried. The
residue was recrystallized from ethanol to yield a white powder (22.5 g, 57.7%).
Thin-layer chromatography (TLC): ethylacetate-hexane (3:2), Rf = 0.54. 1H-NMR(CDCl3) δ: 1.99 (s, 3H, Me), 2.01 (s, 3H, Me), 2.03 (s, 3H, Me), 2.15 (d, 6H, 2Me),
4.10 (m, 2H, 2H6), 4.34 (m, 1H, H4), 5.33 (m, 2H, H5 and H3), 5.49 (t, 1H, H2), 6.37 (s,1H, H1).
2,3,4,6-Tetra-O-acetyl-1-bromo-1-deoxy-D-galactopyranose (3)Ten grams of (2) was dissolved in 50 mL of HBr solution (in glacial acetic acid,
11.5%, w/v) and diluted with 200 mL of CHCl3. The resulted mixture was poured into
1.8 L of ice-water and thoroughly mixed. The organic layer was collected, washedwith saturated NaHCO3 solution (2×100 mL) and water (2×100 mL), dried overMgSO4, and filtered. The filtrate was vacuum dried to syrup (3). Yield: 9.8 g (93.0%).
TLC: ethylacetate-hexane (3:2), Rf = 0.65. 1H-NMR (CDCl3) δ: 1.89 (s, 3H, Me), 1.95
(s, 6H, 2Me), 2.05 (s, 3H, Me), 4.07 (m, 2H, 2H6), 4.37 (m, 1H, H4), 5.01 (m, 2H, H5
and H3), 5.11 (t, 1H, H2), 5.30 (s, 1H, H1).

141

Appendix

1-O-[6’-(N-Benzyloxycarbonyl)aminohexyl]-2,3,4,6-tetra-O-acetyl-Dgalactopyranoside (4)(3) (9.6 g, 23.5 mmol) was mixed with benzyl N-(6-hydroxyhexyl) carbamate(6.52 g, 26 mmol), Hg(CN)2 (6.55g, 26 mmol), Drierite (2.6 g) in a toluenenitromethane mixture (1:1, v/v, 250 mL) and stirred for 24 h. The mixture was filteredand the filtrate was concentrated under reduced pressure. The residue was dissolved in
CHCl3 (200 mL), washed with 1M NaCl solution (2×200 mL) and 0.5M KBr solution(200 mL), dried over MgSO4, and filtered. The filtrate was concentrated to syrup. The
crude product was subjected to silica chromatography using ethylacetate-hexane (3:2,v/v, Rf = 0.39) as the eluent. (4) was obtained as white powder after evaporation of
the solvent from the corresponding fractions (5.2 g, 38.2%). 1H-NMR (CDCl3) δ:
1.21-1.52 (m, 8H, 4CH2), 1.95-2.18 (m, 12H, 4Me), 2.96 (t, 2H, CH2-N), 3.54 (m, 2H,
O-CH2), 4.03 (m, 2H, 2H6), 4.35 (m, 1H, H4), 4.65 (m, 1H, H5), 4.72 (d, 1H, H2), 4.85
(d, 1H, H2), 5.03 (d, 1H, H1).
1-O-[6’-(N-Benzyloxycarbonyl)aminohexyl]-D-galactopyranoside (5)One milliliter of sodium methoxide solution in methanol (5%, w/v) was added toa solution of (4) (5.0 g, 8.6 mmol) in methanol (100 mL). The mixture was stirred for
3 h, followed by adding Dowex 50WX8-200 ion-exchange resin (pretreated with 1N
HCl and washed with methanol) until the pH value of the solution reached 5-6. The
mixture was gently stirred for 0.5 h and filtered. The filtrate was evaporated to yield
yellowish syrup (3.2 g, 90.0%). TLC: ethylacetate-acetic acid (9:1), Rf = 0.78. 1HNMR (D2O) δ: 1.19 (m, 4H, 2CH2), 1.34 (m, 2H, CH2), 1.49 (m, 2H, CH2), 2.98 (t,2H, CH2-N), 3.44 (m, 2H, O-CH2), 3.55 (m, 2H, H3 and H4), 3.70 (d, 2H, H6), 3.79 (m,1H, H5), 3.85 (d, 1H, H2), 4.24 (d, 1H, H1), 4.93 (s, 2H, CH2-Ph), 7.20 (m, 5H, C6H5).

142

Appendix

1-O-(6’-Aminohexyl)-D-galactopyranoside (6)The deacetylated product (5) (3.2 g, 7.7 mmol) was dissolved in methanol (150mL) with Pd-C catalyst (1.6 g). Hydrogen gas was bubbled into the stirred mixture
until benzyloxycarbonyl group was completely removed as determined by TLC. Pd-C
was filtered off and the filtrate was concentrated and vacuum dried to syrup. It wasthen dissolved in distilled water and lyophilized to obtain white powder (6) (1.8 g,
83.3%). TLC: ethanol-acetic acid (9:1), Rf = 0.23. 1H-NMR (D2O) δ: 1.32 (m, 4H,
2CH2), 1.43 (m, 2H, CH2), 1.59 (m, 2H, CH2), 2.61 (t, 2H, N-CH2), 3.43 (m, 2H, OCH2), 3.62 (m, 2H, H3 and H4), 3.73 (m, 2H, H6), 3.88 (m, 2H, H5 and H2), 4.33 (d,1H, H1).

