Expansion and Osteogenic Differentiation of
Human Amniotic Fluid Derived Stem Cells
DISSERTATION
Presented in Partial Fulfillment of the Requirements for the Degree of
Doctor of Philosophy
in the Graduate School of The Ohio State University
By
Meimei Liu, M.S.
Graduate Program in Chemical and Biomolecular Engineering
The Ohio State University
2013
Dissertation Committee:
Dr. Shang-Tian Yang, Advisor
Dr. Jeffrey J. Chalmers
Dr. Andre F. Palmer
i
Copyright by
Meimei Liu
2013
ii
Abstract
Because of their easy accessibility and broad multipotentiality, human amniotic
fluid stem cells (hAFSCs) are emerging as an important cell source for tissue engineering
and regenerative medicine. However, due to the very recent identification of hAFSCs,
few research exploring their mass production, differentiation and medium optimization
have been carried out.
To date, hAFSC cultures are still performed in cell culture dishes, T-flasks and
multiwell plates. These traditional techniques exhibit several serious drawbacks,
including being labor-intensive, time-consuming, expensive, prone to contamination, and
difficult to scale up. For clinical applications, a large number of hAFSCs are required,
and conventional cell culture systems cannot meet this demand. In this study, a threedimensional (3-D) polyethylene terephthalate (PET) fibrous bed bioreactor (FBB) was
developed for hAFSC expansion. This novel bioreactor provide large specific surface
areas favoring hAFSC attachment and proliferation, better mimic in vivo environments,
facilitate nutrient and oxygen diffusion, and protect cells from shear damage. It achieved
155 expansion fold production of functional hAFSCs.
Bone diseases, especially osteoporosis, leading to a high risk of fracture and
deformities, bring serious issues to public health. Estrogen replacement therapy and
ii
biphosphonate have been considered to be the most effective treatment for osteoporosis
in the past 10 years. However, they could increase the risk of cancer and cardiovascular
diseases and cause acute incapacitating bone, joint, and muscle pain. AFSCs provide a
novel cell source for cell therapy, and are reported to have strong potential to differentiate
into osteoblasts and chondrocytes. Recently, some medical plant-derived estrogen-like
chemical compounds have been shown to have antiosteoporotic activity and minimal side
effects. In this study, two natural plant ingredients, naringin and curculigoside, were
investigated for their stimulation effects on proliferation and osteogenic differentiation of
hAFSCs. The results demonstrated that these two natural ingredients could promote the
proliferation and osteogenesis, and concurrently inhibit osteoclastogenesis of hAFSCs.
Moreover, signal transduction pathways underlying the promotion of osteogenic
differentiation of hAFSCs were clearly revealed.
Being a recently identified stem cell for promising clinical applications, hAFSCs
have not been well studied for their in vitro expansion. To date, very few medium
formulae have been optimized for growth and expansion of hAFSCs. Currently, most
hAFSC cultures are carried out in a commonly used serum-rich medium, which is not
qualified for good manufacturing practice (GMP). In this study, by cooperating with
scientists at Irvine Scientific, the author evaluated three newly developed and optimized
media, including a complete medium and two serum-free media, for supporting in vitro
proliferation of hAFSCs. The results indicated that the serum-containing complete (SCC)
medium could support a faster proliferation than the control medium while maintaining
the characteristic immunophenotype and the multilineage differentiation potential of
iii
hAFSCs. However, the two serum-free media SF I and SF II were not satisfactory and
further development and optimization are needed. Nevertheless, this study provided
groundwork for the development of medium for the expansion of hAFSCs for clinical
applications.
iv
Dedication
This document is dedicated to my family.
v
Acknowledgements
The pursuit of my Doctoral degree at the Ohio State University is one of the most
exciting and challenging undertakings of my life. I would love to express my deep
appreciations to my family and all my friends. Without their unconditional support, love,
understanding and encouragements, I could not successfully complete my research and
achieve this work.
The cooperation and help of our research group was essential for me, without
their support I could not accomplish any success. I would particularly like to thank Dr.
Yan Li for her careful revision of my dissertation draft. I also want to express my
appreciation to other previous group members Dr. Yan Li, Dr. Xiaoguang Liu, Dr.
Xudong Zhang, Dr. Yuan Wen, Dr. Ning Liu, and Dr. Ru Zang for their support and
technical advice in my research.
This study was partially funded by Alumni Grants for Graduate Research and
Scholarship (AGGRS) of The Ohio State University. I would also like to thank Dr.
Anthony Atala and Dr. James Yoo of the Wake Forest Institute for Regenerative
Medicine (Winston-Salem, NC) for their kindly providing hAFSCs.
vi
Moreover, I would like to express my gratitude to Dr. Jeffrey J. Chalmers, Dr.
Andre F. Palmer, and Dr. Kenneth D. Koenig for serving as my dissertation committee
members and the graduate faculty representative, respectively, and for their invaluable
and insightful comments for my work.
I want to express my final thanks to my advisor Dr. Shang-Tian Yang for his
continuous support, encouragements, seasoned guidance and friendly presence during
these years. It is his patience and high standard discipline made my study and life at the
Ohio State University productive and joyful. His understanding and patience in my
achievements in study and research were impressive and encouraging. His energetic work
attitude and loving attitude are always examples for me to study.
vii
Vita
June 2004 .......................................................B.S. Chemical Engineering,
Tianjin University, China
March 2007 ....................................................M.S. Chemical Engineering,
Tianjin University, China
September 2007 - present...............................Graduate Research Associate,
Department of Chemical Engineering,
The Ohio State University
Publications
1. Mirkelamoglu B, Liu M, and Ozkan US. 2010. Dual-catalyst aftertreatment of leanburn engine exhaust. Catalysis Today 151: 386-394.
2. Fan J, Wang K, Liu M, and He Z. 2008. In vitro evaluations of konjac glucomannan
and xanthan gum mixture as the sustained release material of matrix tablet. Carbohydrate
Polymers 73: 241-247.
3. Liu M, Fang J, Wang K, and He Z. 2007. Synthesis, characterization, and evaluation
of phosphated crosslinked konjac glucomannan hydrogels for colon-targeted drug
delivery. Drug Delivery 14:397-402.
viii
Fields of Study
Major Field: Chemical and Biomolecular Engineering
Area of Interest: Biotechnology, especially cell culture
ix
Table of Contents
Abstract ............................................................................................................................... ii
Dedication ........................................................................................................................... v
Acknowledgments.............................................................................................................. vi
Vita................................................................................................................................... viii
List of Tables .................................................................................................................. xvii
List of Figures ................................................................................................................ xviii
Chapter 1: Introduction and Literature Review ................................................................. 1
1.1 Cell Culture ................................................................................................................ 1
1.2 Stem Cells .................................................................................................................. 2
1.2.1 Embryonic stem cells (ESCs) .............................................................................. 3
1.2.2 Induced pluripotent stem (iPS) cell...................................................................... 3
1.2.3 Adult stem cells.................................................................................................... 4
1.3 Amniotic Fluid Stem Cells (AFSCs) ......................................................................... 5
1.3.1 Amniotic fluid (AF) ............................................................................................. 5
1.3.2 Amniotic fluid stem cells (AFSCs) ...................................................................... 6
x
1.4 Large-Scale Expansion and Differentiation of Stem Cells ...................................... 13
1.4.1 The motivation of large-scale cell cultures ........................................................ 13
1.4.2 Challenges in large-scale cultures and cell classes to culture ............................ 13
1.4.3 Bioreactors for suspension cell cultures ............................................................ 14
1.4.4 Bioreactors for anchorage-dependent cell cultures ............................................ 16
1.5 3-Dimensional Cell Culture ..................................................................................... 18
1.6 Optimization of Culture Medium............................................................................. 19
1.7 Objectives ................................................................................................................ 20
Chapter 2: Expansion of Human Amniotic Fluid Stem Cells in 3-Dimensional Fibrous
Scaffolds in Bioreactors .................................................................................................... 55
2.1 Introduction .............................................................................................................. 56
2.2 Materials and Methods ............................................................................................. 58
2.2.1 AFSC cultures and media .................................................................................. 58
2.2.2 Preparation of PET fibrous scaffolds ................................................................. 59
2.2.3 Static AFSC cultures in microwells ................................................................... 59
2.2.4 Dynamic AFSC cultures in fibrous fed bioreactor (FBB) ................................. 60
2.2.5 F Flow cytometry ............................................................................................... 60
2.2.6 Scanning electron microscopy (SEM) ............................................................... 61
2.2.7 Osteogenic and adipogenic differentiations ....................................................... 61
xi
2.2.8 Reverse transcriptase polymerase chain reaction (RT-PCR) ............................. 62
2.2.9 Colony-forming unit-fibroblastic (CFU-F) assay .............................................. 63
2.2.10 Analytical methods .......................................................................................... 63
2.2.11 Statistical analysis ............................................................................................ 64
2.3 Results and Discussion ............................................................................................ 64
2.3.1 AFSCs proliferation in 3-D PET scaffolds ........................................................ 64
2.3.2 AFSC expansion in the fibrous bed bioreactor .................................................. 65
2.3.3 Phenotype of bioreactor-expanded human AFSCs ............................................ 68
2.3.4 Multi-lineage differentiation of FBB-expanded human AFSCs ........................ 69
2.4 Conclusions .............................................................................................................. 70
Chapter 3: Effects of Naringin on the Proliferation and Osteogenic Differentiation of
Human Amniotic Fluid Derived Stem Cells ..................................................................... 89
3.1 Introduction .............................................................................................................. 90
3.2 Materials and Methods ............................................................................................. 92
3.2.1 Culture of human amniotic fluid derived stem cells (hAFSCs) ......................... 92
3.2.2 hAFSC treatment with naringin ......................................................................... 92
3.2.3 Cell proliferation analysis .................................................................................. 93
3.2.4 Alkaline phosphatase activity (ALP) assay ....................................................... 93
3.2.5 Alizarin red S (ARS) staining ............................................................................ 94
xii
3.2.6 Calcium assay .................................................................................................... 94
3.2.7 Reverse transcriptase polymerase chain reaction (RT-PCR) ............................. 95
3.2.8 Statistical analysis .............................................................................................. 96
3.3 Results ...................................................................................................................... 96
3.3.1 Effect of naringin on the proliferation of hAFSCs ............................................ 96
3.3.2 Effect of naringin on the ALP activity of hAFSCs ............................................ 97
3.3.3 Effect of naringin on calcium deposition ........................................................... 97
3.3.4 Effect of naringin on the expression of osteogenic markers .............................. 97
3.3.5 Effect of naringin on the osteoclast differentiation of hAFSCs ......................... 98
3.3.6 Effect of naringin on BMP and Wnt/β-catenin pathways .................................. 99
3.4 Discussion ................................................................................................................ 99
3.5 Conclusions ............................................................................................................ 103
Chapter 4: Curculigoside Improves Osteogenesis and Inhibits Osteoclastogenesis of
Human Amniotic Fluid Derived Stem Cells ................................................................... 120
4.1 Introduction ............................................................................................................ 121
4.2 Materials and Methods ........................................................................................... 123
4.2.1 Culture of human amniotic fluid stem cells (hAFSCs) .................................... 123
4.2.2 hAFSC treatment with curculigoside ............................................................... 123
4.2.3 Cell proliferation analysis ................................................................................ 124
xiii
4.2.4 Alkaline phosphatase activity (ALP) assay ..................................................... 124
4.2.5 Assay of calcium deposition ............................................................................ 125
4.2.6 Reverse transcriptase polymerase chain reaction (RT-PCR) ........................... 125
4.2.7 Statistical analysis ............................................................................................ 126
4.3 Results .................................................................................................................... 127
4.3.1 Effect of curculigoside on the proliferation of hAFSCs .................................. 127
4.3.2 Effect of curculigoside on ALP activity of hAFSCs ....................................... 127
4.3.3. Effect of curculigoside on calcium deposition................................................ 128
4.3.4 Effect of curculigoside on the expression of osteogenic genes ....................... 128
4.3.5 Effect of curculigoside on the osteoclast differentiation of hAFSCs .............. 129
4.3.6 Effect of curculigoside on Wnt/β-catenin signaling pathway .......................... 129
4.4 Discussion .............................................................................................................. 130
4.5 Conclusions ............................................................................................................ 133
Chapter 5: Optimization of Serum Containing and Serum Free Media for Expansion of
Human Amniotic Fluid Stem Cells ................................................................................. 150
5.1 Introduction ............................................................................................................ 151
5.2 Materials and Methods ........................................................................................... 152
5.2.1 Cultures and media .......................................................................................... 152
5.2.2 Expansion of hAFSCs over 3 passages ............................................................ 153
xiv
5.2.3 Morphology of hAFSC cultures....................................................................... 153
5.2.4 Flow cytometry analysis of immunophenotype ............................................... 153
5.2.5 Multilineage differentiation assays .................................................................. 154
5.2.6 Statistical analysis ............................................................................................ 155
5.3 Results and Discussion .......................................................................................... 155
5.3.1 Expansion of hAFSCs in Control and SCC medium ....................................... 155
5.3.2 Expansion of hAFSCs in SF I, SF II and SCC medium .................................. 158
5.4 Conclusions ............................................................................................................ 159
Chapter 6: Conclusions and Recomondations ................................................................ 174
6.1 Conclusions ............................................................................................................ 174
6.1.1 Mass production of hAFSCs in a 3-dimensional fibrous bed bioreactor ......... 174
6.1.2 Promoted proliferation and osteogenic differentiation of hAFSCs treated with
naringin and curculigoside ........................................................................................... 175
6.1.3 Medium development and optimization for the expansion of hAFSCs ........... 176
6.2 Recommendations .................................................................................................. 176
Apendix A: Comparasion of Effects of Naringin and Curculigoside on Osteogenesis of
hAFSCs ........................................................................................................................... 178
Apendxi B: Osteogenic Differentiation of Naringin Treated hAFSCs in 3D Dynamic
Bioreactors ...................................................................................................................... 182
Apendix C: Extracts of Chinese Herb Combinations on the Proliferation of hAFSCs .. 186
xv
Comprehensive Bibliography ......................................................................................... 189
xvi
List of Tables
Table 1.1 The early history of cell culture ....................................................................... 41
Table 1.2 Adult stem cells and their applications ............................................................. 42
Table 1.3 Surface markers expressed by hAFSCs ............................................................ 43
Table 1.4 Ex vivo AFSC differentiation induced by chemical-based media .................... 44
Table 1.5 Main characteristics of ESCs, iPS cells, AFSCs and MSCs ............................. 45
Table 1.6 Various preclinical applications of AFSCs ....................................................... 46
Table 1.7 Different types of culture medium .................................................................... 48
Table 2.1. Primers for RT-PCR analysis of osteogenic and adipogenis differentiations. 78
Table 2.2. Cell number and expansion fold of human AFSCs in 3-D bioreactor. ............ 79
Table 2.3. CFU-F assay for human AFSCs before and after expansion in 3-D bioreactor.
........................................................................................................................................... 80
Table 3.1. Primers used in the RT-PCR for osteogenic differentiation of hAFSCs ....... 112
Table 4.1. Primers used in the RT-PCR for osteogenic differentiation of hAFSCs ....... 141
Table 4.2. Effects of curculigoside on the osteoclast differentiation of hAFSCs ........... 142
Table 5.1 Doubling time and seeding density of cultures in Control medium and SCC
medium over three passages ........................................................................................... 163
xvii
Table 5.2 Doubling time and seeding density of cultures in SF I, SF II and SCC medium
over three passages ......................................................................................................... 164
xviii
List of Figures
Figure 1.1 Differentiation of ESCs ................................................................................... 49
Figure 1.2 Isolation and expansion of hAFSCs ................................................................ 50
Figure 1.3 AFSCs differentiation into lineages representative of the three embryonic
germ layers ........................................................................................................................ 51
Figure 1.4 3-D fibrous PET for cell culture ...................................................................... 52
Figure 1.5 Media pyramid: a modular approach for the development of serum-free media
........................................................................................................................................... 53
Figure 1.6 Objectives and outline of this work ................................................................. 54
Figure 2.1 A fibrous bed bioreactor modified from a spinner flask with a PET matrix
around the wall. ................................................................................................................. 81
Figure 2.2 Human AFSCs expansion in 3-D PET matrix................................................. 82
Figure 2.3 Metabolic activities of human AFSCs expanded in 3-D bioreactor ................ 83
Figure 2.4 SEM images of human AFSCs expanded in 3-D PET fibrous matrices ......... 84
Figure 2.5 Phenotype of human AFSCs before and after expansion in 3-D bioreactor ... 85
Figure 2.6 CFU-F assay of hAFSCs before and after FBB expansion ............................. 86
xix
Figure 2.7 Multi-lineage differentiation of human AFSCs after expansion in 3-D
bioreactor ......................................................................................................................... .87
Figure 2.8 RT-PCR analysis of differentiated human AFSCs after expansion in 3-D
bioreactor .......................................................................................................................... 88
Figure 3.1 Chemical structure of naringin ...................................................................... 113
Figure 3.2 Effect of naringin on the proliferation of hAFSCs ........................................ 114
Figure 3.3 ALP activity of hAFSCs after naringin treatment ......................................... 115
Figure 3.4 Osteogenic differentiation of hAFSCs after naringin treatment.................... 116
Figure 3.5 RT-PCR analysis of naringin-enhanced osteogenic differentiation of hAFSCs.
......................................................................................................................................... 117
Figure 3.6 RT-PCR analysis of naringin-enhanced BMP and Wnt signaling of hAFSCs
......................................................................................................................................... 118
Figure 3.7 Schematic illustration of BMP and Wnt-signaling pathways in naringinenhanced osteogenic differentiation of hAFSCs............................................................. 119
Figure 4.1 Chemical structure of curculigoside .............................................................. 143
Figure 4.2 Effect of curculigoside on the proliferation of hAFSCs ................................ 144
Figure 4.3 ALP activity of hAFSCs after curculigoside treatment ................................. 145
Figure 4.4 Calcium deposition of hAFSCs after curculigoside treatment ...................... 146
Figure 4.5 RT-PCR analysis of curculigoside-enhanced osteogenic differentiation of
hAFSCs ........................................................................................................................... 147
Figure 4.6 RT-PCR analysis of curculigoside-enhanced Wnt signaling of hAFSCs ..... 148
Figure 4.7 Schematic illustration of Wnt signaling pathway in osteogenic differentiation
of hAFSCs ....................................................................................................................... 149
xx
Figure 5.1 Scheme of medium optimization experiments .............................................. 165
Figure 5.2 hAFSCs growth over three passages in control medium and SCC medium . 166
Figure 5.3 hAFSC morphology in Control culture and SCC culture over 3 passages.... 167
Figure 5.4 Phenotype of hAFSCs cultured in Control medium and SCC medium after 3
passages analyzed by flow cytometry ............................................................................. 168
Figure 5.5 Histochemistry and immunostaining of multipotent differentiation of hAFSCs
expanded in Control and SCC medium .......................................................................... 169
Figure 5.6 hAFSCs growth over three passages in SF I, SF II and SCC mediums ........ 170
Figure 5.7 Cell viability of hAFSCs in SF I, SF II, and SSC cultures over 3 passages . 171
Figure 5.8 hAFSC morphology in SF I, SF II and SCC mediums over 3 passages ....... 172
Figure 5.9 Phenotype of hAFSCs cultured in SF I, SF II and SCC mediums after 3
passages analyzed by flow cytometry ............................................................................. 173
Figure A.1 ALP activity of hAFSCs treated by naringin and curculigoside .................. 180
Figure A.2 Calcium deposit of hAFSCs treated by naringin and curculigoside ............. 181
Figure B.1. A fibrous bed bioreactor modified from a spinner flask with a PET matrix
around the wall used for osteogenic differentiation of hAFSCs ..................................... 184
Figure B. 2. Calcium deposition of naringin treated hAFSCs in 2D static culture and 3D
dynamic bioreactor.......................................................................................................... 185
Figure C.1 Proliferation of (A) PG and (B) RRPG treated hAFSCs .............................. 188
xxi
Chapter 1
Introduction and Literature Review
1.1 Cell Culture
Cell culture is the process to continually grow animal cells under controlled
conditions after they were removed from animal tissues, which inaugurated a new era in
biology and medicine. Cell culture process has been widely used in a large number of
areas, including studying the biochemistry and physiology of cells, testing the effects of
drugs and other chemical compounds on specific cell types, generating artificial tissues in
vitro, synthesizing valuable biologicals such as therapeutic proteins and viruses from
large-scale cultures, as well as investigating diseases in vitro [1, 2].
The concept of animal cell culture dates back to the 19th century (Table 1.1). In
1885, Wilhelm Roux successfully removed and maintained a portion of the medullary
plate of an embryonic chicken in a warm saline solutions for several days [3]. Ross
Harrison in 1907 demonstrated not only maintenance but also growth of frog embryo
nerve fiber in vitor [4]. Animal cell culture became an established laboratory technique in
the 1950s with the development of defined culture media by Eagle et al. [5]. Another
remarkable milestone came in the 1920s with the discovery of antibiotics by Fleming
1
which forwarded continual cell culture by reducing contamination problems [2]. In the
1940s and 1950s major epidemics stimulated the development of virology research and
the manufacture of vaccines [2, 3]. Recombinant DNA technology developed in the
1970s quickly made the production of therapeutic proteins from animal cells possible [2].
The generation of hybridomas by Kohler and Milstein in 1975 was another milestone that
facilitated the continuous production of antibody molecules [6].
A large number of cell types like epithelial cells, fibroblasts, muscle cells, nerve
cells, cardiac cells, mesenchymal cells, endocrine cells, and stem cells, have been
successfully cultured [1]. Besides cell biology study and drug screening, animal cells
from cultures have also been widely applied as end products, such as artificial skin grafts,
islet cells, hepatocytes, and bone marrow implants, in regeneration medicine, as well as
to be used to produce recombinant and natural proteins including human growth hormone,
nerve growth factor, epidermal growth factor (EGF), monoclonal antibodies (MAb),
vaccines, interferons, and blood clotting factors [1].