143

References

REFERENCES[1] Hubbell JA. Biomaterials in tissue engineering. Biotechnology (NY) 1995;13(6):565-576.
[2] Griffith LG. Emerging design principles in biomaterials and scaffolds for tissue
engineering. Ann N Y Acad Sci 2002;961:83-95.
[3] Peter SJ et al. Polymer concepts in tissue engineering. J Biomed Mater Res
1998;43(4):422-427.
[4] Chaikof EL et al. Biomaterials and scaffolds in reparative medicine. Ann N Y Acad Sci
2002;961:96-105.
[5] Yarlagadda PK et al. Recent advances and current developments in tissue scaffolding.
Biomed Mater Eng 2005;15(3):159-177.
[6] Hammond JS et al. Scaffolds for liver tissue engineering. Expert Rev Med Devices
2006;3(1):21-27.
[7] Bottaro DP et al. Molecular signaling in bioengineered tissue microenvironments. Ann N
Y Acad Sci 2002;961:143-153.
[8] Lutolf MP et al. Synthetic biomaterials as instructive extracellular microenvironments formorphogenesis in tissue engineering. Nat Biotechnol 2005;23(1):47-55.
[9] Dersch R et al. Nanoprocessing of polymers: applications in medicine, sensors, catalysis,
photonics. Polymers for Advanced Technologies 2005;16(2-3):276-282.
[10] Huang ZM et al. A review on polymer nanofibers by electrospinning and theirapplications in nanocomposites. Composites Science and Technology 2003;63(15):22232253.
[11] Jayaraman K et al. Recent advances in polymer nanofibers. Journal of Nanoscience andNanotechnology 2004;4(1-2):52-65.
[12] Subbiah T et al. Electrospinning of nanofibers. Journal of Applied Polymer Science
2005;96(2):557-569.
[13] Wang YK et al. Nanofibres and their influence on cells for tissue regeneration. Australian
Journal of Chemistry 2005;58(10):704-712.
[14] Li WJ et al. Electrospun nanofibrous structure: A novel scaffold for tissue engineering.
Journal of Biomedical Materials Research 2002;60(4):613-621.
[15] Li WJ et al. Biological response of chondrocytes cultured in three-dimensional
nanofibrouspoly(epsilon-caprolactone)
scaffolds.
J
BiomedMater
Res
2003;67A(4):1105-1114.
[16] Li WJ et al. Multilineage differentiation of human mesenchymal stem cells in a threedimensional nanofibrous scaffold. Biomaterials 2005;26(25):5158-5166.
[17] Li WJ et al. A three-dimensional nanofibrous scaffold for cartilage tissue engineering
using human mesenchymal stem cells. Biomaterials 2005;26(6):599-609.
[18] Badami AS et al. Effect of fiber diameter on spreading, proliferation, and differentiationof osteoblastic cells on electrospun poly(lactic acid) substrates. Biomaterials
2006;27(4):596-606.
[19] Shin M et al. Contractile cardiac grafts using a novel nanofibrous mesh. Biomaterials
2004;25(17):3717-3723.

144

References[20] Zong X et al. Electrospun fine-textured scaffolds for heart tissue construct. Biomaterials
2005;26(26):5330-5338.
[21] Mo XM et al. Electrospun P(LLA-CL) nanofiber: a biomimetic extracellular matrix forsmooth muscle cell and endothelial cell proliferation. Biomaterials 2004;25(10):18831890.
[22] Xu C et al. Electrospun nanofiber fabrication as synthetic extracellular matrix and its
potential for vascular tissue engineering. Tissue Eng 2004;10(7):1160-1168.
[23] Xu CY et al. In vitro study of human vascular endothelial cell function on materials withvarious surface roughness. Journal of Biomedical Materials Research Part A
2004;71A(1):154-161.
[24] Xu CY et al. Aligned biodegradable nanofibrous structure: a potential scaffold for bloodvessel engineering. Biomaterials 2004;25(5):877-886.
[25] Yang F et al. Characterization of neural stem cells on electrospun poly(L-lactic acid)
nanofibrous scaffold. J Biomater Sci Polym Ed 2004;15(12):1483-1497.
[26] Williamson MR et al. PCL-PU composite vascular scaffold production for vascular tissue
engineering: attachment, proliferation and bioactivity of human vascular endothelial cells.
Biomaterials 2006;27(19):3608-3616.
[27] Sun T et al. Self-organization of skin cells in three-dimensional electrospun polystyrene
scaffolds. Tissue Eng 2005;11(7-8):1023-1033.
[28] Reneker DH et al. Nanometre diameter fibres of polymer, produced by electrospinning.
Nanotechnology 1996;7(3):216-223.
[29] Li D et al. Electrospinning of nanofibers: Reinventing the wheel? Advanced Materials
2004;16(14):1151-1170.
[30] Zhu Y et al. Surface modification of polycaprolactone membrane via aminolysis andbiomacromolecule immobilization for promoting cytocompatibility of human endothelial
cells. Biomacromolecules 2002;3(6):1312-1319.
[31] Oehr C et al. Plasma grafting - a method to obtain monofunctional surfaces. Surface &Coatings Technology 1999;119:25-35.
[32] Uchida E et al. A Novel Method for Graft-Polymerization Onto Poly(EthyleneTerephthalate) Film Surface by Uv Irradiation Without Degassing. Journal of Applied
Polymer Science 1990;41(3-4):677-687.
[33] Yin C et al. High density of immobilized galactose ligand enhances hepatocyte
attachment and function. J Biomed Mater Res 2003;67A(4):1093-1104.
[34] Ying L et al. Immobilization of galactose ligands on acrylic acid graft-copolymerizedpoly(ethylene terephthalate) film and its application to hepatocyte culture.
Biomacromolecules 2003;4(1):157-165.
[35] Tan WJ et al. Adhesion contact dynamics of primary hepatocytes on poly(ethylene
terephthalate) surface. Biomaterials 2005;26(8):891-898.
[36] Nakajima N et al. Mechanism of amide formation by carbodiimide for bioconjugation inaqueous media. Bioconjug Chem 1995;6(1):123-130.
[37] Reneker DH et al. Bending instability of electrically charged liquid jets of polymer
solutions in electrospinning. Journal of Applied Physics 2000;87(9):4531-4547.
[38] Yarin AL et al. Taylor cone and jetting from liquid droplets in electrospinning of
nanofibers. Journal of Applied Physics 2001;90(9):4836-4846.