1.2 Stem Cells
Stem cells are the cells that can renew themselves through mitotic division and
differentiate into other specialized cell types. Because of these two properties, selfrenewal and differentiation, stem cells have attracted much attention in biology science
and therapy research. According to the potency, the capacity of differentiation, stem cells
are generally classified into two types: adult stem cells and embryonic stem cells. Adult
stem cells are derived from adult tissues and can differentiate into a certain amount of
cell types; while embryonic stem cells are isolated from the inner cell mass of early
2
developing embryo and can differentiate into three germlayer cell types. Stem cell culture
provides cells as end product and has attracted much attention for regenerative medicine
and tissue engineering applications.
1.2.1 Embryonic stem cells (ESCs)
Embryonic stem cells were first derived in 1981 from mouse embryos [7, 8].
Shortly after fertilization, the fertilized egg divides into morula and then forms blastocyst
at day 4-5 day. The inner cell mass of blastocyst is pluripotent and generates the three
germ layers, including ectoderm, endoderm, and mesoderm, that differentiates into all
tissues of the body [9]. ESCs are isolated from the inner cell mass of blastocyst of the 4-5
day embryo. It was not until 1998 researchers first isolated and grew human ESC in cell
culture [10]. ESCs are pluripotent and have been reported to differentiate into all three
germ layer cells, such as cardiogenic cells, myogenic cells, hematopoietic cells,
neurongenic cells, skeletal muscle cells, vascular smooth muscle cells, epithelial-like
cells and pancreatic cells [11]. Figure 1.1 illustrates the differentiation of ESCs. To date,
around 400 human ES cell lines have been established in more than 20 countries to date
[12]. However, ESC research cause serious ethical issues and are limited in many
countries over the world.
1.2.2 Induced pluripotent stem (iPS) cell
Recently, a new embryonic stem like cell, induced pluripotent stem (iPS) cell,
was developed by scientists [13]. They converted mouse fetal fibroblasts to iPS cells by
retrovirally transfecting Oct3/4, Klf4, Sox2, and c-Myc four genes. iPS cells were
3
reported to have the same specific marker and gene expressions as ES cells, and their
ability to form teramas showed their pluripotency [14]. Shortly after mouse iPS cells,
human iPS cells were also derived from human dermal fibroblast cells [15]. iPS cells
possess significant advances, not relating to ethical issues and immunity rejections, and
they have appealed to researchers’ interests. However, some concernful problems were
reported: the differentiation efficiency of iPS cells is much lower than that of ES cells,
and the cells derived from iPS cells exhibited limited expansion and early senescence
[16].
1.2.3 Adult stem cells
Adult stem cells exist in specific tissues or organs in the body. They generate
some or all of the major specialized cell types of the tissue or organ, and are involved in
the continual maintenance, replenishment, and repair of the tissues or organs throughout
the life span of the individual. Adult stem cells are widely found throughout the body,
their sources, differentiations and applications are summarized in Table 1.2. The
discovery of adult stem cells has generated a lot of excitement in cell biology and
transplant clinical areas. It opens new avenues for biological research and applications by
using adult stem cells as an alternative to embryonic stem cells [17].
Research on adult stem cells started about 50 years ago [18]. Becker et al.
determined that hematopoietic stem cells (HSCs) were present in bone marrows and
could restore damaged tissues in 1963 [19]. A few years later, in the 1970s, mesenchymal
stem cells (MSCs) were discovered in bone marrow by Friedenstein and coworkers [20].
4
These non- hematopoietic stem cells compose a small part of bone marrow stromal cells,
and can derive bone, cartilage, fat, and fibrous connective tissues [18]. In 1966, Altman
and Das first observed the existence of neural cells that could divide and ultimately
become neurons in the embryonic mammalian central and peripheral nervous system [21].
It was not until 1992, however, scientists agreed that neural stem cells (NSCs) also
appear in the adult mammalian central nervous system [22].
Traditionally, adult stem cells are considered to only be able to generate a certain
cell types restricted to their tissue or origin. However, in the early 2000s, multipotent
adult progenitor cells have been revealed in several tissues [23]. They possess remarkable
self-renewal ability, express embryonic stem-specific transcription factor Oct3a/4, and
own remarkable differentiation ability akin to ESCs [22]. The attractive pluripotent
capacity has greatly promoted basic and applied researches on these multipotent adult
stem cells.
1.3 Amniotic Fluid Stem Cells (AFSCs)
Due to their easy accessibility and broad multipotentiality, human amniotic fluid
stem cells (hAFSCs) are emerging as a promising adult stem cell for tissue engineering
and cell therapy of human diseases.
1.3.1 Amniotic fluid (AF)
Amniotic fluid (AF) is a clear, slightly yellowish watery liquid that surrounds the
developing fetus within the amniotic sac. It grants the unborn baby to freely move and
grow inside the womb, cushions and protects it from outside injuries, keeps a relatively
5
constant temperature around it, and provides exchange body chemicals between it and the
mother [24-26].
The average volume of AF over the course of gestation is 800 ml [25]. It is
mainly made up by water and electrolytes (98-99%), chemical substances (such as
glucose, lipids, proteins, hormones and enzymes), suspended materials (such as vernix
caseosa, lanugo hair and meconium) and cells [24]. AF cells are believed shed from both
extra-embryonic structures (such as placenta and fetal membranes) and embryonic and
fetal tissues [27, 28]. These cells are indicated to express markers of all three germ layers
[29]. Human AF cells have been used as a procedure to diagnose embryonic
chromosomal, structural, biochemical, and genetic anomalies for more than 50 years [24,
30, 31]. It presents a low risk for both the mother and the fetus.
The majority AF cells have been believed to be terminally differentiated and
possess limited proliferation potentials [32, 33]. It was only in the 1990s, however,
scientists discovered two subsets of AF cells harbouring a proliferation and
differentiation capacity. In 1993, Torricelli et al. demonstrated the presence of
haematopoietic progenitors in AF [34]. Several years later, Streubel et al. derived
myocytes from AF cells in 1996, indicating the presence of non-haematopoietic
precursors in AF [35]
1.3.2 Amniotic fluid stem cells (AFSCs)
The first evidence of the presence of pluripotent stem cells in AF was indicated by
Prusa et al. in 2003 by discovering expression of the pluripotency marker Oct4 in a
6
distinct sub-population (0.1-0.5%) of proliferating AF cells [24, 36]. Oct4 is a specific
transcription factor of ESCs and germ cells, which maintains ESC and germ cell selfrenewal and differentiation capacities [24, 37-39]. Thereafter, different research groups
clearly demonstrated the presence of an AF cell population able to differentiate into all
three embryonic germ layer lineages [40-43]. These cells are called amniotic fluid stem
cells (AFSCs) and characteristically express the surface antigen c-kit (CD117) [24].
1.3.2.1 Isolation and culture of AFSCs
AFSCs can be derived from small amount of 2nd-trimester AF or amniocentesis
waste. Figure 1.2 depicted the isolation and expansion process of hAFSCs. The cells are
isolated from AF according to the two-step protocol in the prior immunological selection
of c-kit positive cells and subsequently expanded in culture [40, 44]. Briefly, hAFSCs are
isolated through positive selection for cells expressing the membrane receptor c-kit, that
binds to the ligand stem cell factor [45]. Fluorescence-activated cell sorting (FACS)
analysis indicated that around 0.8% to 1.4% of AF cells are c-kit positive [46]. For the
first week after they are isolated, the progenitor cells maintain a round shape in nontreated culture dishes and exhibit a very low proliferation capacity. From the second
week, the cells start to adhere to the plate, become more elongated, and grow more
rapidly. They need a subculture upon reaching 80% confluency every 48 to 72 hours.
Isolated hAFSCs are commonly expanded in serum-rich medium, containing 17% of fetal
bovine serum and Chang supplement, and no feeder layers are required. These cells show
a high self-renewal capability with over 250 population doublings [40].
7
1.3.2.2 Chracterization of AFSCs
AFSCs present a fibroblast-like to an oval-round shape, and possess an extensive
clonogenic capacity[40, 47]. They are not tumorigenic and have extensive self-renew
capacity which can expand over 250 population doublings [40]. Additionally, AFSCs can
be derived from amniotic fluid which is regarded as medical waste after amniocentesis
and thus can be easily acquired without ethical issues.
Different studies have determined the cell-surface antigenic profile of AFSCs
through flow cytometry (Table 1.3). Cultured hAFSCs are positive for the human
embryonic stem cell marker Oct4 and the embryonic stage-specific surface marker
SSEA4, both of which are typical of the undifferentiated state of ESCs. AFSCs also
express mesenchymal and neuronal stem cell markers (CD73, CD90, CD105, CD29, and
CD44) and antigens belonging to the major histocompatibility complex I (MHC-I). They
do not express embryonic stage-specific surface marker SSEA1, haematopoietic and
endothelial markers (CD14, CD34, CD45, CD133, CD31), and antigens belonging to the
major histocompatibility complex II (MHC-II) [24, 46]. This immunophenotypic profile
shows that AFSCs express some major markers of ESC phenotype, but not the whole
complement, indicating that AFSCs are not as primitive as ESCs but possess greater
potential than most adult stem cells [46]. Although the embryonic bodies formed from
AFSCs in vitro are stained positive for all three germ layer markers, these cells do not
generate teratomas when they were implanted into immunodeficient mice [10].
8
1.3.2.3 Differentiation of AFSCs
AFSCs are able to differentiate towards tissues representative of all three
embryonic germ layers, including such as adipogenic, osteogenic, myogenic, endothelial,
neurogenic, hepatic lineages and so on [40, 46] (Figure 1.3). Therefore, AFSCs have
emerged as a promising cell source for tissue engineering and regenerative medicine
applications. Each differentiation pattern induced by the chemical-based culture media
conditions was summarized in Table 1.4.
1.3.2.4 Comparison of AFSCs with other stem cells
Currently, ESCs, iPSCs, AFSCs and MSCs are four of the most widely studied
stem cell populations, and their main characteristics were compared and summarized in
Table 1.5. ESCs and iPSCs are pluripotent, however, their efficient differentiations are
difficult to carry out and they form teratomas when injected in vivo [46]. MSCs are less
multipotent and show relatively low proliferation capacity. In contrast, AFSCs exhibit
some significant advantages over these stem cells. First, AFSCs do not involve in any
ethical issues of ESCs and do not generate teratomas in vivo, thus in terms of
practicability, they possess great potential for prospective application in clinical trials. In
addition, AFSC culture does not need feeder cells, have a shorter doubling time and can
be easily induced to three germ layer cell types [40, 46]. At last, AFSCs express
transcriptional factor Oct4 and SSEA4, indicating that they partially maintain the
undifferentiated state and pluripotency in ESCs [48] and possess more broad
multipotency than other adult stem cells.
9
1.3.2.5 Preclinical applications of AFSCs
Although AFSCs were identified very recently, several explorations of their
application in preclinical regenerative medicine have been carried out (Table 1.6).
De Coppi et al. transplanted AFSCs into rat cryo-injured bladders [49]. A few small
smooth muscle bundles and limited vasculogenesis were formed, and cryo-injury
induced hypertrophy of the surviving smooth muscle cells was prevented.
AFSCs were cultured in neuronal differentiation medium and grafted into the
lateral cerebral ventricles of control mice and the ventricles of twitcher mice [40]. It is
found that more AFSCs integrated into the injured twitcher mice brains (70%) than into
the control mice brains (30%). A recent research studied the neuronal differentiation
ability of rat AFSCs and their effects on injured avian embryos [50]. AFSCs were grafted
at the site of an extensive thoracic crush injury in E2.5 chick embryos, and they were
indicated to remarkably reduce hemorrhage and increase survival.
Human AFSCs were microinjected into murine embryonic kidneys and were
found to be able to contribute to the development of various primordial kidney structures
[46]. Moreover, in a renal injury model, injected AFSCs were found to ameliorate acute
tubular necrosis (ATN) as reflected by decreased blood urea nitrogen (BUN) and
creatinine levels as well as to decrease the amount of damaged tubules and apoptosis [51].
Carraro et al. microingected hAFSCs into murine lung after injury and found that
the cells differentiated into pulmonary lineages by expressing specific alveolar versus
bronchiolar epithelial cell lineage [52]. Significantly, cell fusion phenomena were
10
excluded and no tumour formation was observed in the treated animals up to 7 months
after AFSCs injection.
Bollini et al. demonstrated that rat AFSCs could differentiate into myocardial
phenotypes and improve heart function a rat myocardial infarction model, however, their
potential is limited by poor survival in an allogeneic setting [53]. In addition, a variety of
AFSC derived cellular structures were developed and tested in animal heart infarction
models [54]. Spherical cell aggregations [55] and cell sheet fragments [56] generated
from AFSCs on methylcellulose hydrogel systems were implanted into infarcted sites in
rat hearts. Both cellular structures were found to reduce cell loss, produced an enriched
extracellular matrix environment, and include expression of several angiogenic and
cardioprotective factors. Chiavegato et al. studied AFSCs differentiation capacity towards
cardiac and vascular lineages [57]. Under in-vitro cardiovascular inducing conditions,
hAFSCs were found to express cardiomyocyte (Nkx2.5, MLC-2v, GATA-4, β-MyHC),
endothelial (angiopoietin, CD146) and smooth muscle (smoothelin) markers.
Neonatal valve tissues were constructed with biodegradable scaffolds seeded with
hAFSCs. These tissues were found to contain viable endothelium which showed stable
mechanical strength similar to native tissues [58].
Ovine AFSCs were seeded on an acellular hydrogel to construct engineered
tendon structures. Implanted into a partial diaphragmatic defect in newborn lambs, these
cellular tissues facilitated better functional and mechanical outcomes comparing to
acellular bioprostheses [59].
11
AFSCs were osteogenically differentiated and seeded in alginate/collagen
scaffolds, and then were implanted into immunodeficient mice [40]. Highly-mineralized
tissues and blocks of bone-like material were observed in the recipient mice after 18
weeks. Sun et al. induced osteogenic differentiation of hAFSCs by bone morphogenic
protein 7 (BMP-7) [30]. Meanwhile, they used nanofiber scaffolds that mimic in vivo
collagen fibers to facilitate osteogenesis of hAFSCs. Peister et al applied poly (εcaprolactone) (PCL) biodegradable scaffold to support the osteogenic differentiation of
hAFSCs in vitro and subsequent ectopic bone formation after implantation [60]. To cure
postnatal sternal repair, rabbit AFSCs seeded biodegradable nanofibers were implanted
into full-thickness sternal defects [61]. Two months later, chest closure and bone
formation were confirmed by in vivo imaging modalities.
The potential of hAFSCs to differentiate into functional chondrocytes has also
been demonstrated. Human AFSCs treated with TGF-β1 form remarkable amounts of
cartilaginous matrix (such as type II collagen and sulfated glycosaminoglycans) in both
pellet and alginate hydrogel cultures [44].
Very recently, the support effects of AFSC-secreted biological factors have been
studied [55, 56]. Teodelinda et al. indicated that conditioned medium (CM) from AFSC
cultures contained pro-angiogenic soluble factors, including stromalderived factor (SDF)1, interleukin (IL)-8, monocyte chemotactic protein (MCP)-1, and vascular endothelial
growth factor (VEGF) [46, 62]. This CM is found to prevent muscle tissue necrosis and
capillary loss, and consequently induce neo-arteriogenesis and remodeling of pre-existing
collateral arteries, after injected into a mouse hind-limb ischemic model.
12
1.4 Large-Scale Expansion and Differentiation of Animal Cells
1.4.1 The motivation of large-scale cell cultures
Cell cultivation was first carried out over a century ago. However, for several
decades, most cell cultures were restricted in cell culture dishes, T-flasks and multiwell
plates. These traditional technologies are labor-intensive and difficult to scale up.
Moreover, continuous monitoring and control of culture parameters, such as pH, DO, and
shear force, are hard to achieve within a dish culture, where undesired differentiation of
stem cells were observed due to uncontrolled local concentration gradients of nutrients
and regulation reagents [63]. As the demand of mass vaccines, recombinant proteins,
monoclonal antibodies and animal cells, suitable bioreactor processes for large scale cell
cultures are highly sought.
1.4.2 Challenges in large-scale cultures and cell classes to culture
The major obstacles in scale-up cultures are oxygen supply limitation, waste
metabolite accumulation, shear damage to cells, automatic process control, and growing
anchorage-dependent cells [64]. In term of the use of bioreactors, animal cells can be
categorized into two classes: suspension cells and anchorage-dependent cells. In general,
suspension cell cultures are homogenous and easy to automatically control and thus are
preferred in industry. Most animal cells are adapted to suspension cultivation prior to
scale-up. However, anchorage-dependent cell cultures are also performed in industry,
especially for vaccine production. Compared to suspension cell cultures, anchoragedependent cell cultures are more difficult to scale up.
13
1.4.3 Bioreactors for suspension cell cultures
1.4.3.1 Stirred tank reactors
Stirred tank bioreactors (STR) are the simplest and most widely used bioreactor
type for suspension cell cultures. Various suspension cells and cells adapted to growth in
suspension including CHO, BHK 21, HEK 293, hybridoma cells, have been successfully
cultured in STR [65]. STR has been used to produce commercial monoclonal antibodies
[66, 67], recombinant proteins such as tPA, blood coagulation factor VIII and
erythropoietin [68, 69], vaccines [70], growth factors, and interferon [71]. Large-scale
STR for animal cells can be up to several thousand cubic meters, the maximum working
volume has been reported to be 15,000L [65]. STR was adopted from microbial
fermentation. However, animal cells are more fragile and sensitive to shear force and
grow more slowly than bacteria or fungi. Therefore, animal cells require more gentle
culture conditions and control systems which are fitted for lower metabolism [72]. Large
impellers with an axial fluid flow characteristics, for example, marine impellers,
segmented impellers, and large paddle impellers, are typically applied to achieve
nonturbulent bulk flows at minimum shear forces for animal cell culturing in STR [7377]. Turbine impellers used in microbial fermenters that cause turbulence should be
avoided in cell culture bioreactors. Damages to the cells caused by mechanical agitation
and gas sparging are the main issues with STR [78-80]. Another problem is forming
which kills the cells by absorbing cells and bubble rupturing and as well as causes
contamination [81, 82].
14
1.4.3.2 Airlift reactors
Airlift reactor is composed of a tall column with an inner draught tube [83], and it
provides mixing through the introduction of gas bubbles at the bottom of the tall column
[84]. Airlift reactor is easy to scale up and reliable for sterile operation due to its simple
structure [84, 85]. This type of reactor has been used for suspension growth of hybridoma,
BHK, CHO, Namalva cell lines [86-89]. 2 m3 of airlift reactor has been reported to be
routinely used at LONZA for monoclonal antibody production [85, 90]. Celltech used
airlift reactors with over 1000 L to produce monoclonal antibodies from hybridomas [72].
This type reactor has advantages such as flexible working volume and gentle mixing [84],
however, it causes cell damage and CO2 accumulation due to direct sparging [91].
1.4.3.3 Hollow fiber bioreactors
Hollow fiber bioreactors consist of bundles of synthetic and semipermeable
hollow fibers. In this kind of reactor, cells grow in the extra-capillary space, medium
recirculates in the intra-capillary space, and hydrostatic pressure achieves the exchange of
nutrients and metabolic waste across the capillary wall. Very high cell densities, for
example, 2×108 cells/ml, can be obtained in hollow fiber bioreactors [92, 93]. However,
the major limitation of this type of bioreactor is the nutrient gradients and uneven cell
growth due to the pressure difference along the fibers, which increases the difficulties to
scale up. In addition, hollow fiber bioreactor is poorly stable because of fouling of the
biber membrane.
15
1.4.4 Bioreactors for anchorage-dependent cell cultures
1.4.4.1 Microcarrier culture in stirred tank reactors
Microcarrier culture was developed to provide similar scale-up and environmental
control properties as suspension cell cultivation in stirred bioreactors for growing
anchorage-dependent cells. Microcarriers are small solid particles made of dextran,
plastic, gelatin, glass, or cellulose and have diameters of around 100-200 µm [94, 95].
Size, density, surface charge and other chemical and physical properties of the carrier are
important. Generally, nontoxicity, good adhesion properties, and a suitable buoyant
density to be easily suspended are required [65]. Usually the surface of the microcarriers
is coated or modified with collagen, proteoglycans, fibronectin, laminin, elastin, and
chondronectin to improve cell attachment [94]. Porous microcarriers are also used to
increase surface area and protect shear sensitive cells inside the beads [96]. Microcarrier
culture has been used for the large-scale production of vaccines [70, 97, 98], interferon
[99, 100], and recombinant therapeutic proteins [101-103]. This technology has many
advantages for anchorage-dependent cell culture including high growth surface-to-culture
volume ratio, direct monitoring and control, simple cell separation and perfusion.
However, mixing and aeration are challenges for microcarrier cultures. It is reported that
cells grown on smooth carriers are more sensitive to overagitation, and growth arrest and
cell detachment occur at high stirrer rates [74, 104, 105]. Oxygen supply is another issue
for large-scale microcarrier culture and buble-free aeration methods are generally
preferred to reduce foam layers and cell damages [106].
16
1.4.4.2 Packed bed bioreactors
Packed bed bioreactor is composed of a static bed and solid inert particles which
are generally glass beads with 3-5 mm of diameter [72]. This type of bioreactor has been
used to produce vaccine [107, 108], interferon [109], herpes simplex virus [110], tissue
plasminogen activator, and acetylcholinesterase [111]. Besides solid glass beads, porous
glass spheres are also used [112, 113]. Packed bed bioreactor can be applied to
immobilize anchorage-dependent or suspension cells by offering high cell densities in the
macroporous structures for the generation of therapeutic proteins and lytic viruses [65].
Packed bed bioreactors own some major advantages, such as high cell density, low
surface shear rates, high productivity, as well as no particle-particle abrasion. However,
some disadvantages are also reported, for example, poor oxygen transfer, blockage of the
pores because of high cell densities, and the risk of medium channeling in the bed [114].
1.4.4.3 Fluidized bed bioreactors
In fluidized bed bioreactors, cells are immobilized or entrapped in porous
microspheres which are suspended in the column by high-velocity upward flow of culture
medium. The microcarriers can be made by collagen, alginate, borosilicate glass, and
polyethylene [65, 72]. A heavy metal such as noncytotoxic steel can be added to achieve
a high bead density to make the carriers remain suspended in the high-speed upward-fluid
flow of medium. The carriers have sponge-like structure of interconnected pores and
channels to allow cells to enter and populate inside of the carriers [115]. Fluidized bed
bioreactor has been used to large-scale culture CHO cells and hybridomas and other
17
anchorage-dependent and suspension cells to produce proteins [116-118]. A main
disadvantage of this type of bioreactor is the progressive depletion of oxygen along the
axis of the reactor bed. Moreover, reliable measurement of the cell density in
immobilized cultures was another issue [65].