145

References[39] Yarin AL et al. Bending instability in electrospinning of nanofibers. Journal of Applied
Physics 2001;89(5):3018-3026.
[40] Deitzel JM et al. The effect of processing variables on the morphology of electrospun
nanofibers and textiles. Polymer 2001;42(1):261-272.
[41] Fong H et al. Beaded
1999;40(16):4585-4592.

nanofibers

formedduringelectrospinning.

Polymer[42] Zong XH et al. Structure and process relationship of electrospun bioabsorbable nanofiber
membranes. Polymer 2002;43(16):4403-4412.
[43] Luong-Van E et al. Controlled release of heparin from poly(epsilon-caprolactone)electrospun fibers. Biomaterials 2006;27(9):2042-2050.
[44] Huang CB et al. Electrospun polymer nanofibres with small diameters. Nanotechnology
2006;17(6):1558-1563.
[45] Li D et al. Electrospinning of polymeric and ceramic nanofibers as uniaxially aligned
arrays. Nano Letters 2003;3(8):1167-1171.
[46] Dersch R et al. Electrospun nanofibers: Internal structure and intrinsic orientation. Journalof Polymer Science Part A-Polymer Chemistry 2003;41(4):545-553.
[47] Teo WE et al. Electrospun fibre bundle made of aligned nanofibres over two fixed points.
Nanotechnology 2005;16(9):1878-1884.
[48] Theron A et al. Electrostatic field-assisted alignment of electrospun nanofibres.
Nanotechnology 2001;12(3):384-390.
[49] Katta P et al. Continuous electrospinning of aligned polymer nanofibers onto a wire drumcollector. Nano Letters 2004;4(11):2215-2218.
[50] Sun ZC et al. Compound core-shell polymer nanofibers by co-electrospinning. Advanced
Materials 2003;15(22):1929-+.
[51] Li D et al. Nanofibers of conjugated polymers prepared by electrospinning with a twocapillary spinneret. Advanced Materials 2004;16(22):2062-+.
[52] Zhang YZ et al. Coaxial electrospinning of (fluorescein isothiocyanate-conjugated bovine
serum albumin)-encapsulated poly(epsilon-caprolactone) nanofibers for sustained release.
Biomacromolecules 2006;7(4):1049-1057.
[53] Kidoaki S et al. Mesoscopic spatial designs of nano- and microfiber meshes for tissueengineering matrix and scaffold based on newly devised multilayering and mixing
electrospinning techniques. Biomaterials 2005;26(1):37-46.
[54] Ding B et al. Fabrication of blend biodegradable nanofibrous nonwoven mats via multijet electrospinning. Polymer 2004;45(6):1895-1902.
[55] Curtis AS et al. Reactions of cells to topography. J Biomater Sci Polym Ed
1998;9(12):1313-1329.
[56] Flemming RG et al. Effects of synthetic micro- and nano-structured surfaces on cell
behavior. Biomaterials 1999;20(6):573-588.
[57] Dalby MJ et al. In vitro reaction of endothelial cells to polymer demixed nanotopography.
Biomaterials 2002;23(14):2945-2954.
[58] Barbucci R et al. Micro and nano-structured surfaces. J Mater Sci Mater Med
2003;14(8):721-725.
[59] Dalby MJ et al. Use of nanotopography to study mechanotransduction in fibroblasts-methods and perspectives. Eur J Cell Biol 2004;83(4):159-169.

146

References[60] Nishimura S et al. Three-dimensional architecture and distribution of collagencomponents in the goat hypophysis. Anat Rec A Discov Mol Cell Evol Biol
2004;277(2):275-286.
[61] Ojeda JL et al. Evidence of a new transitory extracellular structure within the developing
rhombencephalic cavity. An ultrastructural and immunoelectron-microscopic study in thechick. Anat Embryol (Berl) 2000;202(3):257-264.
[62] Nishida T et al. The network structure of corneal fibroblasts in the rat as revealed byscanning electron microscopy. Invest Ophthalmol Vis Sci 1988;29(12):1887-1890.
[63] He W et al. Fabrication and endothelialization of collagen-blended biodegradable
polymer nanofibers: potential vascular graft for blood vessel tissue engineering. Tissue
Eng 2005;11(9-10):1574-1588.
[64] Li M et al. Electrospun protein fibers as matrices for tissue engineering. Biomaterials
2005;26(30):5999-6008.
[65] Ji Y et al. Electrospun three-dimensional hyaluronic acid nanofibrous scaffolds.
Biomaterials 2006;27(20):3782-3792.
[66] Rho KS et al. Electrospinning of collagen nanofibers: effects on the behavior of normal
human keratinocytes and early-stage wound healing. Biomaterials 2006;27(8):1452-1461.
[67] Chew SY et al. Sustained release of proteins from electrospun biodegradable fibers.
Biomacromolecules 2005;6(4):2017-2024.
[68] Li C et al. Electrospun silk-BMP-2 scaffolds for bone tissue engineering. Biomaterials
2006;27(16):3115-3124.
[69] Liang DH et al. In vitro non-viral gene delivery with nanofibrous scaffolds. Nucleic Acids
Research 2005;33(19):170.
[70] Ma ZW et al. Surface engineering of electrospun polyethylene terephthalate (PET)
nanofibers towards development of a new material for blood vessel engineering.
Biomaterials 2005;26(15):2527-2536.
[71] Ma ZW et al. Grafting of gelatin on electrospun poly(caprolactone) nanofibers to improveendothelial cell spreading and proliferation and to control cell orientation. Tissue Eng
2005;11(7-8):1149-1158.
[72] Robinette EJ et al. Synthesis of polymer-polymer nanocomposites using radiation grafting
techniques. Nuclear Instruments & Methods in Physics Research Section B-BeamInteractions with Materials and Atoms 2005;236:216-222.
[73] Kim TG et al. Biomimicking extracellular matrix: cell adhesive RGD peptide modified
electrospun poly(D,L-lactic-co-glycolic acid) nanofiber mesh. Tissue Eng
2006;12(2):221-233.
[74] Bismuth H et al. Orthotopic liver transplantation in fulminant and subfulminant hepatitis.
The Paul Brousse experience. Ann Surg 1995;222(2):109-119.
[75] McCashland TM et al. The American experience with transplantation for acute liver
failure. Semin Liver Dis 1996;16(4):427-433.
[76] Gridelli B et al. Strategies for making more organs available for transplantation. N Engl J
Med 2000;343(6):404-410.
[77] Kaihara S et al. Tissue engineering: toward new solutions for transplantation andreconstructive surgery. Arch Surg 1999;134(11):1184-1188.
[78] Tzanakakis ES et al. Extracorporeal tissue engineered liver-assist devices. Annu Rev
Biomed Eng 2000;2:607-632.

147

References[79] Allen JW et al. Advances in bioartificial liver devices. Hepatology 2001;34(3):447-455.
[80] Allen JW et al. Engineering liver therapies for the future. Tissue Eng 2002;8(5):725-737.
[81] Chan C et al. Hepatic tissue engineering for adjunct and temporary liver support: criticaltechnologies. Liver Transpl 2004;10(11):1331-1342.
[82] Dich J et al. Long-term culture of hepatocytes: effect of hormones on enzyme activities
and metabolic capacity. Hepatology 1988;8(1):39-45.
[83] Waxman DJ. P450 gene induction by structurally diverse xenochemicals: central role ofnuclear receptors CAR, PXR, and PPAR. Arch Biochem Biophys 1999;369(1):11-23.
[84] Liu J et al. Characterization and evaluation of detoxification functions of anontumorigenic immortalized porcine hepatocyte cell line (HepLiu). Cell Transplant
1999;8(3):219-232.
[85] Kobayashi N et al. Establishment of a reversibly immortalized human hepatocyte cell line
by using Cre/loxP site-specific recombination. Transplant Proc 2000;32(5):1121-1122.
[86] Roy P et al. Analysis of oxygen transport to hepatocytes in a flat-plate microchannel
bioreactor. Ann Biomed Eng 2001;29(11):947-955.
[87] Bissell DM et al. Interactions of rat hepatocytes with type IV collagen, fibronectin andlaminin matrices. Distinct matrix-controlled modes of attachment and spreading. Eur J
Cell Biol 1986;40(1):72-78.
[88] Ben Ze'ev A et al. Cell-cell and cell-matrix interactions differentially regulate theexpression of hepatic and cytoskeletal genes in primary cultures of rat hepatocytes. Proc
Natl Acad Sci U S A 1988;85(7):2161-2165.
[89] Dunn JC et al. Long-term in vitro function of adult hepatocytes in a collagen sandwich
configuration. Biotechnol Prog 1991;7(3):237-245.
[90] Hamilton GA et al. Regulation of cell morphology and cytochrome P450 expression inhuman hepatocytes by extracellular matrix and cell-cell interactions. Cell Tissue Res
2001;306(1):85-99.
[91] Vinken M et al. Rat hepatocyte cultures: collagen gel sandwich and immobilization
cultures. Methods Mol Biol 2006;320:247-254.
[92] Carlisle ES et al. Enhancing hepatocyte adhesion by pulsed plasma deposition andpolyethylene glycol coupling. Tissue Eng 2000;6(1):45-52.
[93] Chia SM et al. Hepatocyte encapsulation for enhanced cellular functions. Tissue Eng
2000;6(5):481-495.
[94] Yin C et al. Microcapsules with improved mechanical stability for hepatocyte culture.
Biomaterials 2003;24(10):1771-1780.
[95] Bhatia SN et al. Effect of cell-cell interactions in preservation of cellular phenotype:
cocultivation of hepatocytes and nonparenchymal cells. FASEB J 1999;13(14):1883-1900.
[96] Bhandari RN et al. Liver tissue engineering: a role for co-culture systems in modifying
hepatocyte function and viability. Tissue Eng 2001;7(3):345-357.
[97] Lu HF et al. Three-dimensional co-culture of rat hepatocyte spheroids and NIH/3T3
fibroblasts enhances hepatocyte functional maintenance. Acta Biomater 2005;1(4):399410.
[98] Peshwa MV et al. Mechanistics of formation and ultrastructural evaluation of hepatocyte
spheroids. In Vitro Cell Dev Biol Anim 1996;32(4):197-203.