1.5 3-Dimensional Cell Culture
As introduced above, most stem cells are conventionally cultured on 2-D culture
surfaces like culture dishes, T-flasks and multiwell plates, which cannot mimic in vivo
environment where multi-dimensional cell-cell contact plays critical roles on maintaining
cellular functions. The cell density in vivo tissue is over 109 cells/ml tissue, while the
maximum cell density reported in 2-D culture is 106-107 cells/ml [12]. Therefore, current
2-D culture systems cannot actually simulate body environment for cell production and
drug toxicity screening. In the contrast, 3-D scaffold culture can provide high cell density
and better mimic in vivo environment. 3-D scaffolds can provide larger specific surface
areas favoring cell attachment and growth, and highly porous structures of 3-D scaffolds
facilitate nutrient and oxygen diffusion. Additionally, 3-D scaffolds can protect cells
from shear force so to decrease its damage to the cells. In addition, it was reported that 3D scaffolds play a significant role in enhancing stem cell proliferation and promoting
desired differentiation [119, 120]. Therefore, 3-D scaffolds have attracted more and more
attention for cellular transplantation and drug screening.
Many materials have been studied as stem cell culture scaffolds, including
polylactic acid, alginate, polyethylenglycol and so on [121, 122]. Our group has been
studying non-woven fibrous polyethylene terephthalate (PET) as stem cell culture
18
scaffold for years and obtained significant success (Figure 1.4). PET is one of the first
synthetic polymers used in regenerative medicine and has been used for vascular
reconstructions [123] as well as bone cells and endothelial cells cultures [124]. In our
group, over 4×108 cells/ml matrix has been reached, which is much higher than 2-D
culture and almost at the same level as in vivo tissue [12, 125, 126]. Compared to 2-D
systems, longer ESC proliferation period was achieved by using PET matrix in our lab
[11]. In addition, besides expansion we also successfully differentiated ESCs into
hematopoietic cells [127] and neutral cells [11, 12] in PET fibrous matrixes, respectively.
It was found that PET scaffold can promote hematopoietic and neutral differentiation of
ESCs.
1.6 Optimization of Culture Medium
Cell culture medium is a mixture consisting of amino acids, a source of energy
(such as glucose), vitamins, growth factors, trace elements, etc. in a pH buffered salt
solution [128]. Traditional mammalian cell culture formulations require further
supplementation with a protein source, such as serum, to maintain and proliferate cells.
Fetal bovine serum (FBS) is the present standard serum. It is a complex mixture
containing a large number of ingredients, such as proteins, growth factors, hormones,
vitamins, trace minerals and so on, which are essential for mammalian cells [129].
However, the serum composition continually varies with season and producing batch and
is ill-defined. Moreover, because of the threat of contamination of viral, bacterial, and
prion pathogens, the use of animal-based products is firmly dejected for production of
medicinal products [130-132]. It is also reported that exposure of human cells to FBS
19
resulted in fixation of animal proteins on the human cell surface thus made the host more
prone to inflammatory and/or adverse immunemediated events [133-135]. Therefore,
mammalian cell culture media are directed to progress from serum-containing to serumfree, to animal-component-free and then to chemically defined formulations (Table 1.7)
[128, 129]. Figure 1.5 briefly depicts a modular approach for the development of serumfree media.
1.7 Objectives
This study aimed to investigate and promote the development of the expansion
and osteogenic differentiation of hAFSCs, and it was divided into three parts (Figure 1.6):
1) develop a PET based 3-dimensional bioprocess for large-scale expansion of
functional hAFSCs to promote theri clinical transplantation applications (Chapter 2);
2) promote the proliferation and osteogenic differentiation of hAFSCs by using
natural plant ingredients, investigate the responsible signaling pathways, and study the
potential treatment strategy to cure osteoporosis and other bone disorders (Chapter 3 and
Chapter 4);
3) develop and optimize media for the expansion of hAFSCs for clinical
applications (Chapter 5).
20
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Table 1.1 The early history of cell culture.
Time
Event in cell culture history
1885
Roux maintained embryonic chick cells in saline solution for several days
[3]
1907
Harrison grow frog nerve cells [4]
1912
Carrel cultured connective tissue cells for prolonged periods and exhibited
heart muscle tissue contractility over 2-3 months [72]
1920s
The antibiotics (e.g. penicillin) were added to culture medium [2]
1948
Earle cloned mouse L fibroblasts [73]
1952
Gey established HeLa cells (the first human cell line) from a cervical
carcinoma [74]
1954
Abercrombie and Heaysman observed contact inhibition between
fibroblasts [75]
1955
Eagle et al. developed chemically defined culture media [5]
1961
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finite lifespan [76]
1962
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from tumor [77]
1968
Yaffe investigated the differentiation of normal myoblasts [78]
1975
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1980
Human insulin was produced from bacteria [79]
41
Table 1.2 Adult stem cells and their applications.
Adult stem cells
Source
Derivates
Applications
Hematopoietic
stem cells
Bone marrow,
cord blood,
peripheral blood
Leukemia, and
blood related
disease [80]
Mesenchymal
stem cells
Bone marrow,
adipose,
peripheral blood,
placenta,
umbilical cord,
amniotic fluid
Subventricular
zone, dentate
gyrus in the
hippocampus
Liver
T-cells, B-cells, NK cells,
granulocyte/monocyte
progenitors,
megakaryocyte/erythrocyte
progenitors
Osteoblasts, chondrocytes,
and adipocytes, neurons,
hepatocytes, pancreatic
islet cells, cardiomyocytes,
endothelial cells, pericytes,
smooth muscle cells
Neurons,
oligodendrocytes,
astrocytes
Hepatocytes, biliary cells
Liver repair [90]
Pancreas
Adult ductal cells, β cells
Islet formation [91]
Adipose
Adipocytes, osteoblasts,
myoblasts, chondroblasts
Soft tissue
cosmesis, cartilage
repair, muscle
repair, bone defeat
repair [92]
Skeletal-muscle
stem cells
Muscle fibers
Muscle
regeneration [93]
Skin stem cells
Epidermis, hair
follicles
Trachea,
bronchiole, lung
Hematopoietic cells,
skeletal muscle, satellite
cells
Epidermis, hair follicles
Neural stem cells
Hepatocyte stem
cells
Pancreatic stem
cells
Adipose stem
cells
Lung epithelial
stem cells
Intestinal
epithelium stem
cells
Epithelium
Mucous and ciliated cells,
pneurnocytes
Inflammation,
tissue injuries and
certain cancer
therapies [81-88]
Central nervous
system repair [89]
Skin regeneration
[12]
Cell therapy [12]
Paneth’s cells, goblet cells, Cell therapy [12]
enteroendocrine cells
42
Table 1.3 Surface markers expressed by hAFSCs. Cited from [24]
Markers
Antigen
CD no.
ESC
SSEA-3
none
SSEA-4
none
Tra-1–60
none
Tra-1–81
none
SH2, SH3, SH4
CD73
Thy1
CD90
Endoglin
CD105
LCA
CD14
gp105–120
CD34
LPS-R
CD45
Prominin-1
CD133
Integrins
β1-integrin
CD29
Ig superfamily
PECAM-1
CD31
ICAM-1
CD54
VCAM-1
CD106
HCAM-1
CD44
I (HLA-ABC)
none
II (HLA-DR,DP,DQ)
none
Mesenchymal
Endothelial and
haematopoietic
MHC
43
Table 1.4 Ex vivo AFSC differentiation induced by chemical-based media (Edited from
[46]).
Germ layer
Tissue-specific cell type
Endoderm
Liver (hepatocytes)
Culture conditions
Hepatocyte growth factor (HGF), insulin,
oncostatin M, dexamethasone,
fibroblast growth factor 4 (FGF-4)
Mesoderm
Muscle (myocytes)
Pre-treatment with 5-azacytidine and horse
serum and chick embryo
extract on Matrigel® coated dish
Blood vessel (endothelial
cells)
Endothelial basal medium (EBM®) on gelatin
coated dish
Bone (osteoblasts)
Dexamethasone, β-glycerophosphate, ascorbic
acid-2-phosphate
Fat (adipocytes)
3-isobutyl-1-methyl-xanthine (IBMX),
insulin, indomethacin
Cartilage (chondrocytes)
Dexamethasone, ascorbic acid-2-phosphate,
sodium pyruvate, proline, transforming
growth factor β1 (TGF-β1)
Ectoderm
Nerve (neuronal cells)
Dimethyl sulfoxide (DMSO), butylated
hydroxyanisole (BHA), nerve growth
factor (NGF)
44
Table 1.5 Main characteristics of ESCs, iPS cells, AFSCs and MSCs (cited from [46]).
ESCs
iPS cells
AFSCs
MSCs
Source
Early stage
embryo
Somatic cells
Feeder cells
Markers
Required
SSEA3/4,
OCT-3/4,
SOX2
Pluripotent
Required
SSEA3/4,
OCT-3/4,
SOX2
Pluripotent
Yes
Yes
Bone marrow
and other
adult tissues
Not required
Not required
SSEA4, OCT4, CD44, CD73,
c-kit, CD44,
CD90, CD105
CD105
Broadly
Multipotent
multipotent
No
No
31-57
Long
Yes
No
48
Long
No
No
36
Long
No
No
Plasticity
Teratoma
formation
Doubling time (h)
Lifespan in vitro
Ethical issues
Clinical trials
45
Amniotic fluid
Variable
Short
No
Yes
Table 1.6 Various preclinical applications of AFSCs (edited from [46]).
Cell types
Scaffolds
Animal model and outcomes
Refs
Muscle
Rat AFSC
N/A
[49]
Nerve
Neuronallyinduced
hAFSC
Rat AFSC
N/A
Cyro-injured rat bladder walls,
prevention of cryo-injury induced
hypertrophy of smooth muscle cells
Twitcher mice, integration with
host neural cells
[50]
Kidney
hAFSC
N/A
Lung
hAFSC
N/A
Heart
Rat AFSC
N/A
N/A
Heart valve
hAFSC and
derived
cellular
structures
hAFSC
Extensive thoracic crush injury of
E2.5 chick embryo, reduction of
hemorrhage and increased survival
Mice with glycerol-induced
rhabdomyolysis and acute tubular
necrosis (ATN), amelioration of
ATN and decrease of damaged
tubules and apoptosis
Mice with hyperoxia and
naphthalene injury, plasticity of
AFS to respond to different lung
damage
Rat heart infarction by
ischemia/reperfusion, improvement
of ejection fraction
Heart infarction in immunesuppressed rats, improved cardiac
function
In vitro formation of neo-tissues by
conditioning in bioreactor system
[58]
Diaphragm
Ovine AFSC
[59]
Bone
hAFSC
Alginate/
collagen
Osteogenic
differentiation
of hAFSC
PLLA
nanofibers
Partial diaphragmatic replacement
of newborn lambs, mechanical and
functional outcomes
Subcutaneous implantation into
immunodeficient mice, ectopic
bone formation
Subcutaneous implantation into
athymic mice, ectopic bone
formation
N/A
Synthetic
polymeric
scaffold
Collagen
hydrogel
[40]
[51]
[52]
[53]
[5456]
[40]
[25]
Continued
46
Table 1.6 Continued.
Rabbit AFSC
PLLA
nanofibers
hAFSC
Porous
PCL
Cartilage
hAFSC
Angiogenesis
CM of
hAFSC
Pellet or
alginate
hydrogel
N/A
Full-thickness sternal defects,
postnatal reconstruction of chest
wall
Subcutaneous implantation into
athymic rats, ectopic bone
formation
In vitro cartilage formation
Hind-limb ischemia in mice, tissue
repair by host stem cell recruitment
mediated by stem cell-secreted
factors
PLLA: poly(L-lactic acid); PCL: poly(ε-caprolactone); CM: conditioned medium.
47
[61]
[60]
[44,
81]
[62]
Table 1.7 Different types of culture medium (Cited from [65]).
Medium
Characteristics
Serum-free media
Serum-free media do not require supplementation with
serum, but may contain discrete proteins or bulk protein
fractions (e.g., animal tissue or plant extracts) and are thus
regarded as chemically undefined (see: chemically defined
media).
Animal-derived
component-free media
Media containing no components of animal or human origin.
These media are not necessarily chemically defined (e.g.,
when they contain bacterial or yeast hydrolysates, or plant
extracts).
Chemically defined
media
Chemically defined media do not contain proteins,
hydrolysates or any other components of unknown
composition. Highly purified hormones or growth factors
added can be of either animal or plant origin, or are
supplemented as recombinant products (see: animal-derived
component-free media).
48
Figure 1.1 Differentiations of ESCs. Cited from [94]
49
Figure 1.2 Isolation and expansion of hAFSCs. Cited from [95]
50
Figure 1.3 Multi-lineage differentiation of AFSCs. AFSCs can differentiate into all three
germ layer cell types (picture adapted from [46]).
51
(A)
(B)
Figure 1.4 3-D fibrous PET for cell culture. (A)The structure formula of PET; (B) SEM
images of ESCs on PET scaffolds [12].
52
Figure 1.5 Media pyramid: a modular approach for the development of serum-free media.
Abbreviations: ADH, antidiuretic hormone; EGF, epidermalgrowth factor; FGF,
fibroblast growth factor; IGF-1, insulin-like growth factor 1; ITS, insulin–transferrin–
sodium selenite supplement; b-ME, b-mercaptoethanol; NGF, nervegrowth factor; PDGF,
platelet-derived growth factor; PGE2, prostaglandin E2; PTH, parathyroid hormone;
TGF-b, transforming growth factor-b; and VEGF, vascularendothelial growth factor.
Cited from [65]
53
Figure 1.6. Objectives and outline of this work.
54
Chapter 2
Expansion of Human Amniotic Fluid Stem Cells in 3Dimensional Fibrous Scaffolds in Bioreactors
Abstract
Human amniotic fluid stem cells (hAFSCs) are emerging as an important cell
source for tissue engineering and regenerative medicine due to their easy accessibility
and broad multi-potentiality. In clinical applications, a large number of hAFSCs are
required, which cannot be provided in conventional 2-dimensional (2-D) culture systems.
To address this issue, the expansion of human AFSCs in 3-dimensional (3-D)
polyethylene terephthalate (PET) scaffolds in a stirred bioreactor was evaluated. The
results showed that 3-D PET scaffold with in vivo-like environment and a large specific
surface area for cell adhesion promoted cell expansion (66-fold vs. 38-fold) compared to
2-D culture. A dynamic fibrous bed bioreactor (FBB) was used to expand AFSCs to
reach a high cell density of 3.2×106 cells/mL. The bioreactor-expanded cells maintained
clonogenic ability and high levels of expression (95.5-99.8%) of characteristic stem cell
surface makers, including CD29, CD44, CD90 and CD105. The differentiation of
bioreactor-expanded AFSCs into osteogenic and adipogeneic lineages was demonstrated
with Alizarin red S and Oil Red O staining, respectively, and further confirmed by
55
reverse transcriptase polymerase chain reaction (RT-PCR) analysis. This study
demonstrated the feasibility of using the FBB to mass-produce hAFSCs for potential
applications in tissue engineering and regenerative medicine.
2.1 Introduction
Mesenchymal stem cells (MSC) are non-hematopoietic cells that have been
identified primarily in bone marrow (BM). The clonogenic potential of these multipotent
BM-MSCs was first reported in the 1970s by Friedenstein et al [1]. The most important
characteristics of MSCs are their ability to differentiate into osteoblasts, chondrocytes,
and adipocytes [2]. In addition, MSCs were later reported to give rise to various cell
types, including neurons [3], hepatocytes [4], pancreatic islet cells [5, 6], cardiomyocytes,
and smooth muscle cells [7]. However, bone marrow aspiration is an invasive procedure
that causes pain and morbidity [8, 9]. Moreover, the frequency of MSCs in BM is low,
about 0.01% in healthy newborns and continues to decline to 0.001-0.0005% with age
[10]. Thus, MSCs from other easily obtainable sources with great expansion capabilities
have been highly sought for potential use in clinical therapies.
Recently, stem cells were identified in amniotic fluid (AF), which fills the
amniotic sac and surrounds the developing fetus [11]. AF contains a heterogeneous
population of cells shed from embryonic and extra-embryonic tissues during fetal
development. Previous studies have indicated that amniotic fluid stem cells (AFSCs)
possess a phenotype of MSCs [12] and can give rise to a wide range of cell types,
including adipogenic, osteogenic, myogenic, neurogenic, and hepatic lineages [13]. AF is
56
regarded as medical waste after amniocentesis and thus can be acquired easily without
ethical issues. Amniocentesis is a widely accepted procedure in prenatal testing and
presents a low risk for both the mother and the fetus [8, 14]. Additionally, AFSCs are not
tumorigenic and have extensive self-renewal capacity that can expand over 250
population doublings [13]. Therefore, AFSCs have emerged as a promising cell source
for tissue engineering and regenerative medicine.
Although there is much interest in cell-based therapies, the clinical use of AFSCs
has been limited partly because of the difficulty in obtaining the quantity of cells needed
for the applied dose, such as 1-5×106 MSCs per kg of patient body weight [15].
Conventionally, MSCs are expanded in petri dishes, T-flasks, and multiwell plates. These
techniques are labor-intensive, time-consuming, expensive, prone to contamination, and
difficult to scale up. Although cell factories have recently been applied to mass-produce
stem cells, they require a large working volume and only provide two-dimensional (2-D)
culture environments. While about 109 cells are needed for a 70 kg patient, the cell
density in 2-D culture is only about 105 cells/mL [16]. Therefore, an efficient and reliable
ex vivo expansion method of MSCs is required to achieve clinically relevant numbers. In
this study, a three-dimensional (3-D) polyethylene terephthalate (PET) fibrous bed
bioreactor (FBB) was developed for human AFSC expansion. PET is one of the first
synthetic polymers used in regenerative medicine and has been used for vascular
reconstructions [17] as well as bone cell cultures [18]. Compared with 2-D culture
systems, 3-D PET scaffolds can provide larger specific surface areas favoring stem cell
attachment, growth and higher cell density, and better mimic in vivo environments [1957
21]. Due to the porous structures, the PET fibrous matrices facilitate nutrient and oxygen
diffusion and can protect cells from shear damage [22, 23]. By providing a 3-D cell
growth environment, the PET-based bioreactor requires a small working volume to
generate a large quantity of human AFSCs.
The aims of this study were to a) investigate the proliferation of hAFSCs in 3-D
PET scaffold in comparison to 2-D cultures under static culture condition; b) expand
hAFSCs in PET-based 3-D dynamic FBB for potential clinical applications; and c)
characterize the bioreactor-expanded hAFSCs for phenotypic expression, clonogenic
ability, and multi-lineage differentiation potential.
2.2 Materials and Methods
2.2.1 AFSC cultures and media
AFSCs were cultured in 175 cm2 T-flasks and harvested by TrypLE™ Select
solution for 5 min at 37 °C. For 2-D static cultures to compare AFSC and BM-MSC, each
well of 48-well plates containing 1 mL growth medium was seeded with 100 cells and
incubated at 37 °C in a humidified 5% CO2 incubator. For the comparison of 2-D and 3D static cultures, each well of 48-well plates was seeded with 5000 cells. For 3-D
cultures, 5000 cells in 30 µl medium were carefully added to the center of the PET matrix
placed in each well of 48-well plate. The cells were kept at 37 °C in a humidified
atmosphere containing 5% CO2 for 3 h to allow cell attachment to PET scaffolds. Then,
the PET matrix was transferred to a new well on a 48-well plate, and 1 mL fresh medium
58
was added to each well. Unless otherwise noted, the culture medium was changed once
every 1-3 days, according to the metabolic activities of cells.
2.2.2 Preparation of PET fibrous scaffolds
Needle-punched PET fabric (thickness, 0.18 cm; fiber diameter, 20 µm; matrix
density, 0.11 g/cm3; pore size, 60-130 µm; porosity, 92.5%; specific surface area, 190
cm2/cm3) was used as cell culture scaffolds [25]. For static microwell cultures, diskshaped scaffolds (0.6 cm diameter) were used. For dynamic spinner-flask cultures, PET
matrix was cut into a 9.0 cm × 1.2 cm sheet. Before use, the fibrous scaffolds were
washed with phosphate-buffered saline (PBS) three times, sterilized in PBS at 121 °C, 15
psig for 30 min, and stored at room temperature. To favor cell attachment, PET scaffolds
were soaked in the culture medium for 30 min prior to seeding.
2.2.3 Static AFSC cultures in microwells
AFSCs were cultured in 175 cm2 T-flasks and harvested by TrypLE™ Select
solution for 5 min at 37 °C. For 2-D static cultures to compare AFSC and BM-MSC, each
well of 48-well plates containing 1 mL growth medium was seeded with 100 cells and
incubated at 37 °C in a humidified 5% CO2 incubator. For the comparison of 2-D and 3D static cultures, each well of 48-well plates was seeded with 5000 cells. For 3-D
cultures, 5000 cells in 30 µl medium were carefully added to the center of the PET matrix
placed in each well of 48-well plate. The cells were kept at 37 °C in a humidified
atmosphere containing 5% CO2 for 3 h to allow cell attachment to PET scaffolds. Then,
the PET matrix was transferred to a new well on a 48-well plate, and 1 mL fresh medium
59
was added to each well. Unless otherwise noted, the culture medium was changed once
every 1-3 days, according to the metabolic activities of cells.
2.2.4 Dynamic AFSC cultures in fibrous fed bioreactor (FBB)
The expansion of hAFSCs was studied in a fibrous bed bioreactor, which was
made of a 25-mL spinner flask with a PET matrix affixed on a stainless steel wire mesh
around the wall (Figure 2.1). The PET matrix (dimension: 1.2 cm × 9.0 cm × 0.18 cm)
had a total surface area of 4104 cm2. After sterilization, the FBB with the PET matrix was
soaked in 10 mL of the growth medium, inoculated with 106 hAFSCs (high seeding cell
number) or 105 hAFSCs (low seeding cell number), and incubated at 37 °C with agitation
at 60 rpm for 18-28 days in a humidified atmosphere containing 5% CO2. Glucose and
lactate concentrations in the medium were monitored daily, and the culture medium was
refreshed every 1-3 days according to the metabolic activities. On day 18 (high seeding
cell number) or day 28 (low seeding cell number), cells in the FBB were harvested by
using TrypLE™ Select and analyzed for morphology, surface marker expression, and
multi-lineage differentiation.