148

References[99] Tzanakakis ES et al. The role of actin filaments and microtubules in hepatocyte spheroid
self-assembly. Cell Motil Cytoskeleton 2001;48(3):175-189.
[100] Abu-Absi SF et al. Structural polarity and functional bile canaliculi in rat hepatocyte
spheroids. Exp Cell Res 2002;274(1):56-67.
[101] Ma M et al. Biochemical and functional changes of rat liver spheroids during spheroid
formation and maintenance in culture: I. morphological maturation and kinetic changes ofenergy metabolism, albumin synthesis, and activities of some enzymes. J Cell Biochem
2003;90(6):1166-1175.
[102] Landry J et al. Spheroidal aggregate culture of rat liver cells: histotypic reorganization,
biomatrix deposition, and maintenance of functional activities. J Cell Biol
1985;101(3):914-923.
[103] Hodgkinson CP et al. Fibronectin-mediated hepatocyte shape change reprograms
cytochrome P450 2C11 gene expression via an integrin-signaled induction ofribonuclease activity. Mol Pharmacol 2000;58(5):976-981.
[104] Lin KH et al. Long-term maintenance of liver-specific functions in three-dimensional
culture of adult rat hepatocytes with a porous gelatin sponge support. Biotechnol Appl
Biochem 1995;21 ( Pt 1):19-27.
[105] Ijima H et al. Formation of a spherical multicellular aggregate (spheroid) of animal cells
in the pores of polyurethane foam as a cell culture substratum and its application to a
hybrid artificial liver. J Biomater Sci Polym Ed 1998;9(7):765-778.
[106] Powers MJ et al. A microfabricated array bioreactor for perfused 3D liver culture.
Biotechnol Bioeng 2002;78(3):257-269.
[107] Mizumoto H et al. Liver regeneration using a hybrid artificial liver support system. Artif
Organs 2004;28(1):53-57.
[108] Koide N et al. Formation of multicellular spheroids composed of adult rat hepatocytes indishes with positively charged surfaces and under other nonadherent environments. Exp
Cell Res 1990;186(2):227-235.
[109] Koide N et al. Continued high albumin production by multicellular spheroids of adult rat
hepatocytes formed in the presence of liver-derived proteoglycans. Biochem Biophys Res
Commun 1989;161(1):385-391.
[110] Yagi K et al. Rapid formation of multicellular spheroids of adult rat hepatocytes byrotation culture and their immobilization within calcium alginate. Artif Organs
1993;17(11):929-934.
[111] Sakai Y et al. Large-scale preparation and function of porcine hepatocyte spheroids. Int J
Artif Organs 1996;19(5):294-301.
[112] Cho CS et al. Galactose-carrying polymers as extracellular matrices for liver tissue
engineering. Biomaterials 2006;27(4):576-585.
[113] Cho CS et al. Effect of ligand orientation on hepatocyte attachment onto the poly(N-pvinylbenzyl-o-beta-D-galactopyranosyl-D-gluconamide) as a model ligand ofasialoglycoprotein. J Biomater Sci Polym Ed 1996;7(12):1097-1104.
[114] Gutsche AT et al. Engineering of a sugar-derivatized porous network for hepatocyte
culture. Biomaterials 1996;17(3):387-393.
[115] Weigel PH et al. Specific adhesion of rat hepatocytes to beta-galactosides linked topolyacrylamide gels. J Biol Chem 1978;253(2):330-333.
[116] Griffith LG et al. Microdistribution of substratum-bound ligands affects cell function:
hepatocyte spreading on PEO-tethered galactose. Biomaterials 1998;19(11-12):979-986.