2.2.5 Flow cytometry
To identify the effects of bioreactor expansion on the immunophenotype of
hAFSCs, flow cytometric analysis of anti-CD29, anti-CD44, anti-CD105 (Developmental
Studies Hybridoma Bank, Iowa City, Iowa), anti-CD90, and anti-CD34 (BD, Franklin
Lakes, NJ) was performed. About 5×105 cells, after harvesting from the reactor and
dissociation, were used for flow cytometric analysis. Samples were fixed with 4%
60
paraformaldehyde (PFA) in PBS for 20 min. After washing with PBS three times, the
fixed cells were blocked in 3% FBS for 1 h and incubated overnight at 4 oC with the
primary antibodies in 1% FBS in PBS. Stained cells were washed and incubated with
Alexa Fluor® 488 secondary antibody (IgG1, or IgG2a for CD90) for 1 h. Positive cells
were detected and quantified against isotype control using FACS Calibur instrument and
CellQuest software (Becton Dickinson, Franklin Lakes, NJ).
2.2.6 Scanning electron microscopy (SEM)
The morphology and distribution of cells in the PET scaffolds were observed
using a scanning electron microscope. PET scaffolds containing cells were washed with
PBS and fixed with 2.5% (v/v) glutaraldehyde (Sigma-Aldrich, St. Louis, MO) overnight
at 4 oC. The fixed samples were rinsed with distilled water and progressively dehydrated
in ethanol solutions from 10% (v/v) to 100% (v/v) with 10% increment for 30 min at each
concentration. Then, the samples were dried using hexamethyldisilazane (HMDS)
(Sigma-Aldrich) and ethanol mixtures with ascending HMDS concentrations of 25%,
50%, and 75% (v/v) for final dehydration. The dried samples were sput-coated with
gold/palladium at an argon pressure of 14 Pa and a current of 15 mA for 140 seconds, and
then viewed under a Quanta 200 scanning electron microscope (FEI Worldwide,
Hillsboro, Oregon, USA) with 5-25 kV accelerating voltage.
2.2.7 Osteogenic and adipogenic differentiations
To test the differentiation potential, human AFSCs were induced for osteogenic
and adipogenic differentiation. The hAFSCs were seeded into six-well plates at 10,000
61
cells/cm2 and cultured until 70-80% confluence. Cells cultured in the growth medium
were used as negative control. For osteogenic differentiation, cells were cultured for 21
days in osteogenic medium composed of α-MEM supplemented with 16% FBS, 10 mM
β-glycerol phosphate, 1 nM dexamethasone, 50 µg/mL thyroxine (Sigma), 2 mM Lglutamine, 100 U/mL penicililin, and 100 µg/mL streptomycin. Media were changed
every 3 days. After 3 weeks, the cells were fixed with 10% (v/v) formalin and stained
with 1% (w/v) Alizarin red S solution. The presence of calcium was observed with a light
microscope (Olympus IX71, Olympus Corporation, Tokyo, Japan). For adipogenic
differentiation, confluent hAFSCs were cultured for 21 days in adipogenic medium
composed of Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10%
FBS, 1 µM dexamethasone, 1 mM 3-isobutyl-1-methylxanthine, 10 µg/mL insulin, 60
µM indomethacin, 2 mM L-glutamine, 100 U/mL penicililin, and 100 µg/mL
streptomycin. Media were changed every 3 days. After 21 days, the cells were stained
with 1% (w/v) Oil Red O solution and the intracellular lipid vacuoles were visualized
with a light microscope (Olympus IX71).
2.2.8 Reverse transcriptase polymerase chain reaction (RT-PCR)
The total RNA was isolated using TRIZOL reagent (Invitrogen, Carlsbad, CA)
from bioreactor expanded hAFSCs after osteogenic and adipogenic differentiations for 21
days. RNA concentrations were measured using a ND-1000 spectrophotometer
(NanoDrop Technologies, Wilmington, DE). After that, 1 µg of RNA was initially
reverse transcribed into cDNA using SuperScriptTM III First-Strand Synthesis System
(Invitrogen). Then, the cDNA (~200 ng) was used as a template for the amplification of
62
the target genes using the Quick-Load® Taq 2X Master Mix Kit (BioLabs, Ipswich, MA)
and the primer sequences listed in Table 2.1. Two osteogenic genes encoding runt related
transcription factor 2 (RUNX2) and osteopontin (OPN), and three adipogenic genes
encoding adipose fatty acid-binding protein (aP2), peroxisome proliferative activated
receptor γ (PPAR-γ) and lipoprotein lipase (LPL), respectively, were analyzed. The
housekeeping gene for glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used
as an endogenous reference gene. Amplified products were fractionated in a 2% agarose
(Fisher Scientific, Pittsburgh, PA) gel at 70 V for 80 min and visualized and
photographed with a Gel Doc 2000 Gel Documentation System (Bio-Rad, Hercules, CA).
2.2.9 Colony-forming unit-fibroblastic (CFU-F) assay
CFU-F assay was used to examine the progenitor content of bioreactor-expanded
human AFSCs. Cells before (control) and after bioreactor expansion were seeded into 15cm cell culture dishes at a density of 1000 cells/dish. The cells were cultured at 37 °C in a
humidified atmosphere containing 5% CO2 for 21 days. The cells were then washed with
PBS and incubated with a 3% crystal violet (Sigma-Aldrich) solution in methanol for 10
min. Stained colonies were rinsed with distilled water and counted manually. Three
samples from each condition were used for each data point, and two independent runs
were performed.
2.2.10 Analytical methods
Cell proliferation was measured using the Alamar Blue assay (AbD Serotec,
Raleigh, NC). The cells were incubated with 500 µl of 10% Alamar Blue solution at
63
37 °C for 3 h. The fluorescence of resorufin was monitored in triplicate in a 96-well plate
(BD Optilux™, Black/clear bottom) at 535 nm excitation wavelength and 590 nm
emission wavelength using a GENios Pro plate reader (Tecan, Research Triangle Park,
NC). The fluorescence intensity, which was linearly correlated to the cell number, was
used to calculate the expansion fold and the specific growth rate. The specific growth rate
was determined using a simple first order kinetic model as: dX/dt = μapp·X, where X is
the cell number, and μapp and t are the apparent growth rate and culture time, respectively.
The concentrations of glucose and lactate were measured using a biochemistry analyzer
YSI 2700 (YSI Life Sciences, Yellow Springs, OH).
2.2.11 Statistical analysis
Unless otherwise noted, all experiments and samples were triplicated.
Experimental results were presented as mean ± standard deviation (SD) (n = 3) and
analyzed by Student’s t-test using JMP 7.0 (SAS Institute Inc., Cary, NC) with p < 0.05
as statistically significant.
2.3 Results and Discussion
2.3.1 AFSCs proliferation in 3-D PET scaffolds
The in vitro proliferation potential of AFSCs was first compared with BM-MSCs
in 2-D static cultures (Figure 2.2A). Compared to BM-MSC, AFSC proliferated more
quickly with a shorter lag phase and grew significantly faster in the exponential phase.
The expansion fold of AFSCs over 18 days was 31-fold while it was only 15.8-fold for
BM-MSC. On average, AFSCs had a specific growth rate of 0.46 day-1 and doubling time
64
of 37 h, while BM-MSCs had a specific growth rate of 0.26 day-1 and doubling time of 63
h. Clearly, AFSCs representing an intermediate state between embryonic and adult
mesenchymal stem cells [26] exhibited an extensive proliferation capacity and would be
advantageous over BM-MSCs as a promising cell source for tissue engineering and
regenerative medicine applications.
The proliferation of AFSCs in 3-D fibrous scaffolds was then investigated and
comapted to 2-D static cultures (Figure 2.2B). During the first six days, AFSCs on 2-D
surface grew faster than those in 3-D PET scafflods. The 2-D culture reached its
maximum density with a 37.6-fold expansion on day 6, but then rapidly decreased
because of cell death and detachment from the surface due to contact inhibition upon
confluence. However, in 3-D culture, AFSCs continued to proliferate and expanded 65.9fold by day 12. The specific growth rate in 2-D culture was 0.53 day-1 with a doubling
time of 32 h, while 3-D culture had a lower specific growth rate of 0.43 day-1 and a
longer doubling time of 39 h, which however were within the variation of 2-D cultures.
The 3-D scaffold with a large specific surface area allowed cells to grow to a higher cell
density, and thus is more desirable for the large-scale expansion of AFSCs.
2.3.2 AFSC expansion in the fibrous bed bioreactor
Given the high proliferation of AFSCs in 3-D fibrous scaffolds in static culture,
AFSCs were then expanded in a dynamic FBB for 18 days. Figure 2.3 shows the
metabolic activities of AFSCs seeded at 1×106 cells (high seeding cell number) or 1×105
cells (low seeding cell number) in the FBB with periodically medium refreshing. Initially,
65
the glucose consumption and lactate production were low due to the low seeding cell
density, especially for low seeding cell number. As cells grew to a higher cell density,
both glucose consumption and lactate generation rates also increased in each subsequent
feeding and more frequent medium refreshing was necessary to maintain the culture. For
the culture with a high seeding number, on day 11, 15 mL growth medium were added
instead of 10 mL, which could not provide sufficient substrate to keep up with the
consumption. On day 12, 20 mL growth medium were added so cell growth would not be
limited by glucose or inhibited by lactate. During the expansion period, the lactate
concentration was kept well below the reported inhibitory level of 3.2 g/L for human
MSC [27]. By the end of day 18, the total expansion fold was 31.7, reaching a total cell
number of 3.17×107 AFSCs and final density of 3.17×106 cells/mL (Table 2.2). For the
bioreactor initially seeded with 1×105 cells, it took 28 days to reach the final density of
1.55×106 cells/mL or a total cell number of 1.55×107 AFSCs, corresponding to an
expansion fold of 155.
Glucose is an important energy source in cellular metabolism. Mammalian cells
utilize glucose as the primary source to produce ATP, either through mitochondrial
oxidative phosphorylation, which yields around 3038 moles ATP per mole glucose, or
anaerobic glycolysis, which yields 2 moles ATP and 2 moles lactate per mole glucose
[28]. A lactate yield of higher than 2 mol/mol indicates that lactate is being generated
from other sources such as glutamine [29]. During the exponential growth phase of
AFSCs in the 3-D bioreactor, the apparent yield of lactate/glucose varied between 1.66
and 1.83, with an average of 1.73, which was significantly lower than that in T-flasks
66
(2.47), indicating that the energy metabolism of AFSCs in the FBB was more efficient
than that in conventional 2-D cultures. Moreover, our bioreactor also supported more
efficient metabolism of glucose than a reported fixed bed bioreactor for BM-MSC
expansion in which the apparent yield of lactate from glucose was 2.74 mol/mol [30]. In
our PET-based 3-D dynamic bioreactor, oxygen and nutrient transfers were improved by
agitation and by using a high-porous scaffold. Providing sufficient oxygen and nutrients
facilitated cells to undergo more efficient metabolism. In addition, daily and average
lactate yields of 1.71.9 in the FBB culture indicated that AFSC metabolism was mainly
through anaerobic glycolysis combined with oxidative phosphorylation.
Cell morphology in the PET scaffold was observed at the end of 18-day culture
using SEM (Figure 2.4). The 3-D PET scaffold consisting of non-woven, randomly
oriented fibers with a high aspect ratio, provided high surface area and sufficient void
volume for cell growth. The morphology and distribution of AFSCs in the PET scaffold
were multi-dimensional (Fig. 2.4A and 4C). Most cells attached on fibers and bridged
between fibers, while some cells formed clusters within the fibrous matrix (Fig. 2.4B and
4D). Figure 2.4B also shows that AFSCs adopted a variety of morphologies within the
scaffold and secreted an extensive extracellular matrix (ECM) network, which might
include collagen I, collagen IV, laminin, and fibronectin as reported for BM-MSCs
cultured in a 3-D fibrous matrix [21, 31].
Biomaterial scaffolds were found to play important roles in directing cellular
behavior and function in tissue engineering and regenerative medicine [32]. AFSCs are
anchorage-dependent cells; viable and healthy cells were found to attach to the fibrous
67
matrix due to the interactions between the cells and the scaffold. The ability to retain
viable and healthy cells in the fibrous matrix contributed to the high cell density during
long-term culture in the FBB. Additionally, the highly porous network of PET scaffold
permitted diffusion of oxygen, nutrients, growth factors, as well as metabolites, and
protected the cells from shear stress, all of which contributed to the prolonged growth
phase of AFSCs in the 3-D scaffold [33]. Large-scale expansion of MSCs in a hollowfiber membrane bioreactor [34] and a rotary bioreactor [35] has also been reported.
However, hollow-fiber membrane and rotary bioreactor systems are expensive and
difficult to scale up. In addition, the operation life of hollow-fiber bioreactor is relatively
short because of the accumulation of dead cells over time. Therefore, PET-based FBB is
advantageous for AFSC long-term expansion.
2.3.3 Phenotype of bioreactor-expanded human AFSCs
To investigate the influence of 3-D expansion in the FBB on AFSC phenotype,
cells after expansion in the bioreactor were examined for their surface markers with flow
cytometry analyses and compared to the cells before expansion (control) (Figure 2.5). In
general, human MSCs express CD29 (β-integrins), CD44 (hyaluronan receptor), CD90
and CD105 (endoglin), but do not express hematopoietic lineage markers, such as CD34
(hematopoietic progenitors receptor) [13, 36]. As expected, AFSCs from the T-flask
cultures (control) had high expression levels of CD29 (99.1%), CD44 (95.5%), CD90
(99.8%), and CD105 (98.7%) and did not express CD34 (0.1%). AFSCs expanded in the
FBB also expressed high levels of CD29 (98.3%), CD44 (97.6%), CD90 (99.6%) and
CD105 (99.4%), and negative in CD34 (0.4%). So AFSCs expanded in the FBB did not
68
change their surface marker expression. In contrast, decreased expression of surface
marker including CD29, CD44, CD90, and CD105 has been observed for MSC
aggregates expanded in stirred bioreactors and rotating wall vessels [37]. The presence of
shear stress in a dynamic bioreactor has been shown to mediate several signaling proteins
including mitogen activated protein kinases (MAPK) and Wnt [38]. Apparently, the
protection from 3-D fibrous scaffold minimized the effect of shear stress on AFSCs in
our study.
The progenitor content of FBB-expanded AFSCs was further confirmed by
examining their clonogenic capacity using CFU-F assay (Table 2.3 and Figure 2.6). The
results showed that the total number of colonies were similar for both the control (87±7)
and cells harvested from the FBB (79±8), although fewer larger colonies with 50 or more
cells (51±4 vs. 76±5) and more smaller colonies with 25 to 50 cells (28±4 vs. 11±3) were
formed by cells harvested from the FBB as compared to the control. This slight decline in
the total colony number and larger colonies of 50 or more cells might be attributed to the
reduced cell viability due to trypsin treatment for cell harvesting from 3-D scaffolds,
which also happened during normal passaging. Similar phenomena have also been
reported by other researchers [29, 39].
2.3.4 Multi-lineage differentiation of FBB-expanded human AFSCs
To confirm the multipotency of the FBB-expanded AFSCs, cells harvested from
the 3-D bioreactor were induced to differentiate along osteogenic and adipogenic lineages,
respectively, under two different conditions. As expected, AFSCs cultured in the growth
69
medium (negative control) did not produce any mineralized matrix after Alizarin red
staining nor showing any oil droplets with Oil Red O staining (Figure 2.7A). On the
other hand, the AFSCs in the osteogenic medium developed a calcium-rich mineralized
bone matrix along the cell membrane as large red aggregates embedded in the ECM
(Figure 2.7B), while the AFSCs cultured in the adipogenic medium exhibited
morphological changes and small lipid vesicles in the cytoplasm were revealed by Oil
Red O staining (Figure 2.7C). RT-PCR analysis exhibited high expressions of RUNX2
and OPN in cells after osteogenic induction (Figure 2.8A) and high expressions of
PPAR-γ, LPL and aP2 in cells after adipogenic induction (Figure 2.8B). These results
confirmed that the FBB-expanded AFSCs were capable of differentiating into osteogenic
and adipogenic lineages, suggesting that AFSCs expansion in the FBB maintained their
multipotency after an extended culturing period of 1828 days.
2.4 Conclusions
Because of the easy and safe accessibility, abundant cell numbers, and lack of
ethical concerns, human AFSCs have emerged as an attractive source of stem cells for
basic research and clinical applications. AFSCs with a superior proliferation capacity to
BM-MSCs are more promising for use in regenerative medicine where rapid proliferating
progenitor cells are required. Compared to 2-D cultures, AFSCs grown in 3-D
microenvironments of PET had stable long-term proliferation with a significantly higher
expansion fold, suggesting that the PET fibrous matrix is an effective 3-D support for
anchorage-dependent AFSCs. Furthermore, a PET-based 3-D dynamic fibrous-bed
bioreactor was applied to expand AFSCs. The dynamic bioreactor enhanced nutrient,
70
oxygen and metabolite transfers and consequently promoted cell expansion to reach a
high cell number required by clinical dose. Additionally, our results indicated that the
bioreactor-expanded AFSCs maintained the profile of surface markers, clonogenic ability,
and multi-lineage differentiation potential. In conclusion, the PET-based 3-D dynamic
fibrous-bed bioreactor system can be easily implemented for clinical-scale expansion to
maximize AFSC yield while maintaining cell product quality suitable for regenerative
medicine and cell therapy.
Acknowledgments
This work was supported in part by Alumni Grants for Graduate Research and
Scholarship (AGGRS) of The Ohio State University. We would like to acknowledge Dr.
Anthony Atala and Dr. James Yoo of the Wake Forest Institute for Regenerative
Medicine (Winston-Salem, NC) for kindly providing hAFSCs used in this study.
71
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Table 2.1. Primers for RT-PCR analysis of osteogenic and adipogenis differentiations.
Gene
Sequence (forward/reverse)
RUNX2
AGTGGACGAGGCAAGAGTTTC
CCTTCTGGGTTCCCGAGGT
GAGACCCTTCCAAGTAAGTCCA
GATGTCCTCGTCTGTAGCATCA
AAGAAGTAGGAGTGGGCTTTGC
CCACCACCAGTTTATCATCCTC
TTGGTGACTTTATGGAGCCC
CATGTCTGTCTCCGTCTTCTTG
AGAGAGGACTTGGAGATGTGGA
GGAAGACTTTGTAGGGCATCTG
GTGGTCTCCTCTGACTTCAACA
CTCTTCCTCTTGTGCTCTTGCT
OPN
aP2
PPAR-γ
LPL
GAPDH
Product
size (bp)
Annealing
Temp (°C)
Cycle
Gene ID
117
62
35
NM_004348
354
62
35
285
62
35
NM_0010400
60
NM_001442
311
62
35
NM_005037
264
62
35
NM_000237
211
62
24
NM_002046
*Osteogenic genes: related transcription factor 2 (RUNX2), osteopontin (OPN);
Adipogenic genes: adipose fatty acid-binding protein (aP2), peroxisome proliferative
activated receptor γ (PPAR-γ) and lipoprotein lipase (LPL). Glyceraldehyde-3-phosphate
dehydrogenase (GAPDH) was used as an endogenous reference gene.
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Table 2.2. Cell number and expansion fold of human AFSCs in 3-D bioreactor.
Parameters
Low Seeding Density
High Seeding Density
Initial seeding cell number (106 cells)
0.1
1.0
Final harvested cell number (106 cells)
15.5
31.7
28
18
Final cell density (106 cells/mL)
1.55
3.17
Expansion fold
155
31.7
Culture duration (days)
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Table 2.3. CFU-F assay for human AFSCs before and after expansion in 3-D bioreactor.
Condition
Colonies with ≥50 cellsb Colonies with 25-50 cellsb
Total coloniesa
Before expansion
76 ±5
11 ±3
87 ±7
After expansion
51 ±4
28 ±4
79 ±8
a
No significant difference in total colony number was observed before and after expansion in 3-D
bioreactor (p > 0.05). bMore large colonies were observed before expansion, while more small
colonies were observed after expansion (p < 0.05).
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Figure 2.1 A fibrous bed bioreactor modified from a spinner flask with a PET
matrix around the wall. The fibrous PET matrix was fixed on a stainless steel wire
mesh before placed in the spinner flask.
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Figure 2.2. Human AFSCs expansion in 3-D PET matrix. (A) Growth kinetics of 2-D
static cultures of AFSCs and BM-MSCs; (B) Growth kinetics of 2-D and 3-D static
cultures of AFSCs. * indicate p <0.05.
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Figure 2.3. Metabolic activities of human AFSCs expanded in 3-D bioreactor. The
growth medium was periodically refreshed (indicated by the sudden changes in glucose
and lactate concentrations). (A) Glucose and lactate concentrations; (B) Glucose
consumption rate, lactate production rate, and lactate yield for the reactor with the high
seeding cell number of 1×106; (C) Glucose and lactate concentrations; (D) Glucose
consumption rate, lactate production rate, and lactate yield for the reactor with the low
seeding cell number of 1×105.
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Figure 2.4. SEM images of human AFSCs expanded in 3-D PET fibrous matrices.
(A) AFSCs uniformly distributed in 3-D scaffolds; Low magnification, scale bar: 500 μm;
(B) AFSC clusters with extracellular matrix fibers; High magnification, scale bar: 100 μm.
(C) AFSC cell sheet was formed in 3-D scaffolds; Low magnification, scale bar: 500 μm;
(D) AFSC cell sheet at high magnification, scale bar: 200 μm.
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Figure 2.5. Phenotype of human AFSCs before and after expansion in 3-D
bioreactor. The surface markers including CD29, CD44, CD90, CD105 and CD34 were
analyzed by flow cytometry against isotype control. The histograms were compared for
the samples before and after expansion in bioreactor for 18 days.
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Before expansion
After expansion
Figure 2.6. CFU-F assay of hAFSCs before and after FBB expansion.
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Figure 2.7. Multi-lineage differentiation of human AFSCs after expansion in 3-D
bioreactor. (A) AFSC control before induction; (B) Alizarin Red S-staining for
osteogenic differentiation; (C) Oil Red O staining for adipogenic differentiation.
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Figure 2.8. RT-PCR analysis of differentiated human AFSCs after expansion in 3-D
bioreactor. (A) Osteogenic differentiation; osteogenic genes including runt related
transcription factor 2 (RUNX2) and osteopontin (OPN) were detected; (B) Adipogenic
differentiation; adipogenic genes including adipose fatty acid-binding protein (aP2),
peroxisome proliferative activated receptor γ (PPAR-γ), and lipoprotein lipase (LPL)
were detected. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as an
endogenous reference gene and shown as weak bands.
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Chapter 3
Effects of naringin on the proliferation and osteogenic
differentiation of human amniotic fluid derived stem cells
Abstract:
Human amniotic fluid derived stem cells (hAFSCs) are a novel cell source for
generating osteogenic cells to treat bone diseases. Effective induction of osteogenic
differentiation from hAFSCs is critical to fulfill their therapeutic potential. In this study,
naringin, the main active compound of rhizome drynariae (a Chinese herbal medicine),
was used to stimulate the proliferation and osteogenic differentiation of hAFSCs. The
results showed that naringin enhanced the proliferation and alkaline phosphatase activity
(ALP) of hAFSCs in a dose-dependent manner in the range of 1-100 μg/mL, while
inhibition effect was observed at 200 μg/mL. Consistently, the calcium content also
increased with increasing naringin concentration up to 100 μg/mL. The enhanced
osteogenic differentiation of hAFSCs by naringin was further confirmed by the dosedependent up-regulation of marker genes including osteopontin (OPN) and Collagen I
from RT-PCR analysis. The increased osteoprotegerin (OPG) expression and minimal
expression of receptor activator of nuclear factor kappa-B ligand (RANKL) suggested
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that naringin also inhibited osteoclastogenesis of hAFSCs. In addition, the gene
expressions of bone morphogenetic protein 4 (BMP4), runt related transcription factor 2
(RUNX2), β-catenin, and Cyclin D1 also increased significantly, indicating that naringin
promotes the osteogenesis of hAFSCs via the BMP and Wnt/β-catenin signaling
pathways. These results suggested that naringin can be used to upregulate the osteogenic
differentiation of hAFSCs, which could provide an attractive and promising treatment for
bone disorders.
3.1 Introduction
Bone diseases, especially osteoporosis, bring serious issues to public health.
Osteoporosis is characterized by low bone mineral density and microarchitecture
deterioration, resulting in structural instability of bone tissue and a high fracture risk.
Estrogen withdrawal is the most well-recognized cause of osteoporosis [1], which
happens more commonly in the senior society and results in excess morbidity, mortality,
and the decreased quality of life. Recently, it is estimated that more than 200 million
people in the world [2] and 44 million in the US [3] suffer from osteoporosis. In America,
more than 1.5 million fractures associated with osteoporosis occur each year [4]. National
costs on the medical care expenses related to the bone fractures was more than $17
billion in 2005, and a cumulative cost of $474 billion is estimated for the next two
decades [5]. Estrogen replacement therapy (ERT) has been considered to be the most
effective treatment for osteoporosis in the past 10 years. However, a long-term use of
estrogen could increase the risk of breast cancer, endometrial carcinoma, and
cardiovascular diseases [1, 6]. Bisphosphonate therapy is another method developed in
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the last decade. Nevertheless, it only inhibits the resorption of osteoclast and can cause
acute incapacitating bone, joint, and muscle pain [7, 8].
Amniotic fluid derived stem cells (AFSCs) are a novel cell source for tissue
engineering and regenerative medicine because they have high potential to differentiate
into osteoblasts, chondrocytes, and adipocytes [9] and possess a phenotype of
mesenchymal stem cells (MSCs) [10]. MSCs have promising capacities to heal bone
fractures and thus obtained much attention in treating bone diseases [11]. Animal studies
showed that MSCs could possibly involve in bone formation through intravenous
infusion to target bones [12]. Compared to bone marrow-derived MSCs, AFSCs have
fetal origin and an extensive self-renewal capacity [9, 13]. In addition, AFSCs are not
tumorigenic and have no ethical concerns for clinical use. Therefore, AFSCs are superior
candidates for cell-based therapies especially for the treatment of bone disorders.
Rhizoma drynariae is a traditional Chinese herbal medicine, which has been
commonly used to treat orthopedic disorders and bone healing for thousands of years [14].
Modern pharmacological study indicates that naringin, a polymethoxylated flavonoid
(Figure 3.1), is the main active compound of rhizome drynariae. Naringin has been
found to enhance the bone morphogenetic protein (BMP) level of osteoblasts [15] and
stimulate the proliferation and osteogenic differentiation of bone marrow-derived MSCs
[16]. However, osteoblast is only in the downstream of the osteogenesis tree and no
underlying signal transduction pathways have been investigated for the effect of naringin
on MSCs. Thus, for the first time, the effects of naringin on the proliferation and
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osteogenic differentiation of hAFSCs and the responsible signaling pathways were
elucidated and are reported in this study.
3.2 Materials and Methods
3.2.1 Culture of human amniotic fluid derived stem cells (hAFSCs)
The hAFSCs were isolated and cultured as previously described [9]. hAFSCs at
passages 16-18 were used in this study. All culture reagents were purchased from Life
Technologies unless otherwise noted. The cells were cultured in alpha-minimum essential
medium (α-MEM) supplemented with 15% embryonic stem cell qualified-fetal bovine
serum (ES-FBS), 100 U/mL penicillin, 100 µg/mL streptomycin, 2 mM L-glutamine, 18%
Chang B, and 2% Chang C (Irvine Scientific, Santa Ana, CA). The hAFSCs were
maintained at 37 °C in a humidified 5% CO2 incubator and sub-cultured at 70%
confluence. The culture medium was changed every 3 days.
3.2.2 hAFSC treatment with naringin
Naringin (≥ 90% purity) was purchased from Sigma-Aldrich (St. Louis, MO). The
hAFSCs (1×104 cells/ml) were seeded in 48, 24, and 6-well plates and cultured in the
growth medium until 70-80% confluence. Then, the cells were cultured in the
differentiation medium, which contained α-MEM, 17% FBS (Atlanta Biologicals, Atlanta,
GA), 2 mM L-glutamine, 100 U/mL penicillin, and 100 µg/mL streptomycin. Various
amounts of naringin were added in the differentiation medium to final concentrations of 1,
10, 100, and 200 µg/ml, respectively. Cells cultured in the medium without naringin were
used as negative control. Media were changed every 3 days.
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3.2.3 Cell proliferation analysis
The hAFSCs (5×103 per well) were seeded in a 48-well plate. After 24 h of
incubation, the growth medium was changed into naringin-containing media at a
concentration of 0 (Control), 1, 10, 100, and 200 µg/ml accordingly. Cells were incubated
at 37 °C in a humidified 5% CO2 incubator for 1, 2, 3 or 4 days. After that, the medium
was replaced with 500 µl of 10% Alamar Blue (AbD Serotec, Raleigh, NC) solution at
37 °C for 3 h. The fluorescence of the medium was then monitored in triplicate at 535 nm
excitation wavelength and 590 nm emission wavelength using a GENios Pro plate reader
(Tecan, Research Triangle Park, NC). The fluorescence intensity can be correlated to the
cell number, using a standard calibration curve.
3.2.4 Alkaline phosphatase activity (ALP) assay
Osteogenesis of hAFSCs was induced in the differentiation medium containing
naringin. At day 7, cells were washed with PBS twice and lysed with the lysis buffer
consisting of 20 mM Tris-HCl (pH 7.5), 150 mM NaCl, and 1% Triton X-100 for 5 min.
The chromogenic substrate for ALP was p-nitrophenyl phosphate (pNPP; Sigma-Aldrich).
A 50 µL of lysed sample was mixed with 50 µL pNPP (1 mg/ml) substrate solution
containing 1.0 mg/mL pNPP, 0.2 M Tris buffer and 5 mM MgCl2 at 37 °C for 15 min on
a Belly Button Shaker (MidSci, St. Louis, MO). The reaction was stopped by adding 25
µL of 3 N NaOH. Absorbance of p-nitrophenol released in the samples was measured at
405 nm using a SpectraMAX 250 microplate reader (Molecular Devices, Sunnyvale, CA).
The protein concentration of cell lysate was determined using the Bradford assay at 595
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nm on a microplate spectrophotometer (Bio-Rad, USA). The ALP activity was
normalized according to the total protein content of cell lysate and expressed as nmol (pnitrophenyl)/min/mg protein.
3.2.5 Alizarin red S (ARS) staining
ARS staining was performed to evaluate the calcium deposition in cells of the
osteogenic lineage obtained from hAFSCs after 28 days of treatment with naringin.
Briefly, cells cultured in 24-well plate were rinsed with PBS twice, fixed with 10% (v/v)
formalin, and then stained with 1% (w/v) ARS solution. Orange red staining indicated the
location and intensity of the calcium deposition. The presence of calcium was observed
with a light microscope Olympus IX71 (Olympus Corporation, Tokyo, Japan).
3.2.6 Calcium assay
To quantify mineralization, the calcium deposition in hAFSCs after 21 days was
measured using the Calcium Assay (Genzyme Diagnostics, Charlottetown, PE, Canada).
Samples were added with 1 M acetic acid and placed on a vortex overnight at 4 °C to
extract the calcium from the mineralized matrix. In a 96 well clear polycarbonate plate,
15 µL of cell extract was mixed with 150 µL of the Calcium Assay reagent and incubated
for 30 s at room temperature. The absorbance at 650 nm was determined on a
SpectraMAX 250 microplate reader. The samples were measured in triplicate and
compared to the calcium calibration curve.
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3.2.7 Reverse transcriptase polymerase chain reaction (RT-PCR)
Total RNA was isolated from hAFSCs treated with different concentrations of
naringin using TRIZOL reagent (Invitrogen, Carlsbad, CA). RNA concentrations were
measured using a ND-1000 spectrophotometer (NanoDrop Technologies, Wilmington,
DE). After that, 1 µg of RNA was initially reverse transcribed into cDNA using
SuperScript™ III First-Strand Synthesis System (Invitrogen). Then, 200 ng of the cDNA
was used as a template for the amplification of the target genes using the Quick-Load®
Taq 2X Master Mix Kit (BioLabs, Ipswich, MA). The primer sequences of the analyzed
genes and PCR conditions are listed in Table 3.1. For osteogenic differentiation, genes of
osteopontin (OPN), collagen I, and ALP were measured. For bone morphogenetic protein
(BMP) pathway, genes of runt related transcription factor 2 (RUNX2) and BMP4 were
measured. For Wnt pathway, β-catenin and Cyclin D1 were analyzed. For osteoclast
differentiation, osteoprotegerin (OPG) and receptor activator of nuclear factor kappa-B
ligand (RANKL) were measured. The housekeeping gene, glyceraldehyde-3-phosphate
dehydrogenase (GAPDH), was used as an endogenous reference gene. Amplified
products were fractionated in a 2% agarose (Fisher Scientific, Pittsburgh, PA) gel at 70 V
for 80 min and visualized and photographed with a Gel Doc 2000 Gel Documentation
System (Bio-Rad, Hercules, CA). The expression level of each gene was analyzed using
Image J Software and normalized to the GAPDH expression.
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3.2.8 Statistical analysis
Unless otherwise noted, all experiments and samples were triplicated. Experimental
results were presented as mean ±standard deviation (SD) (n = 3) and analyzed using
ANOVA followed by paired Tukey-Kramer analysis using JMP 7.0 (SAS Institute Inc.,
Cary, NC). p < 0.05 was considered as statistically significant.
3.3 Results
3.3.1 Effects of naringin on the proliferation of hAFSCs
The stimulation effect of naringin on the proliferation of hAFSCs during a 4-day
culture was evaluated at various concentrations (1-200 µg/mL) (Figure 3.2). In the
presence of naringin, the proliferation fold of hAFSCs increased in a dose-dependent
manner in the range of 1-100 µg/mL. For example, on day 3, naringin increased the
proliferation from 19-fold in the control to 20, 23, 26-fold at 1, 10, and 100 µg/mL,
respectively. The use of 100 µg/mL naringin, the most effective concentration, increased
the proliferation by 28%, 32%, 35% and 19% compared to the control on day 1, 2, 3, and
4, respectively. However, 200 µg/mL naringin slightly inhibited the proliferation of
hAFSCs by 2-5% compared to control, indicating that high concentration (≥200 µg/mL)
of naringin may be harmful to cell growth. Thus, naringin within a range of 0-100 µg/mL
had no cytotoxic effect and stimulated the proliferation of hAFSCs.
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3.3.2 Effect of naringin on the ALP activity of hAFSCs
ALP activity was used to indicate the early osteogenic differentiation of hAFSCs.
Naringin was shown to increase the ALP activity of hAFSCs in a dose-dependent manner
in the range of 1-100 µg/mL after a 7-day culture (Figure 3.3). Compared to the control,
ALP activity increased 44%, 57% and 163% in the presence of 1 µg/mL, 10 µg/mL, and
100 µg/mL naringin, respectively. The ALP activity of hAFSCs treated by 200 µg/mL
naringin also increased 74%, but was lower than that treated with 100 µg/mL naringin.
3.3.3 Effect of naringin on calcium deposition
The ARS staining after 28 days of naringin treatment was performed to detect the
presence of calcium. In the presence of naringin, more calcium deposition was observed
compared to the control group (Figure 3.4A). At the concentration of 1-100 µg/mL
naringin, calcium deposition increased in a dose-dependent manner, while cells treated
with 200 µg/mL naringin (N200) produced less calcium than the 100 µg/mL naringin
group (N100). The osteogenic differentiation of the cells was further investigated by
quantifying the calcium content (Figure 3.4B). Consistently, naringin increased calcium
deposition in a dose-dependent manner at concentrations of 1-100 µg/mL. Compared to
the control group, the calcium content increased 31%, 44% and 239% in the presence of
1 µg/mL, 10 µg/mL and 100 µg/mL naringin, respectively. The calcium content in the
N200 group was much lower than that of N100 group, only increased 15% compared to
control. Hence, both calcium quantification and ARS staining results confirmed that
naringin promoted calcium deposition in hAFSCs.
97
3.3.4 Effect of naringin on the expression of osteogenic markers
The RT-PCR results for osteogenic differentiation of hAFSCs showed that the
osteogenic marker genes, including OPN and Collagen I, were significantly up-regulated
with naringin treatment compared to the control group at day 21 (Figure 3.5A, 3.5B).
ALP expression was weak for all the samples except for the negative expression in N200
group. In general, naringin increased the expression of osteogenic differentiation markers
in a dose-dependent manner at concentrations of 1-100 µg/mL, while at 200 µg/mL, the
expression levels were reduced.
3.3.5 Effect of naringin on the osteoclast differentiation of hAFSCs
The gene expression of OPG was also markedly increased by naringin in a dosedependent manner at concentrations of 1-100 µg/mL (Figure 3.5C, 3.5D). Meanwhile,
little expression of RANKL was observed in the presence of naringin. It should be noted
that RANKL expression was observed in the presence of curculigoside, an active
component in another Chinese herbal medicine (unpublished data). Hence, the minimal
RANKL expression was not due to the detection method. The ratio between mRNA
expressions of OPG to RANKL (OPG/RANKL) is usually used as an indicator of
osteoclastogenesis. The dose-dependent expression of OPG and minimal expression of
RANKL indicated that naringin inhibited osteoclastogenesis by increasing the relative
portion of OPG expression and thus enhancing the osteogenic differentiation of hAFSCs.
98
3.3.6 Effect of naringin on BMP and Wnt/β-catenin pathways
BMP and Wnt signaling pathways were reported to involve in osteogenic
differentiation of MSCs [17]. Therefore, effects of different concentrations of naringin on
the BMP and Wnt signaling pathways in hAFSCs were studied on day 21 (Figure 3.6).
As can be seen in Figure 3.6A and 3.6B, two BMP-related regulators, BMP4 and
RUNX2, were up-regulated in naringin groups in a dose-dependent manner from 1
μg/mL to 100 μg/mL, while the expression was reduced at 200 μg/mL. In addition, the
gene expression levels of two Wnt-related regulators, β-catenin and its target gene Cyclin
D1 were evaluated (Figure 3.6C, 3.6D). Both of these two regulators were significantly
up-regulated in the presence of naringin in a dose-dependent manner from 1 μg/mL to
100 μg/mL, while 200 μg/mL significantly inhibited their expressions.
3.4 Discussion
The present study demonstrated that naringin, the main effective component of
Chinese herb rhizoma drynariae, could promote the proliferation and osteogenesis of
hAFSCs. The enhancing effect of naringin on hAFSCs’ osteogenic differentiation might
explain the mechanism which credits the common use of rhizome drynariae for bone
formation in Asia.
The addition of naringin exhibited a biphasic effect on cell proliferation and ALP
activity of hAFSCs. At a concentration of 200 µg/mL, naringin suppressed the growth
and moderately increased ALP activity of hAFSCs, while at lower concentrations (1-100
µg/mL), naringin significantly enhanced cell proliferation and ALP activity in a dose99
dependent manner. The process of osteogenesis of up-stream stem cells can be depicted
into three major stages: stem cells to osteoprogenitor, then to preosteoblast, and finally to
mature osteoblast [18]. In this study, it was found that osteogenesis of hAFSCs was
promoted by naringin at both early and later stages. The earlier stage promotion was
evidently observed by the up-regulation of ALP activity on day 7. ALP, a significant
enzyme in the process of bone formation, enhances the mineralization of bone matrix by
transforming the phosphoric ester into inorganic phosphorous to increase the
phosphorous concentration [6]. In this study, ALP was used as an indicator of early
osteogenic differentiation of hAFSCs. The later stages of osteogenesis enhancement was
demonstrated by the expression of marker genes including OPN and Collagen I on day 21
and extracellular mineralization and calcium content on day 21 and 28. Therefore,
naringin could enhance osteogenesis of hAFSCs at both early and late stages.
Bone remodeling includes two processes, bone formation and bone resorption.
Osteoblasts are responsible to secrete new bone (bone formation), and osteoclasts deal
with breaking bone down (bone resorption). An imbalance in the regulation of bone
formation and bone resorption results in many bone diseases such as osteoporosis [19].
RANKL (receptor activator of nuclear factor ҡB ligand) is critical in the maturation and
activity of osteoclasts [20]. OPG is a soluble factor regulating bone mass [18] and a
decoy receptor binding to RANKL, therefore inhibiting osteoclast differentiation [20].
Therefore, the ratio of OPG to RANKL is a good indicator in the regulation of osteoblast
and osteoclast formation. An increase in the OPG/RANKL ratio favors bone formation
while a decrease in the ratio favors bone resorption. Our study showed that naringin
100
significantly increased the OPG expression with minimal RANKL expression during
osteogenic differentiation of hAFSCs, indicating the inhibition effect of naringin on
osteoclastogenesis. Thus, naringin may be used to enhance osteogenic differentiation of
hAFSCs and other MSCs to heal bone resorbing diseases, such as osteoporosis and boneerosive rheumatoid arthritis.
Osteogenesis is a complicated process involving several signaling pathways,
including BMP and canonical Wnt pathways [17, 21, 22]. The up-regulation of the
mRNA expression of BMP-related regulators (BMP4 and RUNX2) and Wnt-related
regulators (β-catenin and Cyclin D1) in our study suggested an involvement of both BMP
and Wnt signaling pathways in the hAFSC osteogenic process (Figure 3.7). BMPs are
responsible for maintaining the skeleton and facilitate the recruitment of osteoblast
precursors to a certain location during the later embryogenesis development.23 They are
essential to inducing ectopic bone formation, especially for the osteogenic differentiation
of non-bone cells [24, 25]. It has been reported that BMPs promoted osteoblastic
differentiation by up-regulating the expression of structural bone proteins, such as
Collagen I, and enhancing the mineralization of bone matrix [26]. RUNX2 is an
important downstream regulator of the BMP pathway [23, 27]. It is considered as the
master osteogenic transcription factor in the osteoblast maturation process [28] and plays
an essential role in osteoblast marker gene expression [29]. It was reported that BMP
regulator induced the expression of RUNX2 which regulates the expression of other
factors that act during the terminal osteogenic differentiation and bone-specific
extracellular matrix secretion [23, 30, 31]. In our study, the BMP-related regulator,
101
RUNX2, was found closely associated with the enhancing effect of naringin, suggesting
the involvement of BMP signaling pathway in the naringin-promoted osteogenic process
of hAFSCs.
The Wnt/β-catenin signaling is another critical pathway for osteogenic
differentiation and bone formation [17, 32].
Wnts participate in embryonic skeletal
patterning, fetal skeletal development, and adult skeletal remodeling [17]. Activation of
canonical Wnt signaling resulted in higher bone density [33, 34] and higher expression of
alkaline phosphatase, an early osteoblast marker [26, 35]. Previous studies have shown
that Wnt signaling contributed to osteoblast differentiation through the activation of βcatenin [35-37], whose activity is significant for the differentiation of mature osteoblasts
and bone formation [32, 38, 39]. Cyclin D1 is a target gene of Wnt pathway, which was
up-regulated when Wnt/β-catenin signaling was activated [40]. In this study, the mRNA
expressions of β-catenin and Cyclin D1 were enhanced in the presence of naringin,
suggesting that Wnt/β-catenin signaling was involved in the naringin-enhanced
osteogenesis of hAFSCs. RUNX2 was reported to integrate Wnt signaling for mediating
the process of osteoblast differentiation [41, 42] and it was also involved in BMP
signaling as discussed above. Therefore, RUNX2 behaves as cross-talking regulator
between BMP and Wnt/β-catenin signaling pathways.
Most of osteoprotective medicines have some adverse effects. For instance,
increased risk of cancer [43] and cardiovascular diseases [44] were reported to be
associated with hormone replacement therapy. It has also been reported that antiresorptive bisphosphonate might result in upper gastrointestinal tract complications [45]
102
as well as long-range effects on skeletons, especially in regard to bone turnover and
strength [46]. For Chinese herbal medicine, the dose-dependent cytotoxicity effects have
been observed in our cell-based screening study [47, 48]. The present results showed that
1100 µg/mL naringin had no cytotoxicity to hAFSCs. Moreover, naringin is a natural
gradient in Rhizoma drynariae and grapefruit, and has been widely used as a nutrient
supplement. Therefore, naringin itself could be an osteoprotecitve medicine as well as the
promising agent to enhance osteogenic differentiation of hAFSCs with low toxicity.