149

References[117] Chung TW et al. Preparation of alginate/galactosylated chitosan scaffold for hepatocyte
attachment. Biomaterials 2002;23(14):2827-2834.
[118] Park TG. Perfusion culture of hepatocytes within galactose-derivatized biodegradablepoly(lactide-co-glycolide) scaffolds prepared by gas foaming of effervescent salts. J
Biomed Mater Res 2002;59(1):127-135.
[119] Yang J et al. Galactosylated alginate as a scaffold for hepatocytes entrapment.
Biomaterials 2002;23(2):471-479.
[120] Yoon JJ et al. Surface immobilization of galactose onto aliphatic biodegradable polymers
for hepatocyte culture. Biotechnol Bioeng 2002;78(1):1-10.
[121] Wen J et al. Preparation and characterization of poly(D,L-lactide-co-ethylene methyl
phosphate). Polym Int 1998;47(4):503-509.
[122] Mao HQ. Biodegradable polyphosphoesters. In: Mathiowitz E, editor. Encyclopedia ofControlled Drug Delivery. New York, NY: Johns Wiley & Sons, Inc., 1999.
[123] Wen J et al. Poly(D,L-lactide-co-ethyl ethylene phosphate)s as new drug carriers. J
Control Release 2003;92(1-2):39-48.
[124] Friend JR et al. Formation and characterization of hepatocyte spheroids. In: Morgan JR,
Yarmush ML, editors. Tissue engineering methods and protocols. Totowa, NJ: Humana
Press Inc., 1999: 245-252.
[125] Jauregui HO et al. Xenobiotic induction of P-450 PB-4 (IIB1) and P-450c (IA1) andassociated monooxygenase activities in primary cultures of adult rat hepatocytes.
Xenobiotica 1991;21(9):1091-1106.
[126] Hansen T et al. Cytochrome P450 enzyme activity and protein expression in primary
porcine enterocyte and hepatocyte cultures. Xenobiotica 2000;30(1):27-46.
[127] Behnia K et al. Xenobiotic metabolism by cultured primary porcine hepatocytes. Tissue
Eng 2000;6(5):467-479.
[128] Xu J et al. Characterisation of some cytotoxic endpoints using rat liver and HepG2
spheroids as in vitro models and their application in hepatotoxicity studies. II. Spheroid
cell spreading inhibition as a new cytotoxic marker. Toxicol Appl Pharmacol
2003;189(2):112-119.
[129] Shimada T et al. Dose-response studies on the induction of liver cytochromes P4501A1
and 1B1 by polycyclic aromatic hydrocarbons in arylhydrocarbon-responsive C57BL/6J
mice. Xenobiotica 2003;33(9):957-971.
[130] Chen XQ et al. Prediction of aqueous solubility of organic compounds using aquantitative structure-property relationship. J Pharm Sci 2002;91(8):1838-1852.
[131] Rytting E et al. Aqueous and cosolvent solubility data for drug-like organic compounds.
AAPS J 2005;7(1):E78-E105.
[132] Emerson SG et al. In vitro expansion of hematopoietic cells for clinical application.
Cancer Treat Res 1995;76:215-223.
[133] Alcorn MJ et al. Ex vivo expansion of haemopoietic progenitor cells. Blood Rev
1996;10(3):167-176.
[134] Collins PC et al. Ex vivo culture systems for hematopoietic cells. Curr Opin Biotechnol
1996;7(2):223-230.
[135] McAdams TA et al. Hematopoietic cell culture therapies (Part II): Clinical aspects andapplications. Trends Biotechnol 1996;14(10):388-396.

150

References[136] Bremers AJ et al. Immunology and immunotherapy of human cancer: present concepts
and clinical developments. Crit Rev Oncol Hematol 2000;34(1):1-25.
[137] Noll T et al. Cultivation of hematopoietic stem and progenitor cells: biochemicalengineering aspects. Adv Biochem Eng Biotechnol 2002;74:111-128.
[138] Bonnet D. Biology of human bone marrow stem cells. Clin Exp Med 2003;3(3):140-149.
[139] Robinson S et al. Ex vivo expansion of umbilical cord blood. Cytotherapy 2005;7(3):243250.
[140] Takagi M. Cell processing engineering for ex-vivo expansion of hematopoietic cells. J
Biosci Bioeng 2005;99(3):189-196.
[141] Migliaccio AR et al. Cell dose and speed of engraftment in placental/umbilical cord bloodtransplantation: graft progenitor cell content is a better predictor than nucleated cell
quantity. Blood 2000;96(8):2717-2722.
[142] Eridani S et al. Cytokine effect on ex vivo expansion of haemopoietic stem cells fromdifferent human sources. Biotherapy 1998;11(4):291-296.
[143] Stewart DA et al. Factors predicting engraftment of autologous blood stem cells: CD34+
subsets inferior to the total CD34+ cell dose. Bone Marrow Transplant 1999;23(12):12371243.
[144] Wagner JE et al. Ex vivo expansion of umbilical cord blood hemopoietic stem andprogenitor cells. Experimental Hematology 2004;32(5):412-413.
[145] Sorrentino BP. Clinical strategies for expansion of haematopoietic stem cells. Nature
Reviews Immunology 2004;4(11):878-888.
[146] Holyoake TL et al. CD34+ positive haemopoietic cells: biology and clinical applications.
Blood Rev 1994;8(2):113-124.
[147] Majdic O et al. Signaling and induction of enhanced cytoadhesiveness via thehematopoietic progenitor cell surface molecule CD34. Blood 1994;83(5):1226-1234.
[148] Healy L et al. The stem cell antigen CD34 functions as a regulator of hemopoietic celladhesion. Proc Natl Acad Sci U S A 1995;92(26):12240-12244.
[149] Krause DS et al. CD34: structure, biology, and clinical utility. Blood 1996;87(1):1-13.
[150] Tada J et al. A common signaling pathway via Syk and Lyn tyrosine kinases generated
from capping of the sialomucins CD34 and CD43 in immature hematopoietic cells. Blood
1999;93(11):3723-3735.
[151] Gratama JW et al. Flow cytometric enumeration and immunophenotyping ofhematopoietic stem and progenitor cells. J Biol Regul Homeost Agents 2001;15(1):14-22.
[152] Lanza F et al. Structural and functional features of the CD34 antigen: an update. J Biol
Regul Homeost Agents 2001;15(1):1-13.
[153] Prosper F et al. Regulation of hematopoiesis through adhesion receptors. J Leukoc Biol
2001;69(3):307-316.
[154] Drew E et al. CD34 and CD43 inhibit mast cell adhesion and are required for optimal
mast cell reconstitution. Immunity 2005;22(1):43-57.
[155] Taussig DC et al. Hematopoietic stem cells express multiple myeloid markers:
implications for the origin and targeted therapy of acute myeloid leukemia. Blood
2005;106(13):4086-4092.