3.5 Conclusions
Osteoporosis can lead to fracture and deformities, and is a crucial public health
problem. Osteogenic cells differentiated from hAFSCs could be used to augment bone
formation and consequently in the treatment of osteoporosis and other bone-related
diseases. Chinese herb rhizome drynariae, which is safe and cheap, has been used for
fracture and bone healing for thousands of years. The present study demonstrated that
naringin, the main effective component of rhizome drynariae, could promote the
proliferation and osteogenesis and concurrently inhibit osteoclastogenesis of hAFSCs.
Moreover, the results also suggested that naringin may promote the osteogenic
differentiation of hAFSCs through both BMP and Wnt/β-catenin signal transduction
pathways. Due to the therapeutic efficiency, economic and safety advantages, naringinenhanced osteogenesis of hAFSCs would be an attractive treatment strategy to augment
bone formation in patients with osteoporosis and other bone disorders.
103
Acknowledgments
This work was supported in part by Alumni Grants for Graduate Research and
Scholarship (AGGRS) of The Ohio State University. We would like to acknowledge Dr.
Anthony Atala and Dr. James Yoo of the Wake Forest Institute for Regenerative
Medicine (Winston-Salem, NC) for kindly providing hAFSCs used in this study. We also
would like to acknowledge Dr. Sebastien Sart for the assistance in Image J analysis.
104
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Table 3.1. Primers used in the RT-PCR for osteogenic differentiation of hAFSCs.
Gene
Forward primera
Reverse primera
RUNX2
AGTGGACGAGGCAAGAGTTTC
CCTTCTGGGTTCCCGAGGT
OPN
GAGACCCTTCCAAGTAAGTCCA
GATGTCCTCGTCTGTAGCATCA
Collagen I ACAGCCGCTTCACCTACAGC
TGCACTTTTGGTTTTTGGTCAT
ALP
CTGGTAGTTGTTGTGAGC
CCCAAAGGCTTCTTCTTG
Cyclin D1 CCCTCGGTGTCCTACTTCA
GTTTGTTCTCCTCCGCCTCT
β-catenin
TGGCAACCAAGAAAGCAAG
CTGAACAAGAGTCCCAAGGAG
BMP4
CGAATGCTGATGGTCGTTT
CAGGGATGCTGCTGAGGTTA
OPG
TGCTGTTCCTACAAAGTTTACG
CTTTGAGTGCTTTAGTGCGTG
RANKL
CCAGCATCAAAATCCCAAGT
CCCCAAAGTATGTTGCATCCTG
GAPDH
GTGGTCTCCTCTGACTTCAACA
CTCTTCCTCTTGTGCTCTTGCT
a
Tm (ºC)b
62
62
52
Touch downb
55
55
Touch down
52
Touch down
62
Sequences are depicted in 5’-3’ direction.
b
Tm is the annealing temperature at which the primer binds to the RNA template during
polymerase chain reaction. Touch down: Tm from 62 to 52 °C, decrease 0.5 °C per cycle and the
following cycles were run at 52 °C. All the genes used 35 cycles.
Abbreviations: Osteogenic genes: osteopontin (OPN), collagen I, and alkaline phosphatase (ALP);
Genes in bone morphogenetic protein (BMP) pathway: runt related transcription factor 2
(RUNX2) and BMP4. Genes in Wnt pathway: β-catenin and Cyclin D1. Genes in osteoclast
differentiation: osteoprotegerin (OPG) and receptor activator of nuclear factor kappa-B ligand
(RANKL). Housekeeping gene: glyceraldehyde-3-phosphate dehydrogenase (GAPDH).
112
Figure 3.1 Chemical structure of naringin.
113
Figure 3.2. Effect of naringin on the proliferation of hAFSCs. N0: Control; N1: 1
µg/ml naringin; N10: 10 µg/ml naringin; N100: 100 µg/ml naringin; N200: 200 µg/ml
naringin. * indicated p < 0.05.
114
Figure 3.3. ALP activity of hAFSCs after naringin treatment. The percentage of
increase was calculated as
-
N0: Control; N1: 1 µg/ml naringin; N10: 10
µg/ml naringin; N100: 100 µg/ml naringin; N200: 200 µg/ml naringin.
115
Figure 3.4. Osteogenic differentiation of hAFSCs after naringin treatment. (A)
Alizarin red S (ARS) staining of hAFSCs after naringin treatment. (B) Calcium
deposition of naringin treated hAFSCs. The percentage of increase was calculated as
-
N0: Control; N1: 1 µg/ml naringin; N10: 10 µg/ml naringin; N100: 100
µg/ml naringin; N200: 200 µg/ml naringin.
116
Figure 3.5. RT-PCR analysis of naringin-enhanced osteogenic differentiation of
hAFSCs. (A) OPN, Collagen I, and ALP gene expression; (B) OPN, Collagen I, and
ALP expression normalized to GAPDH; (C) OPG and RUNXL gene expression; (D)
OPG and RUNXL gene expression normalized to GAPDH. N0: Control; N1: 1 µg/ml
naringin; N10: 10 µg/ml naringin; N100: 100 µg/ml naringin; N200: 200 µg/ml naringin.
117
Figure 3.6. RT-PCR analysis of naringin-enhanced BMP and Wnt signaling of
hAFSCs. (A) Gene expression of BMP pathway related regulators BMP4 and RUNX2;
(B) BMP4 and RUNX2 gene expression normalized to GAPDH; (C) gene expression of
Wnt pathway related regulators β-catenin and Cyclin D1; (D) β-catenin and Cyclin D1
gene expression normalized to GAPDH. N0: control; N1: 1 µg/ml naringin; N10: 10
µg/ml naringin; N100: 100 µg/ml naringin; N200: 200 µg/ml naringin.
118
Figure 3.7. Schematic illustration of BMP and Wnt-signaling pathways in naringinenhanced osteogenic differentiation of hAFSCs. BMP induces RUNX2 expression
which regulates the expression of other factors that act during terminal osteogenic
differentiation. Wnt signaling contributes to osteoblast differentiation through β-catenin
activation, which is responsible for the differentiation of mature osteoblasts and bone
formation. Cyclin D1 is a target gene of Wnt pathway, which is up-regulated when
Wnt/β-catenin signaling is activated
119
Chapter 4
Curculigoside Improves Osteogenesis and Inhibits
Osteoclastogenesis of Human Amniotic Fluid Derived Stem
Cells
Abstract:
Curculigoside, a phenolic glycoside, is the main active compound of curculigo
orchioides (Amaryllidaceae, rhizome). Curculigo orchioides is a traditional Chinese
herbal medicine and has been commonly used to treat orthopedic disorders and bone
healing in Asia. This study evaluated the effect of curculigoside on osteogenic
differentiation of human amniotic fluid derived stem cells (hAFSCs). The results showed
that curculigoside stimulated alkaline phosphatase activity (ALP) and calcium deposition
of hAFSCs during osteogenic differentiation in a dose-dependent manner (1-100 μg/mL)
while the effects were reduced at high concentration (200 μg/mL). From RT-PCR
analysis, the osteogenic genes osteopontin (OPN) and Collagen I were up-regulated with
120
curculigoside treatment (1-100 μg/mL). Concurrently, the ratio of osteoprotegerin (OPG)
to receptor activator of nuclear factor kappa-B ligand (RANKL) was increased, indicating
the inhibition of osteoclastogenesis by curculigoside. Moreover, the role of Wnt/β-catenin
signaling during curculigoside treatment was revealed by the upregulation of β-catenin
and Cyclin D1. In summary, curculigoside improved osteogenesis and inhibited
osteoclastogenesis of hAFSCs, demonstrating to be a novel approach to regulate hAFSC
osteogenic differentiation for treating bone disorders.
4.1 Introduction
Osteoporosis, characterized by low bone mineral density and microarchitecture
deterioration, results in structural instability of bone tissue and a high fracture risk,
causing serious issues to public health [1,2]. Osteoporosis has become a major health
hazard affecting more than 200 million people worldwide, and is considered to be one of
the highest disease incidences in aged people [3-5]. In the United States, national costs on
the medical care expenses associated with bone fractures was more than $17 billion in
2005 and the estimated cumulative cost is $474 billion by 2025 [6]. Current treatments
include estrogen replacement therapy (ERT) and bisphosphonate therapy. However, a
long-term use of estrogen could increase the risk of breast cancer, endometrial carcinoma
and cardiovascular diseases, while bisphosphonate therapy only inhibited the resorption
of osteoclast and caused acute incapacitating bone, joint, and muscle pain [1,7,8]. Hence,
there is an urgent need to develop effective therapies to ease the economic burden and
promote the life quality especially for the elder population.
121
Amniotic fluid derived stem cells (AFSCs) are reported to have strong potential to
differentiate into osteoblasts and arising as a novel cell source for treating bone diseases
[9,10]. AFSCs possess a phenotype of mesenchymal stem cells (MSCs), which have the
ability to migrate and engraft into multiple musculoskeletal tissues, especially sites of
injury, and undergo site-specific osteogenic differentiation [11]. Amniocentesis is a
widely accepted procedure in prenatal testing and presents a low risk for both the mother
and the fetus, enabling the easy derivation of AFSCs [12]. Compared to MSCs derived
from bone marrow, AFSCs have fetal tissue origin and an extensive self-renewal capacity
[13]. Compared to pluripotent stem cells, AFSCs have no ethical concerns and no
tumorigenic risk involved in their usage [14]. Therefore, AFSCs have become ideal
candidates for cell-based therapies to improve bone formation in patients who suffer from
diverse metabolic and genetic bone diseases, including osteoporosis.
Curculigo orchioides (Amaryllidaceae, rhizome) is a traditional Chinese herbal
medicine, which has been commonly used to treat orthopedic disorders and bone healing
for thousands of years [15,16]. Modern pharmacological study indicates that
curculigoside, a phenolic glycoside (Figure 4.1), is the main active compound of
curculigo orchioides. Curculigoside has been found to enhance the proliferation of mouse
pre-osteoblastic cells and stimulate the secretion of vascular endothelial growth factor
(VEGF) and bone morphogenetic protein (BMP)-2 [17,18]. It is also reported that
curculigoside prevented oxidative damage and inhibited osteoclastogenesis of rat bone
marrow cells [16,19]. However, no study has investigated the effects of curculigoside on
MSC osteogenic differentiation to date. In addition, no detailed osteogenic differentiation
122
and the responsible signal transduction pathways have been elucidated for curculigosidetreated stem cells. Thus, this present study aims to investigate the effects of curculigoside
on the osteogenic differentiation of hAFSCs and the related signaling pathway. To our
knowledge, it is the first study to use curculigoside to enhance stem cell lineage
commitment.
4.2 Materials and Methods
4.2.1 Culture of human amniotic fluid stem cells (hAFSCs)
The hAFSCs were isolated and cultured as previously described [9]. hAFSCs at
passages 16-18 were used in this study. All culture regents were from Life Technologies
unless otherwise noted. The cells were maintained in alpha-minimum essential medium
(α-MEM) supplemented with 15% embryonic stem cell qualified-fetal bovine serum (ESFBS), 100 U/mL penicillin, 100 µg/mL streptomycin, 2 mM L-glutamine, 18% Chang B,
and 2% Chang C (Irvine Scientific, Santa Ana, CA). The hAFSCs were subcultured at 70%
confluence. Culture medium was changed every 3 days.
4.2.2 hAFSC treatment with curculigoside
Curculigoside (≥99.6% purity) was purchased from Chinese National Institute
for the Control of Pharmaceutical and Biological Products (Guangzhou, China). The cells
(1×104 cells/ml) were seeded in 48, 24, and 6-well plates and cultured in growth medium
until 70-80% confluence. Then the cells were treated with differentiation medium which
contains α-MEM, 17% FBS (Atlanta Biologicals, Atlanta, GA), 2 mM L-glutamine, 100
U/mL penicillin, and 100 µg/mL streptomycin. Various amounts of curculigoside were
123
supplemented in the differentiation medium at concentrations of 1, 10, 100, or 200 µg/ml.
Cells cultured in medium in the absence of curculigoside were used as negative control.
Media were changed every 3 days.
4.2.3 Cell proliferation analysis
hAFSCs (1×104 per well) were seeded in a 48-well plate. After 24 h of incubation,
the growth medium was changed into curculigoside-containing media at a concentration
of 0 (Control), 1, 10, 100, and 200 µg/ml of curculigoside accordingly. Cells were
incubated at 37 °C in a humidified 5% CO2 incubator for 1, 2, 3 or 4 days. After that, the
medium was replaced with 500 µl of 10% Alamar Blue (AbD Serotec, Raleigh, NC)
solution at 37 °C for 3 h. The fluorescence of the medium was then monitored in
triplicate at 535 nm excitation wavelength and 590 nm emission wavelength using a
GENios Pro plate reader (Tecan, Research Triangle Park, NC). The fluorescence intensity
was correlated to the cell number, using a standard calibration curve.
4.2.4 Alkaline phosphatase activity (ALP) assay
Osteogenesis of hAFSCs was induced by the differentiation medium containing
different concentrations of curculigoside (0-200 μg/ml). At day 7, cells were washed with
PBS twice and lysed with the lysis buffer consisting 20 mM Tris-HCl (pH 7.5), 150 mM
NaCl, and 1% Triton X-100 for 5 min. The chromogenic substrate for ALP was pnitrophenyl phosphate (pNPP; Sigma-Aldrich). A 50 µL supernatant of lysate was mixed
with 50 µL pNPP (1mg/ml) substrate solution containing 1.0 mg/mL pNPP, 0.2 M Tris
buffer and 5 mM MgCl2 and incubated at 37 °C for 15 min on a Belly Button Shaker
124
(MidSci, St. Louis, MO). The reaction was stopped by the addition of 25 µL of 3 N
NaOH. Absorbance of p-nitrophenol released by the samples was measured at 405 nm
using a SpectraMAX 250 microplate reader (Molecular Devices, Sunnyvale, CA). Protein
concentration of cell lysate was determined using the Bradford assay at 595 nm on the
SpectraMAX 250 microplate reader. ALP activity was normalized to the total protein
content of cell lysate and expressed as nmol (p-nitrophenyl)/min/mg protein.
4.2.5 Assay of calcium deposition
To quantify mineralization, the calcium deposited by hAFSCs after 14 days was
measured using the Calcium Assay (Genzyme Diagnostics, Charlottetown, PE, Canada).
Briefly, samples were added with 1 M acetic acid and placed on a vortex overnight at
4 °C to extract the calcium from the mineralized matrix. In a 96 well clear polycarbonate
plate, 15 µL of cell extract was mixed with 150 µL of the Calcium Assay reagent and
incubate for 30 s at room temperature. The absorbance was determined at 650 nm on a
SpectraMAX 250 microplate reader. The samples were measured in triplicate and
compared to the calcium calibration curve.
4.2.6 Reverse transcriptase polymerase chain reaction (RT-PCR)
Total RNA was isolated using TRIZOL reagent (Invitrogen, Carlsbad, CA) from
hAFSCs treated with different concentrations of curculigoside after 21 days. RNA
concentrations were measured using a ND-1000 spectrophotometer (NanoDrop
Technologies, Wilmington, DE). After that, 1 µg of RNA was initially reverse
transcribed into cDNA using SuperScript™ III First-Strand Synthesis System
125
(Invitrogen). Then 200 ng of the cDNA was used as a template for the amplification of
target genes using the Quick-Load® Taq 2X Master Mix Kit (BioLabs, Ipswich, MA).
The primer sequences of the analyzed genes and PCR conditions are listed in Table 4.1.
For osteogenic differentiation, expressions of osteopontin (OPN) and collagen I were
measured. For osteoclast differentiation, osteoprotegerin (OPG) and receptor activator of
nuclear factor kappa-B ligand (RANKL) were assessed. For Wnt pathway, β-catenin,
Cyclin D1, and runt related transcription factor 2 (RUNX2) were analyzed. The
housekeeping gene, glyceraldehyde-3-phosphate dehydrogenase (GAPDH), was used as
an endogenous reference gene. Amplified products were fractionated in a 2% agarose
(Fisher Scientific, Pittsburgh, PA) gel at 70 V for 80 min and visualized and
photographed with a Gel Doc 2000 Gel Documentation System (Bio-Rad, Hercules, CA).
The expression level of each gene was analyzed by Image J Software and normalized to
the GAPDH expression.
4.2.7 Statistical analysis
Unless otherwise noted, all experiments and samples were triplicated.
Experimental results were presented as mean ± standard deviation (SD) (n = 3) and
analyzed using ANOVA followed by paired Tukey-Kramer analysis using JMP 7.0 (SAS
Institute Inc., Cary, NC). p < 0.05 was considered as statistically significant.
126
4.3 Results
4.3.1 Effect of curculigoside on the proliferation of hAFSCs
The stimulation effect of curculigoside on the proliferation of hAFSCs during a 4day culture was evaluated at various concentrations (1  200 µg/mL) (Figure 4.2). In the
presence of curculigoside at 1 or 10 µg/mL, the growth of hAFSCs from day 1 to day 4
was significantly increased compared to the control (Figure 4.2A). Specifically, 1 µg/mL
curculigoside increased the proliferation by 13%, 33%, 34% and 15% on day 1, 2, 3, and
4, respectively; and 10 µg/mL curculigoside increased the proliferation by 21%, 39%, 13%
and 19%, respectively (Figure 4.2B). In 100 µg/mL curculigoside group, the cell growth
was similar on day 1 but remarkably increased on day 2 (i.e. 47%) compared to control.
Then on day 3 and day 4, the cell number became close to the control group again.
However, 200 µg/mL curculigoside inhibited the proliferation of hAFSCs over the 4-day
culture (-7% to -32%), indicating that high concentration (≥200 µg/mL) of curculigoside
was harmful to cell growth. Therefore, there was no cytotoxic effect of curculigoside
within a range of 1-100 µg/mL on hAFSCs and curculigoside could be used to stimulate
hAFSC proliferation.
4.3.2 Effect of curculigoside on ALP activity of hAFSCs
ALP activity was used to indicate the early stage of osteogenic differentiation of
hAFSCs. ALP is a significant enzyme in the process of bone forming. It enhances the
mineralization of bone matrix by increasing the phosphorous concentration [8,20].
Curculigoside was shown to increase the ALP activity of hAFSCs in a dose-dependent
127
manner in the range of 1-100 µg/mL during a 7-day culture (Figure 4.3). ALP activity
was increased by 26.3% and 356% in the presence of 10 µg/mL and 100 µg/mL
curculigoside, respectively. The ALP activity of hAFSCs treated by 200 µg/mL
curculigoside was increased by 251%, lower than hAFSCs treated by 100 µg/mL
curculigoside.
4.3.3. Effect of curculigoside on calcium deposition
The osteogenic differentiation of hAFSCs was also investigated by quantifying
the calcium deposition on day 14 (Figure 4.4). In the presence of curculigoside, more
deposited calcium was formed than in the control group. Comparing to control group,
calcium content was increased by 45.5%, 92.7%, and 367% in the presence of 1 µg/mL,
10 µg/mL, and 100 µg/mL curculigoside, respectively. The calcium content in 200
µg/mL curculigoside group was less than that of 100 µg/mL group, increasing 243%
comparing to control. Thus, calcium deposition increased in a dose-dependent manner at
the concentration of 1-100 µg/mL curculigoside.
4.3.4 Effect of curculigoside on the expression of osteogenic genes
To further confirm the effects of curculigoside on osteogenic differentiation of
hAFSCs, RT-PCR was performed to detect osteogenic marker genes OPN and Collagen I.
The results showed that OPN and Collagen I expressions were significantly upregulated
in curculigoside-containing media comparing to the control group on day 21 (Figure
4.5A, 4.5B). In addition, curculigoside increased the expression of OPN and Collagen I in
128
a dose-dependent manner at concentrations of 1, 10, and 100 µg/mL, while at 200 µg/mL
the expressions of OPN and Collagen I were significantly reduced.
4.3.5 Effect of curculigoside on the osteoclast differentiation of hAFSCs
Expressions of OPG and RANKL were evaluated to assess the effects of
curculigoside on the osteoclast differentiation of hAFSCs. The OPG expression was
markedly increased with the concentration of curculigoside at the range of 1-100 µg/mL
and then decreased at 200 µg/mL (Figure 4.5C, 4.5D). RANKL, although weakly
expressed, also showed dose-dependent increase in the range of 1-100 µg/mL and the
decreased expression at 200 µg/mL. The ratio of OPG to RANKL (OPG/RANKL) is a
critical indicator for the regulation of osteoclast formation [21]. In this study, the
OPG/RANKL ratio increased from 1.6 to 5.3 as the curculigoside concentration increased
from 0 to 100 µg/mL (Table 4.2). Comparing to control, the OPG/RANKL ratio
increased by 28.9%, 30.7%, 233% and 104% at concentrations of 1, 10, 100 and 200
µg/mL, respectively, indicating that curculigoside inhibited osteoclastogenesis by
increasing the relative portion of OPG expression.
4.3.6 Effect of curculigoside on Wnt/β-catenin signaling pathway
Canonical Wnt signaling pathway was reported to involve in osteogenic
differentiation of MSCs [22-24]. Therefore, effects of curculigoside on the Wnt signaling
pathway were studied in the cells harvested on day 21. As seen in Figure 4.6, the mRNA
expression level of Wnt-related regulators, including β-catenin and Cyclin D1, were
significantly up-regulated in the presence of curculigoside in a dose-dependent manner at
129
1, 10, and 100 μg/mL. However, at 200 μg/mL, expressions of β-catenin and Cyclin D1
were significantly reduced compared to 100 μg/mL. The expression of RUNX2 was
weakly detected in the presence of curculigoside. Although BMP signaling was also
reported to regulate osteogenic differentiation, BMP-related regulators, such as BMP2
and BMP4, were not detected in our study (data not shown).