151

References[156] Tan PC et al. Na+/H+ Exchanger Regulatory Factor-1 Is a Hematopoietic Ligand for a
Subset of the CD34 Family of Stem Cell Surface Proteins. Stem Cells 2006;24(5):11501161.
[157] Giebel B et al. Segregation of lipid raft markers including CD133 in polarized humanhematopoietic stem and progenitor cells. Blood 2004;104(8):2332-2338.
[158] Goussetis E et al. In vitro identification of a cord blood CD133+CD34-Lin+ cell subsetthat gives rise to myeloid dendritic precursors. Stem Cells 2006;24(4):1137-1140.
[159] Bhatia M et al. A newly discovered class of human hematopoietic cells with SCIDrepopulating activity. Nat Med 1998;4(9):1038-1045.
[160] Hogge DE et al. Enhanced detection, maintenance, and differentiation of primitive humanhematopoietic cells in cultures containing murine fibroblasts engineered to produce
human steel factor, interleukin-3, and granulocyte colony-stimulating factor. Blood
1996;88(10):3765-3773.
[161] Denning-Kendall P et al. Cobblestone area-forming cells in human cord blood areheterogeneous and differ from long-term culture-initiating cells. Stem Cells
2003;21(6):694-701.
[162] Fibbe WE et al. Ex vivo expansion and engraftment potential of cord blood-derived
CD34+ cells in NOD/SCID mice. Ann N Y Acad Sci 2001;938:9-17.
[163] Lewis ID et al. Umbilical cord blood cells capable of engrafting in primary, secondary,and tertiary xenogeneic hosts are preserved after ex vivo culture in a noncontact system.
Blood 2001;97(11):3441-3449.
[164] Dexter TM et al. Regulation of haemopoietic stem cell proliferation in long term bonemarrow cultures. Biomedicine 1977;27(9-10):344-349.
[165] Thalmeier K et al. Establishment of two permanent human bone marrow stromal cell lines
with long-term post irradiation feeder capacity. Blood 1994;83(7):1799-1807.
[166] Tsuji T et al. A murine stromal cell line promotes the expansion of CD34high+-primitive
progenitor cells isolated from human umbilical cord blood in combination with humancytokines. Growth Factors 1999;16(3):225-240.
[167] da Silva CL et al. A human stromal-based serum-free culture system supports the ex vivo
expansion/maintenance of bone marrow and cord blood hematopoietic stem/progenitor
cells. Exp Hematol 2005;33(7):828-835.
[168] Breems DA et al. Stroma-conditioned media improve expansion of human primitive
hematopoietic stem cells and progenitor cells. Leukemia 1997;11(1):142-150.
[169] Bhatia R et al. A clinically suitable ex vivo expansion culture system for LTC-IC and
CFC using stroma-conditioned medium. Exp Hematol 1997;25(9):980-991.
[170] Verfaillie CM et al. Macrophage inflammatory protein 1 alpha, interleukin 3 anddiffusible marrow stromal factors maintain human hematopoietic stem cells for at least
eight weeks in vitro. J Exp Med 1994;179(2):643-649.
[171] Kadereit S et al. Expansion of LTC-ICs and maintenance of p21 and BCL-2 expression incord blood CD34(+)/CD38(-) early progenitors cultured over human MSCs as a feeder
layer. Stem Cells 2002;20(6):573-582.
[172] Baksh D et al. Adult human bone marrow-derived mesenchymal progenitor cells arecapable of adhesion-independent survival and expansion. Exp Hematol 2003;31(8):723732.

152

References[173] Miller CL et al. Ex Vivo Expansion of Human and Murine Hematopoietic Stem Cells. In:
Klug CA, Jordan CT, editors. Hematopoietic Stem Cell Protocols. Totowa, New Jersey:
Humana Press Inc., 2002: 189-208.
[174] Brandt J et al. Role of c-kit ligand in the expansion of human hematopoietic progenitor
cells. Blood 1992;79(3):634-641.
[175] Piacibello W et al. Extensive amplification and self-renewal of human primitive
hematopoietic stem cells from cord blood. Blood 1997;89(8):2644-2653.
[176] Ohmizono Y et al. Thrombopoietin augments ex vivo expansion of human cord bloodderived hematopoietic progenitors in combination with stem cell factor and flt3 ligand.
Leukemia 1997;11(4):524-530.
[177] Keil F et al. Effect of interleukin-3, stem cell factor and granulocyte-macrophage colonystimulating factor on committed stem cells, long-term culture initiating cells and bonemarrow stroma in a one-step long-term bone marrow culture. Ann Hematol
2000;79(5):243-248.
[178] Kawada H et al. Rapid ex vivo expansion of human umbilical cord hematopoietic
progenitors using a novel culture system. Exp Hematol 1999;27(5):904-915.
[179] Li Y et al. Human cord cell hematopoiesis in three-dimensional nonwoven fibrous
matrices: in vitro simulation of the marrow microenvironment. J Hematother Stem CellRes 2001;10(3):355-368.
[180] Okamoto T et al. Effect of heparin addition on expansion of cord blood hematopoietic
progenitor cells in three-dimensional coculture with stromal cells in nonwoven fabrics. J
Artif Organs 2004;7(4):194-202.
[181] LaIuppa JA et al. Culture materials affect ex vivo expansion of hematopoietic progenitor
cells. J Biomed Mater Res 1997;36(3):347-359.
[182] Rosenzweig M et al. Enhanced maintenance and retroviral transduction of primitive
hematopoietic progenitor cells using a novel three-dimensional culture system. Gene Ther
1997;4(9):928-936.
[183] Astori G et al. Evaluation of ex vivo expansion and engraftment in NOD-SCID mice ofumbilical cord blood CD34+ cells using the DIDECO 'Pluricell System'. Bone Marrow
Transplant 2005;35(11):1101-1106.
[184] Jiang XS et al. Surface-immobilization of adhesion peptides on substrate for ex vivo
expansion of cryopreserved umbilical cord blood CD34(+) cells. Biomaterials
2006;27(13):2723-2732.
[185] Feng Q et al. Expansion of engrafting human hematopoietic stem/progenitor cells inthree-dimensional scaffolds with surface-immobilized fibronectin. J Biomed Mater Res A
2006. In press.
[186] Hann E et al. Development of a delivery system for the continuous endogenous release ofan anti-idiotypic antibody against ovarian carcinoma. Hybridoma (Larchmt )2005;24(3):133-140.
[187] Unger RE et al. Growth of human cells on polyethersulfone (PES) hollow fiber
membranes. Biomaterials 2005;26(14):1877-1884.
[188] Yamagishi H et al. Development of A Novel Photochemical Technique for Modifying
Poly(Arylsulfone) Ultrafiltration Membranes. Journal of Membrane Science
1995;105(3):237-247.
[189] Keselowsky BG et al. Surface chemistry modulates focal adhesion composition andsignaling through changes in integrin binding. Biomaterials 2004;25(28):5947-5954.

153

References[190] Kakabakos SE et al. Colorimetric determination of reactive solid-supported primary andsecondary amino groups. Biomaterials 1994;15(4):289-297.
[191] Fujihara K et al. Guided bone regeneration membrane made of polycaprolactone /calcium carbonate composite nano-fibers. Biomaterials 2005;26(19):4139-4147.
[192] Bico J et al. Rough wetting. Europhysics Letters 2001;55(2):214-220.
[193] Wang R et al. Light-induced amphiphilic surfaces. Nature 1997;388(6641):431-432.
[194] Donaldson C et al. The CD34(+)CD38(neg) population is significantly increased inhaemopoietic cell expansion cultures in serum-free compared to serum-replete conditions:dissociation of phenotype and function. Bone Marrow Transplant 2001;27(4):365-371.
[195] Wilson CJ et al. Mediation of biomaterial-cell interactions by adsorbed proteins: a review.
Tissue Eng 2005;11(1-2):1-18.
[196] Calvi LM et al. Osteoblastic cells regulate the haematopoietic stem cell niche. Nature
2003;425(6960):841-846.
[197] Zhang J et al. Identification of the haematopoietic stem cell niche and control of the nichesize. Nature 2003;425(6960):836-841.
[198] Wright DE et al. Physiological migration of hematopoietic stem and progenitor cells.
Science 2001;294(5548):1933-1936.
[199] Bakowsky U et al. Cooperation between lateral ligand mobility and accessibility forreceptor recognition in selectin-induced cell rolling. Biochemistry 2002;41(14):47044712.
[200] Houseman BT et al. The microenvironment of immobilized Arg-Gly-Asp peptides is an
important determinant of cell adhesion. Biomaterials 2001;22(9):943-955.
[201] Francis K et al. Murine Sca-1(+)/Lin(-) cells and human KG1a cells exhibit multiple
pseudopod morphologies during migration. Exp Hematol 2002;30(5):460-463.
[202] Williams JL et al. Thrombopoietin requires additional megakaryocyte-active cytokines
for optimal ex vivo expansion of megakaryocyte precursor cells. Blood
1998;91(11):4118-4126.
[203] Piacibello W et al. Extensive amplification and self-renewal of human primitive
hematopoietic stem cells from cord blood. Blood 1997;89(8):2644-2653.
[204] Gupta P et al. Stromal fibroblast heparan sulfate is required for cytokine-mediated ex vivo
maintenance of human long-term culture-initiating cells. Blood 1996;87(8):3229-3236.
[205] Dao MA et al. Adhesion to fibronectin maintains regenerative capacity during ex vivo
culture and transduction of human hematopoietic stem and progenitor cells. Blood
1998;92(12):4612-4621.

154