4.4 Discussion
Chinese herb curculigo orchioides, growing in subtropical regions in Asia, has
been used to maintain healthy energy and support bone healing for thousands of years
[15,16]. The hypothesis of this study is that components in curculigo orchioides should
affect osteogenic differentiation of hAFSCs. Our results demonstrated that curculigoside,
the main effective component of curculigo orchioides, promoted osteogenic
differentiation of hAFSCs and concurrently inhibited the osteoclastogenesis.
The process of osteogenesis from stem cells was depicted into three major stages:
early stage from stem cell to osteoprogenitor, intermediate stage from osteoprogenitor to
preosteoblast, and late stage from preosteoblast to mature osteoblast (Figure 4.7A)
[22,25]. In this study, osteogenesis of hAFSCs was found to be promoted by
curculigoside at early, intermediate, and late stages, which were proved by the increased
ALP activity on day 7, the increased calcium deposition on day 14, and the up-regulated
expressions of OPN and Collagen I on day 21. The addition of curculigoside exhibited a
biphasic effect on cell proliferation and ALP activity of hAFSCs. At a concentration of
200 µg/mL, curculigoside suppressed the growth of hAFSCs; while at lower
130
concentrations (< 200 µg/mL), curculigoside enhanced cell proliferation. The ALP
activity was enhanced by curculigoside in a dose-dependent manner in the range of 1-100
µg/mL. Similarly, gene expressions of OPN and collagen I peaked at 100 µg/mL.
Therefore, at the appropriate dose level, curculigoside significantly enhanced
osteogenesis of hAFSCs.
Interactions between osteoblasts and osteoclasts maintain the balance between the
bone formation and bone resorption [26]. Osteoblasts are responsible to secrete new bone
(bone formation), and osteoclasts deal with breaking bone down (bone resorption). An
imbalance in the regulation of bone formation and bone resorption results in many bone
diseases such as osteoporosis [27]. RANKL is known to be the major factor responsible
for osteoclast differentiation by providing a signal to osteoclast progenitors through the
membrane-anchored receptor activator of NF-kB (RANK) [21]. OPG is a decoy receptor
binding to RANKL and its expression inhibits osteoclast differentiation [21]. Therefore,
the ratio of OPG/RANKL is an essential indicator in the regulation of osteoblast and
osteoclast formation. An increase in the OPG/RANKL ratio favors bone formation while
a decrease in the ratio favors bone resorption. Our study showed that curculigoside
significantly increased the OPG/RANKL ratio during hAFSC osteogenic differentiation
especially at 100 µg/mL, indicating the inhibiting effect of curculigoside on
osteoclastogenesis. Thus, curculigoside provides a new approach to treat hAFSCs and
other MSCs for cell-based therapy in bone resorbing diseases such as osteoporosis.
Osteogenesis is a complicated process which involves several signaling pathways
especially canonical Wnt pathway [28-30]. In our study, the mRNA expression of β131
catenin and Cyclin D1 were up-regulated by curculigoside along with OPN, collagen I,
and OPG/RANKL. The Wnt/β-catenin signaling is critical for osteogenic differentiation
and bone formation (Figure 4.7B) and has been shown to participate in embryonic
skeletal patterning, fetal skeletal development, and adult skeletal remodeling [22,28].
Wnt signaling represents both a cell autonomous mechanism for inducing osteoblasts and
a mechanism in fully differentiated osteoblasts for stimulating OPG production to inhibit
osteoclast formation [29,31]. Therefore, activation of canonical Wnt signaling resulted in
higher bone density and higher expression of alkaline phosphatase [32,33]. Wnt signaling
contributed to osteoblast differentiation through β-catenin activation [30]. High level of
β-catenin enhances bone formation, whereas knockdown of the β-catenin gene at an early
developmental stage causes abnormal osteoblast differentiation [29,34,35]. β-catenin also
regulates osteoclastogenesis through effects on the expressions of OPG and RANKL [35].
Cyclin D1 is the target gene of Wnt pathway and is up-regulated when Wnt/β-catenin
signaling is activated [36]. RUNX2 was reported to integrate Wnt signaling for mediating
the process of osteoblast differentiation and play an essential role in secretion of bonespecific extracellular matrix [37,38]. In this study, the expressions of β-catenin and
Cyclin D1 were enhanced in the presence of curculigoside, indicating that Wnt/β-catenin
signaling is involved in the stimulating effect of curculigoside on osteogenesis of
hAFSCs and the inhibiting effect of osteoclastogenesis.
Generally, osteoprotective medicines have potential adverse effects. For instance,
increased risk of cardiovascular diseases was reported to associate with hormone
replacement therapy [39] and anti-resorptive bisphosphonate might result in upper
132
gastrointestinal tract complications as well as long-range effects on skeletons [40].
Curculigoside is a natural ingredient in curculigo orchioides and can be used as a nutrient
supplement in the diet. Our results showed that 1-100 µg/mL curculigoside had no
cytotoxicity to hAFSCs while the high concentration of 200 µg/mL may inhibit the
growth. The dose effect of Chinese herbal medicine has been observed in our previous
study during cell-based assay development [41,42]. Therefore, with appropriate dose,
curculigoside-enhanced hAFSC osteogenic differentiation could be a promising strategy
to treat bone disorders such as osteoporosis.
4.5 Conclusions
Osteoporosis is a crucial public health problem which can lead to fracture and
deformities. Osteogenic differentiation of hAFSCs could be used for the treatment of
osteoporosis and other bone-related diseases. Chinese herb curculigo orchioides, which is
safe and cheap, has been used for fracture and bone healing for thousands of years. The
present study demonstrated that curculigoside, the main effective component of curculigo
orchioides, promoted the proliferation and osteogenesis and concurrently inhibited
osteoclastogenesis of hAFSCs. Moreover, it is elucidated that curculigoside functions
through the Wnt/β-catenin signal transduction pathway. Due to the economic and safety
advantages, curculigoside-enhanced osteogenesis of AFSCs would be a promising and
attractive treatment strategy to augment bone formation in patients with osteoporosis and
other bone disorders.
133
Acknowledgments
This work was supported in part by Alumni Grants for Graduate Research and
Scholarship (AGGRS) of The Ohio State University. We would like to acknowledge Dr.
Anthony Atala and Dr. James Yoo of the Wake Forest Institute for Regenerative
Medicine (Winston-Salem, NC) for kindly providing hAFSCs used in this study.
134
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Table 4.1. Primers used in the RT-PCR for osteogenic differentiation of hAFSCs.
Gene
Forward primera
Reverse primera
RUNX2
AGTGGACGAGGCAAGAGTTTC
CCTTCTGGGTTCCCGAGGT
OPN
GAGACCCTTCCAAGTAAGTCCA
GATGTCCTCGTCTGTAGCATCA
Collagen I ACAGCCGCTTCACCTACAGC
TGCACTTTTGGTTTTTGGTCAT
Cyclin D1 CCCTCGGTGTCCTACTTCA
GTTTGTTCTCCTCCGCCTCT
β-catenin
TGGCAACCAAGAAAGCAAG
CTGAACAAGAGTCCCAAGGAG
OPG
TGCTGTTCCTACAAAGTTTACG
CTTTGAGTGCTTTAGTGCGTG
RANKL
CCAGCATCAAAATCCCAAGT
CCCCAAAGTATGTTGCATCCTG
GAPDH
GTGGTCTCCTCTGACTTCAACA
CTCTTCCTCTTGTGCTCTTGCT
a
Tm (ºC)b
62
62
52
55
55
52
Touch down
62
Sequences are depicted in 5’-3’ direction.
b
Tm is the annealing temperature at which the primer binds to the RNA template during
polymerase chain reaction. Touch down: Tm from 62 to 52 °C, decrease 0.5 °C per cycle and the
following cycles were run at 52 °C. All the genes used 35 cycles.
Abbreviations: Osteogenic genes: osteopontin (OPN), and collagen I; Genes in Wnt pathway: βcatenin, Cyclin D1, and runt related transcription factor 2 (RUNX2). Genes in osteoclast
differentiation: osteoprotegerin (OPG) and receptor activator of nuclear factor kappa-B ligand
(RANKL). Housekeeping gene: glyceraldehyde-3-phosphate dehydrogenase (GAPDH).
141
Table 4.2. Effects of curculigoside on the osteoclast differentiation of hAFSCs
Curculigoside
OPG/RANKL
Percentage of increase
concentration
ratio
compared to control (%)
0
1.6
0.0
1
2.0
28.9
10
2.1
30.7
100
5.3
233.0
200
3.2
104.0
(μg/mL)
142
Figure 4.1. Chemical structure of curculigoside.
143
Figure 4.2. Effect of curculigoside on the proliferation of hAFSCs. (A) Proliferation
fold of initial cell number; (B) Percentage of proliferation increase relative to control,
calculated as
-
. C0: Control; C1: 1 µg/ml curculigoside; C10: 10 µg/ml
curculigoside; C100: 100 µg/ml curculigoside; C200: 200 µg/ml curculigoside. * p<0.05.
144
Figure 4.3. ALP activity of hAFSCs after curculigoside treatment. hAFSCs were
treated with curculigoside at different concentrations (1-200 μg/mL). The percentage of
increase was calculated as
145
-
Figure 4.4. Calcium deposition of hAFSCs after curculigoside
↑367% treatment. hAFSCs
were treated with curculigoside at different concentrations (1-200 μg/mL). The
↑243%
percentage of increase was calculated as
146
-
Figure 4.5. RT-PCR analysis of curculigoside-enhanced osteogenic differentiation of
hAFSCs. (A) OPN and Collagen I gene expression; (B) OPN, Collagen I, and ALP
expression normalized to GAPDH; (C) OPG and RUNXL gene expression; (D) OPG and
RUNXL gene expression normalized to GAPDH. C0: Control; C1: 1 µg/ml curculigoside;
C10: 10 µg/ml curculigoside; C100: 100 µg/ml curculigoside; C200: 200 µg/ml
curculigoside.
147
Figure 4.6. RT-PCR analysis of curculigoside-enhanced Wnt signaling of hAFSCs.
(A) Gene expression of Wnt pathway related regulators β-catenin, Cyclin D1, and
RUNX2; (B) β-catenin, Cyclin D1, and RUNX2 gene expression normalized to GAPDH.
C0: Control; C1: 1 µg/ml curculigoside; C10: 10 µg/ml curculigoside; C100: 100 µg/ml
curculigoside; C200: 200 µg/ml curculigoside.
148
Figure 4.7. Schematic illustration of Wnt signaling pathway in osteogenic
differentiation of hAFSCs. (A) A scheme of developmental stages during osteogenic
differentiation of hAFSCs. (B) Wnt signaling contributes to osteoblast differentiation
through β-catenin activation, which is responsible for the differentiation of mature
osteoblasts and bone formation. Cyclin D1 is a target gene of Wnt pathway, which is upregulated when Wnt/ β-catenin signaling is activated.
149
Chapter 5
Optimization of serum containing and serum free media for
expansion of human amniotic fluid stem cells
Abstract
Due to their easy and safe accessibility, abundant cell numbers, and lack of ethical
concerns, human amniotic fluid stem cells (hAFSCs) have emerged as an important cell
source for tissue engineering and regenerative medicine. However, little is done about the
development and optimization of culture mediums for superior growth and expansion of
hAFSCs for clinical application. In this study, the performance of a complete medium
and two serum free medium developed by Irvine Scientific in supporting in vitro
proliferation of hAFSCs was investigated and compared with a commonly used hAFSC
growth medium containing 15% embryonic stem cell qualified-fetal bovine serum (ESFBS), 2 mM L-glutamine, 18% Chang B, and 2% Chang C. Our results indicated that the
complete medium can support better cell growth than commonly used 15% ES-FBS
150
medium
while maintaining their immunophenotypic profile
and
multilineage
differentiation capacity.
5.1 Introduction
Amniotic fluid stem cells (AFSCs) is a novel cell source for tissue engineering
and regenerative medicine. Being present in amniotic fluid, for the first time AFSCs were
described to possess mesenchymal features and extensive proliferation abilities by
Kaviani et al. in 2001[1]. Then their phenotype and multilineage differentiation potential
similar to bone marrow mesenchymal stem cell were demonstrated by In ’t Anker et al. in
2003 [2]. AFSCs are not tumorigenic and have no ethical concerns involved in the their
usage, therefore they have become superior candidates for cell based therapies. However,
currently, most hAFSC proliferations are carried out in a commonly used serum-rich
medium as described [3], and to date, very few optimized medium formulae have been
developed for superior growth and expansion of hAFSCs.
Cell culture medium is a mixture consisting of amino acids, a source of energy
(such as glucose), vitamins, growth factors, trace elements, etc. in a pH buffered salt
solution [4]. Traditional mammalian cell culture formulations require further
supplementation with a protein source, such as serum, to maintain and proliferate cells.
Fetal bovine serum (FBS) is the present standard serum. It is a complex mixture
containing a large number of ingredients, such as proteins, growth factors, hormones,
vitamins, trace minerals and so on, which are essential for mammalian cells [5]. However,
the serum composition continually varies with season and producing batch and is ill151
defined. Moreover, because of the threat of contamination of viral, bacterial, and prion
pathogens, the use of animal-based products is firmly dejected for production of
medicinal products [6-8]. It is also reported that exposure of human cells to FBS resulted
in fixation of animal proteins on the human cell surface thus made the host more prone to
inflammatory and/or adverse immunemediated events [9-11]. Therefore, mammalian cell
culture media are directed to progress from serum-containing to serum-free, to animalcomponent-free and then to chemically defined formulations [4, 5].
In this study, a serum-containing and two serum-free media developed by Irvine
Scientific were investigated for supporting the expansion of hAFSCs. The cell specific
growth rate, doubling time and viability in these cultures were determined and compared
with those in a commonly used serum-containing medium. Further, the
immunophenotype and multipotency of expanded cells were studied. In conclusion, this
study performed a contribution to the development of medium for the expansion of
hAFSCs for clinical applications.
5.2 Materials and methods
5.2.1 Cultures and media
The hAFSCs were isolated and cultured as previously described [12]. hAFSCs at
passages 16-18 were used in this study. They were cultured in a currently commonly used
serum containing fresh medium (Control medium), Irvine Scientific developed serum
containing complete medium (SCC medium) and serum free medium (SF I and SF II
medium), respectively. Control medium contains alpha-minimum essential medium (α152
MEM) supplemented with 15% embryonic stem cell qualified-fetal bovine serum (ESFBS), 100 U/mL penicililin, 100 µg/mL streptomycin, 2 mM L-glutamine (Gibco, Grand
Island, NY), 18% Chang B, and 2% Chang C (Irvine Scientific, Santa Ana, CA).
Formulae of SCC, SF I and SF II medium are confidential. These cells were subcultured
and expanded at 70% confluence. Culture medium was changed every 1-3 days,
according to the actual metabolic activities of cells..
5.2.2 Expansion of hAFSCs over 3 passages
Cells were seeded in a 6-well plate at a density of 5×103 cells/cm2 and cultured in
Control, SCC and SF medium, respectively, until 70-80% confluence. Then cells were
harvested and seeded in another 6-well plate for the 2nd passage. For the 3rd passage, cells
were cultured in 75 cm2 T-flasks to generate more cells (Figure 5.1). Cell numbers and
viabilities were measured for each passage.
5.2.3 Morphology of hAFSC cultures
hAFSCs were cultured in each medium for 3 passages. The cell morphologies
were observed with a light microscope Olympus IX71 (Olympus Corporation, Tokyo,
Japan) and images were documented.
5.2.4 Flow cytometry analysis of immunophenotype
To identify the effects of the expansion in different growth mediums on the
characteristic immunophenotype of hAFSCs, flow cytometric analysis of anti-CD29,
anti-CD44 (Developmental Studies Hybridoma Bank, Iowa City, Iowa), anti-CD90, and
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anti-CD34 (BD, Franklin Lakes, NJ) was performed. Cells were harvested using TriplE
Select (10X) (Gibco) and dissociated into individual cells in solution prior to the flow
cytometric analysis. Samples were fixed with 4% paraformaldehyde in PBS at room
temperature for 20 min. After being washed with PBS three times, the fixed cells were
permeabilized and blocked in 3% FBS and 0.1% Triton X-100 in PBS for 1 h at room
temperature and incubated overnight at 4 oC with the primary antibody in 1% FBS in
PBS. Stained cells were washed three times in PBS and incubated with an appropriate
isotype-matched secondary antibody for 1 h at room temperature. Positive cells were
detected and quantified using an FACS Calibur instrument and CellQuest software
(Becton Dickinson, Franklin Lakes, NJ). Cells labeled with only the secondary antibody
were used as controls to evaluate the non-specific binding or background fluorescence
reading.
5.2.5 Multilineage differentiation assays
To test their multipotency potential, hAFSCs harvested from different mediums
were cultured under conditions allowing for osteogenic or adipogenic differentiation. The
hAFSCs were seeded into six-well plates at 10000 cells/cm2 and cultured until 70-80%
confluence. Cells cultured in growth medium were used as negative control. For
osteogenic differentiation, cells were cultured for 21 days in the osteogenic medium
composed of the alpha-minimum essential medium (α-MEM) (Gibco) supplemented with
17% fetal bovine serum (FBS) (Atlanta Biologicals), 10 mM β-glycerol phosphate, 1 nM
dexamethasone, 50 µg/mL thyroxine (Sigma), 2 mM L-glutamine, 100U/mL penicililin,
and 100 µg/mL streptomycin (Gibco). Mediums were changed every 3 days. After 3
154
weeks, the cells were fixed with a 10% (v/v) solution of formalin and stained with 1%
(w/v) Alizarin red S solution, or immunostained with osteocalcin and DAPI. The
presence of calcium was observed with an Olympus IX71 microscope (Olympus
Corporation, Tokyo, Japan). For adipogenic differentiation, confluent hAFSCs were
cultured for 21 days in adipogenic medium composed of Dulbecco’s modified Eagle’s
medium (DMEM) supplemented with 10% fetal bovine serum (FBS) (Atlanta Biologicals,
Atlanta, GA), 1 µM dexamethasone, 1 mM 3-isobutyl-1-methylxanthine, 10 µg/mL
insulin, 60 µM indomethacin, 2 mM L-glutamine, 100 U/mL penicililin, and 100 µg/mL
streptomycin (Gibco). Mediums were changed every 3 days. After 21 days, the presence
of adipocyte in the cultured cells was stained with Oil Red O solution (1% (w/v).
Intracellular lipid vacuoles were visualized with a light microscope Olympus IX71
(Olympus Corporation, Tokyo, Japan).
5.2.6 Statistical analysis
Experimental results were presented as mean ± standard deviation (SD) (n = 3)
and analyzed by Student’s t-test with p < 0.05 as statistically significant.
5.3 Results and Discussion
5.3.1 Expansion of hAFSCs in Control and SCC medium
5.3.1.1 hAFSCs expansion in Control and SCC cultures over three passages
Cells were cultured in Control and SCC medium over three passages, respectively.
Figure 5.2 depicted growth curves for both cultures. The cell number harvested from
155
SCC culture was 44.2% and 115% higher than that from Control culture at the end of the
2nd and 3rd passage, respectively. Doubling time for each culture was calculated in Table
5.1. The doubling time of hAFSCs in SCC culture was 28.8% and 30.3% shorter than that
in Control culture for the 2nd and 3rd passage, respectively. Cell viabilities were > 98% for
both cultures. Therefore, hAFSCs presented a significantly faster proliferation in SCC
medium compared to in the currently commonly used Control medium.
5.3.1.2 Morphology of hAFSCs in Control and SCC cultures over three passages
Cell morphology was observed to investigate cell changes as response to different
culture mediums. As shown in Figure 5.3, both mediums supported proper hAFSC
morphology, showing well-grown epithelium-like cell layers of healthy cells. No changes,
such as vacuoles in the cytoplasma, were observed, providing no apparent evidences for
cell changes, such as apoptosis. These results indicate that normal cell morphology can
be maintained in both Control medium and SCC medium.
5.3.1.3 Flow cytometry analysis of immunophenotype
To investigate the influence of culture medium on the antigenic phenotype of
hAFSCs, cells from Control and SCC groups were examined for their MSC markers by
flow cytometry analysis. Considerable attempts have been made to define the phenotypic
profile of MSCs, and several surface markers have been proposed to characterize hMSCs
[13]. Generally, hMSCs express CD29 (β-integrins), CD44 (hyaluronan receptor), CD90
and CD105 (endoglin), but do not express markers of the hematopoietic lineage, such as
CD34 (hematopoietic progenitors receptor) [12, 14]. The results of flow cytometry
156
analysis are shown in Figure 4. hAFSCs from SCC group had high expression levels of
CD29 and CD44 (95.1% and 98.7%, respectively) and did not express CD34 (0.1%).
hAFSCs expanded in Control medium were also positive in expressing CD29 (88.8%)
and CD44 (98.1%), and negative in CD34 (0.3%). However, cultures expanded by
Control medium lost part CD29 expression (88.8%) while SCC medium maintained
hAFSC immunophenotypic profile well (95.1%) after 3 passages. Therefore, SCC
medium
showed a
better performance in
maintaining hAFSC
characteristic
immunophenotype, indicating it can prevent hAFSCs differentiating early and keep their
stemness and multilineage differentiation potential.
5.3.1.4 Multilineage differentiation of Control and SCC medium-expanded hAFSCs
To confirm the multipotency of the Control and SCC medium-expanded hAFSCs,
cells harvested from the two cultures were stimulated to differentiate along osteogenic
and adipogenic lineages and evaluated by histochemistry and immunostaining. As shown
in Figure 5.5A, the Control medium expanded-hAFSCs developed a calcium-rich
mineralized bone matrix along the cell membrane as large red aggregates embedded in
the ECM after 21 day osteogenic induction, as well as exhibited morphological changes
and small lipid vesicles in the cytoplasm as revealed by Oil Red O staining after 21 day
adiopogenic induction. SCC medium expanded-hAFSCs were stimulated to osteogenic
differentiation for 21 days and immunostained with osteocalcin and DAPI. Figure 5.5B
exhibits that a large number of osteoblasts which were stained green fluorescence were
developed from SCC medium expanded-hAFSCs. These results confirmed that the
Control and SCC medium-expanded hAFSCs were both capable of differentiating into
157
osteogenic and/or adipogenic lineages, suggesting that hAFSCs expansion in these two
cultures did not change their multipotency after 3 passages.
5.3.2 Expansion of hAFSCs in SF I, SF II and SCC medium
5.3.2.1 hAFSCs expansion in SF I, SF II and SCC cultures over three passages
Cells were also cultured in SF I, SF II and SCC medium over three passages,
respectively. Figure 5.6 depicted growth curves for these cultures. Unfortunately,
hAFSCs grew very slowly in SF I and SF II mediums. The cell number harvested from
SCC culture was 281% and 216% higher than that in SF I and SF II culture at the end of
the 2nd respectively, and 1020% and 606% higher than that in SF I and SF II culture at the
end of the 3rd passage respectively. Doubling time over 3 passages for these cultures was
calculated in Table 5.2. The doubling time of hAFSCs in SF I culture was 31.0%, 92.7%
and 88.1% longer than that in SCC culture for each passage, respectively; while the cell
doubling time in SF II culture was 53.3%, 114% and 170% longer than that in SCC
culture for each passage, respectively. In addition, as shown in Figure 5.7, cell viability
for SCC culture were above 98% over 3 passages, however, cell viability for SF I culture
decreased to 97.1% and 96.7% at the end of 2nd and 3rd passage respectively and cell
viability for SF II culture decreased to 96.1% and 94.9% at the end of 2nd and 3rd passage
respectively. Therefore, SF I medium showed a little better performance in supporting
hAFSC expansion than SF II medium did; however, both of SF I and SF II medium could
not support hAFSC growth as well as SCC medium and need further development and
optimization.
158
5.3.2.2 Morphology of hAFSCs in SF I, SF II and SCC culture over three passages
Cell morphology observation was conducted to investigate cell changes as
response to different culture mediums. As shown in Figure 5.8, in consistent with 5.3.2.1,
cells sparsely grew in SF I and SF II cultures and the cell density in these two cultures
were significantly lower than that in SCC culture, especially since the 2nd passage.
Additionally, in SF I and SF II cultures, cell morphology showed a sign of adipogenic
differentiation starting from the 2nd passage. These results indicated that SF I and SF II
medium could not support fast cell proliferation and normal cell morphology.
5.3.2.3 Flow cytometry analysis of immunophenotype
To investigate the influence of SF I and SF II medium on the antigenic phenotype
of hAFSCs, cells from SF I, SF II and SCC groups were examined for their MSC markers
by flow cytometry analyses. The results of flow cytometry analysis are shown in Figure
5.9. hAFSCs from SCC group exhibited high expression levels of CD90 (92.4%), while
cells cultured in SF I and SF II medium evidently lost part CD90 expression (62.8% and
82.6%, respectively) after 3 passages. These results are in consistent with 5.3.2.2,
indicating that SCC medium could maintain hAFSC characteristic immunophenotype,
however, SF I and SF II medium could not prevent hAFSCs differentiating early and
partially lost their stemness.
5.4 Conclusions
hAFSCs is a recently derived cell source and processes great potentials for
regenerative medicine and tissue engineering applications. However, very few mediums
159
are available for the expansion of hAFSCs. In this study, a serum containing complete
medium (SCC) and two serum free medium (SF I and SF II) were developed. hAFSCs
were cultured in these mediums and compared with a currently commonly used hAFSC
growth medium (Control) over 3 passages. Our results indicated that SCC medium could
support a faster proliferation than Control medium, meanwhile SCC medium maintained
the characteristic immunophenotype and the multilineage differentiation potential of
hAFSCs. However, the two serum free mediums SF I and SF II were not very satisfactory
and need further development and optimization.
160
References
1.
Kaviani A., Perry T.E., Dzakovic A., et al. The amniotic fluid as a source of cells
for fetal tissue engineering. Journal of Pediatric Surgery, 2001. 36(11): 16621665.
2.
in 'tAnker P.S., Scherjon S.A., Kleijburg-van der Keur C., et al. Amniotic fluid as
a novel source of mesenchymal stem cells for therapeutic transplantation. Blood,
2003. 102(4): 1548-1549.
3.
De Coppi P., Bartsch G., Siddiqui M.M., et al. Isolation of amniotic stem cell
lines with potential for therapy. Nature Biotechnology, 2007. 25(1): 100-106.
4.
Price P.J. Design, Optimization and Handling of Mammalian Cell Culture Media.
In Vitro Cellular & Developmental Biology-Animal, 2009. 45: S19-S19.
5.
van der Valk J., Brunner D., De Smet K., et al. Optimization of chemically defined
cell culture media - Replacing fetal bovine serum in mammalian in vitro methods.
Toxicology in Vitro, 2010. 24(4): 1053-1063.
6.
van der Valk J., Mellor D., Brands R., et al. The humane collection of fetal bovine
serum and possibilities for serum-free cell and tissue culture. Toxicology in Vitro,
2004. 18(1): 1-12.
7.
Schiff L.J. Review: Production, characterization, and testing of banked
mammalian cell substrates used to produce biological products. In Vitro Cellular
& Developmental Biology-Animal, 2005. 41(3-4): 65-70.
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8.
Kunisaki S.M., Armant M., Kao G.S., et al. Tissue engineering from human
mesenchymal amniocytes: a prelude to clinical trials. Journal of Pediatric Surgery,
2007. 42(6): 974-980.
9.
Mackensen A., Drager R., Schlesier M., et al. Presence of IgE antibodies to
bovine serum albumin in a patient developing anaphylaxis after vaccination with
human peptide-pulsed dendritic cells. Cancer Immunology Immunotherapy, 2000.
49(3): 152-156.
10.
Chachques J.C., Herreros J., Trainini J., et al. Autologous human serum for cell
culture avoids the implantation of cardioverter-defibrillators in cellular
cardiomyoplasty. International journal of Cardiology, 2004. 95: S29-S33.
11.
Spees J.L., Gregory C.A., Singh H., et al. Internalized antigens must be removed
to prepare hypoimmunogenic mesenchymal stem cells for cell and gene therapy.
Molecular Therapy, 2004. 9(5): 747-756.
12.
De Coppi P., Bartsch G., Siddiqui M.M., et al. Isolation of amniotic stem cell
lines with potential for therapy. Nature biotechnology, 2007. 25(1): 100-106.
13.
Baghaban E.M., Jahangir S., and Aghdami N. Mesenchymal stem cells from
murine amniotic fluid as a model for preclinical investigation. Archives of Iranian
medicine, 2011. 14(2): 96.
14.
Cao Y., Li D., Shang C., et al. Three-dimensional culture of human mesenchymal
stem cells in a polyethylene terephthalate matrix. Biomedical Materials, 2010. 5:
065013.
162
Table 5.1 Doubling time and seeding density of cultures in Control medium and SCC
medium over three passages.
Control
Medium
SCC Medium
Seeding Density
( cells/ cm2)
passage 16
47.63
57.72
5180
passage 17
51.63
40.10
2083
passage 18
63.00
48.33
5000
Doubling Time (h)
163
Table 5.2 Doubling time and seeding density of cultures in SF I, SF II and SCC medium
over three passages.
SF I
SF II
SCC
Medium
Medium
Medium
Seeding
Density
( cells/ cm2)
passage 16
124.30
145.47
94.91
5214
passage 17
78.22
87.05
40.58
5236
passage 18
109.51
157.06
58.23
5268
Doubling Time (h)
164
Figure 5.1 Scheme of medium optimization experiments.
165
12
Control medium
SCC medium
Viable Cell Count (M)
10
8
6
4
2
0
0
2
4
6
8
10
12
14
16
Day of Culture (Day)
Figure 5.2 hAFSCs growth over three passages in control medium and SCC medium.
166
Figure 5.3 hAFSC morphology in Control culture and SCC culture over 3 passages.
167
Figure 5.4 Phenotype of hAFSCs cultured in Control medium and SCC medium after 3
passages analyzed by flow cytometry.
168
Figure 5.5 Histochemistry and immunostaining of multipotent differentiation of
hAFSCs expanded in Control and SCC medium. (A) Osteogenic and adipogenic
differentiation of hAFSCs harvested from Control culture; (B) Osteogenic differentiation
of hAFSCs harvested from SCC culture.
169
Viable cell count (M)
5
4.5
SF I
4
SF II
3.5
SCC
3
2.5
2
1.5
1
0.5
0
0
2
4
6
8
10
12
14
16
Day of culture (Day)
Figure 5.6 hAFSCs growth over three passages in SF I, SF II and SCC media.
170
Viability (%)
100
90
SF I
SF II
SCC
80
0
5
10
15
Day of culture (Day)
Figure 5.7 Cell viability of hAFSCs in SF I, SF II, and SSC cultures over 3 passages.
171
Figure 5.8 hAFSC morphology in SF I, SF II and SCC media over 3 passages.
172
Figure 5.9 Phenotype of hAFSCs cultured in SF I, SF II and SCC media after 3 passages
analyzed by flow cytometry.
173
Chapter 6
Conclusions and Recomondations
6.1 Conclusions
Possessing many remarkable advantages, such as easy and safe accessibility,
extensive self-renewal capacity, abundant cell numbers, not tumorigenic, and lack of
ethical concerns, hAFSCs has emerged as a promising candidate for various therapeutic
applications. Due to their recent derivation, very few researches have been done on basic
research and clinical applications of hAFSCs. In this study, a PET based fibrous
bioreactor was develop to perform mass production of functional hAFSCs. Osteogenic
differentiation of hAFSCs aimed for osteoporosis therapy was investigated and promoted
by using natural plant ingredients, naringin and curculigoside. Expansion media for
hAFSCs were developed and optimized. The important results presented in previous
chapters are summarized below.
6.1.1 Mass production of hAFSCs in a 3-dimensional fibrous bed bioreactor
Compared to 2-D cultures, hAFSCs grown in 3-D microenvironments of PET
exhibited more stable long-term proliferation with a significantly higher expansion fold,
174
suggesting that the PET fibrous matrix is an effective 3-D support for culturing
anchorage-dependent hAFSCs. The dynamic culturing condition in the PET-based 3-D
dynamic fibrous-bed bioreactor stimulated nutrient, oxygen and metabolite transfers to
the hAFSCs residing within the 3-D scaffold and, consequently, promoted cell expansion
to reach a high cell yield, 32 folds of initial number. Meanwhile, the energy metabolism
in this 3-D bioreactor was significantly increased with apparent yield of lactate from
glucose reduced by 42.0% and 57.5% comparing to T-flasks and a reported glass carrier
based fixed bed bioreactor, respectively. Additionally, the bioreactor-expanded hAFSCs
were demonstrated to be able to maintain their immunophenotypic profile, multilineage
differentiation potential, and clonogenic ability. In conclusion, this PET-based 3-D
dynamic fibrous-bed bioreactor system can be easily and rapidly implemented for
clinical-scale expansion to maximize hAFSC yield while maintaining cell product quality
for regenerative medicine and cell therapy applications.
6.1.2 Promoted proliferation and osteogenic differentiation of hAFSCs treated with
naringin and curculigoside
Naringin and curculigoside, the main effective components of natural medicinal
plants, rhizome drynariae and curculigo orchioides, were demonstrated to be able to
enhance the proliferation of hAFSCs by 35% and 21.9%, respectively. Furthermore, the
osteogenic differentiation of hAFSCs were indicated to be markedly promoted by
naringin and curculigoside with with ALP activity enhanced by 163% and 356% and
calcium deposit enhanced by 239% and 367%. The responsible signaling pathways were
investigated and revealed: BMP and Wnt/β-catenin pathways are involved in naringin
enhanced-osteogenesis of hAFSCs, while Wnt/β-catenin pathway is involved in
175
curculigoside enhanced osteogenesis of hAFSCs. In addition, osteoclastogenesis was
found to be inhibited by naringin and curculigoside with the ratio of OPG/RANKL
enhanced by 273% and 231%, respectively.
6.1.3 Medium development and optimization for the expansion of hAFSCs
A serum containing complete medium (SCC) and two serum free medium (SF I
and SF II) developed by Irvine Scientific were investigated for supporting the expansion
of hAFSCs. SCC medium was indicated to support a faster proliferation than a currently
commonly used hAFSC growth medium (Control). Meanwhile, SCC medium
maintained the characteristic immunophenotype and the multilineage differentiation
potential of hAFSCs. However, the two serum free mediums SF I and SF II were not very
satisfactory and need further development and optimization.
6.2 Recommendations
The clinical stem cell dose is reported to be 1-5 ×106 cells per kg of patient body
weight [1]. Therefore, a large amount of hAFSCs will be required for clinical therapies.
In this study, 3.2 ×107 hAFSCs were produced in a 25ml 3-D bioreactor with a very high
cell density. Author suggested that in further this process should be scaled up to
bioreactors with a larger working volume in order to generate hAFSCs with higher yields.
Moreover, to meet good manufacturing practice (GMP) standards, automatically
controlled process with online measurement and adjustment of culture parameters, such
as temperature, pH, dissolved oxygen (DO), and shear force, should be developed. In
addition, two extracts of Chinese herbal medicine combinations were found to be able to
176
promote the proliferation of hAFSCs in 2D static culture, and the scale-up of this process
in a 3D dynamic bioreactor should be further investigated and developed.
Promoted osteogenic differentiation of hAFSCs was successfully achieved by
using naringin or curculigoside in 2-D culture in this work. Author thinks that scaling up
the differentiation process in bioreactors would be an interesting project. Other types of
bioreactors could also be taken into consideration for this purpose.
Chemically defined medium for clinical applications is another requirement of
GMP. In this study, a serum containing complete medium (SCC) was demonstrated to be
able to support a faster proliferation of functional hAFSCs than the commonly used
serum containing medium. However, the two serum free media (SF I and SF II) were not
satisfactory and need further optimization. In future research, hAFSC culture media
should be directed to progress from serum-containing to serum-free, to animalcomponent-free and finally to chemically defined formulations.
References
1.
Subbanna P.K.T. Mesenchymal stem cells for treating GVHD: in-vivo fate and
optimal dose. Medical hypotheses, 2007. 69(2): 469-470.
177
Appendix A
Comparison of effects of naringin and curculigoside on
osteogenesis of hAFSCs
Both naringin and curculigoside were demonstrated to be able to promote the
osteogenic differentation of hAFSCs in Chpater 3 and Chapter 4, respectively. Their
effects on osteogenesis of hAFSCs were compared in terms of ALP activity and calcium
deposit in this appendix.
Figure A. 1 shows ALP activities of hAFSCs after 7 days treatment by naringin
and curculigoside, respectively. Both of naringin and curculigoside promoted ALP
activity of induced hAFSCs in a dose-dependent manner at the concentration of 1-100
µg/ml, and both of their ALP activities reached the maximum at 100 µg/ml. However,
compared to the control, 100 µg/ml naringin enhanced ALP activity of hAFSCs by 163%,
while curculigoside enhanced ALP activity of hAFSCs by as high as 356%. Therefore,
the promotion effect of curculigoside on osteogenesis of hAFSCs is significantly stronger
than that of naringin.
178
In addition, Figure A. 2 exhibits calcium deposit of hAFSCs treated by naringin
on day 21 and curculigoside on day 14. Both naringin and curculigoside increased
calcium deposit in a dose-dependent manner at the concentration of 1-100 µg/ml and
reached the maxium calcium deposit at 100 µg/ml. However, 100 µg/ml naringin
enhanced calcium content of hAFSCs 239%, while curculigoside enhanced calcium
content of hAFSCs as high as 367%, compared to the control, respectively (naringin
calcium content was measured on day 21 and curculigoside group was measured on day
14, thus they were compared using the increace compared to each own control).
Therefore, curculigoside showed remarkably stronger promotion effect on osteogenic
differentiation of hAFSCs in terms of calcium deposit.
Moreover, as discussed in Chapter 3 and Chapter 4, the responsible signaling
pathways of naringin or curculigoside-enhanced osteogenic differentiation of hAFSCs are
different. Naringin promoted osteogenesis of hAFSCs through both BMP and Wnt
signaling pathways, while curculigoside promoted osteogenesis of hAFSCs via Wnt
pathway.
In summary, both curculigoside and naringin could markedly promote the
osteogenesis of hAFSCs, and curculigoside exhibited stronger effects than naringin in
terms of ALP activity and calcium deposit. However, the price of curculigoside is much
higher than that of naringin, which is a shortcoming for the applications of curculigoside.
179
ALP activity
(nmol (p-nitrophenyl)/min/mg protein)
1.8
↑163%
1.6
1.4
1.2
1
↑74%
↑57%
↑44%
0.8
0.6
0.4
0.2
0
0 µg/ml
1 µg/ml
10 µg/ml
100 µg/ml
200 µg/ml
Naringin concentration
ALP activity
(nmol (p-nitrophenyl)/min/mg protein)
2.5
↑356%
2
↑251%
1.5
1
↑26.3%
↓1.8%
0.5
0
0 µg/ml
1 µg/ml
10 µg/ml
100 µg/ml
200 µg/ml
Curculigoside concentration
Figure A.1 ALP activity of hAFSCs treated by naringin and curculigoside.
180
25
Calcium content (10-10 mmol/cell)
↑239%
20
15
10
↑31%
↑44%
1 µg/ml
10 µg/ml
↑15%
5
0
0 µg/ml
100 µg/ml
200 µg/ml
Naringin concentration
Calcium content (10-10 mmol/cell)
25
↑367%
20
↑243%
15
↑92.7%
10
↑40.5%
5
0
0 µg/ml
1 µg/ml
10 µg/ml
100 µg/ml
200 µg/ml
Curculigoside concentration
Figure A.2 Calcium deposit of hAFSCs treated by naringin and curculigoside.
181
Appendix B
Osteogenic differentiation of naringin treated hAFSCs in 3D
dynamic bioreactors
After 2D static culture of osteogenic differentiation of naringin treated hAFSCs,
the process was scaled up to 3D dynamic bioreactors. Briefly, the differentiation of
hAFSCs was carried out in a fibrous bed bioreactor (FBB), which was made of a 25-mL
spinner flask with a PET matrix affixed on a stainless steel wire mesh around the wall
(Figure B.1). The PET matrix (dimension: 1.2 cm × 9.0 cm × 0.18 cm) had a total
surface area of 4104 cm2. After sterilization, the FBB with the PET matrix was soaked in
10 mL of the medium without naringin (Control) or with 100 µg/mL naringin, inoculated
with 106 hAFSCs, and incubated at 37 °C with agitation at 60 rpm for 21 days in a
humidified atmosphere containing 5% CO2. The culture media were refreshed every 1-3
days according to the metabolic activities. On day 21, the cell number and the calcium
deposition of the cells in the FBB were measured. Figure B. 2 exhibits the cell calcium
deposition in 2D static culture and 3D dynamic bioreactor on day 21. In 2D static culture,
the cells treated with naringin produced significantly more calcium deposition (3.4-fold)
182
than the control group; however, in the 3D dynamic bioreactor, the control group
produced more calcium than in the 2D static culture, while the calcium deposition in 3D
dynamic naringin group was lower than that in the 2D static culture. The calcium content
in control group and naringin group in the 3D dynamic bioreactors turned out to be very
close. However, the FBB promoted osteogenic differentiation of hAFSCs in term of
calcium deposition when no naringin was used.
183
Figure B.1. A fibrous bed bioreactor modified from a spinner flask with a PET matrix
around the wall used for osteogenic differentiation of hAFSCs.
184
Calcium content (10^(-10) mmol/cell)
30
2D static culture
25
3D dynamic bioreactor
20
15
10
5
0
0 µg/ml
100 µg/ml
Naringin concentration
Figure B. 2. Calcium deposition of naringin treated hAFSCs in 2D static culture and 3D
dynamic bioreactor.
185
Appendix C
Extracts of Chinese herb combinations on the proliferation of
hAFSCs
Panax notoginseng (PN), Rhizoma Atractylodis macrocephalae (RAM), Rhizoma
chuanxiong (RC) and Ganoderma lucidum spores (GLS) are four traditional Chinese
herbal medicines from which two extracts of Chinese herbal medicine combinations were
prepared, PN/GLS (PG) and RAM/RC/PN/GLS (RRPG). Briefly, the herbal materials
were cut, smashed into small pieces and combined and added in distilled water (4 g each
herb to 100 ml water), which was then refluxed at 120 °C for 30 min in a 500 ml roundbottom flask equipped with a condenser. The robtained extract solution was centrifuged
at 4,000 rpm for 10 min. The supernatant was filturated throught 0.22 µm membrane and
stored at -20°C until use.
The effects of the two extracts of Chinese herbal medicine combinations on the
proliferation of hAFSCs were tested. 5000 cells were seeded into each well of 48-well
plates and cultured in the growth medium. After 24 h of incubation, the growth medium
186
was changed into PG or RRPG-containing media at a concentration of 0 (Control), 0.005,
0.01, 0.1, and 0.2 g/ml accordingly. Cells were incubated at 37 °C in a humidified 5%
CO2 incubator for 1, 2, 3 or 4 days. After that, the medium was replaced with 500 µl of
10% Alamar Blue (AbD Serotec, Raleigh, NC) solution at 37 °C for 3 h. The
fluorescence of the medium was then monitored in triplicate at 535 nm excitation
wavelength and 590 nm emission wavelength using a GENios Pro plate reader (Tecan,
Research Triangle Park, NC). The fluorescence intensity can be correlated to the cell
number, using a standard calibration curve. The results (Figure C.1) showed that in the
presence of PG or RRPG, the proliferation fold of hAFSCs increased in a dose-dependent
manner in the range of 0.005-0.2 g/mL. For example, on day 4, PG increased the
proliferation by 14%, 22%, 40% and 53% at 0.005, 0.01, 0.1 and 0.2 g/mL, respectively.
The use of 0.2 g/mL PG or RRPG had the most effective promotion and increased the
proliferation by 53% and 42%, respectively, compared to the control on day 4. Thus, PG
and RRPG within the range of 0.005-0.2 g/mL had no cytotoxic effect and stimulated the
proliferation of hAFSCs. In addition, PG showed a little stronger promotion effect on the
proliferation of hAFSCs than RRPG.
The promotion effects of PG and RRPG on the proliferation of hAFSCs should be
further investigated in 3-D dynamic bioreactors.
Acknowledgements
Special thanks to Ding Li for kindly providing PG and RRPG extract solutions.
187
16
14
Proliferation fold
12
Ctrl
PG 0.005
PG 0.01
10
8
PG 0.1
PG 0.2
6
4
2
0
24
72
Time (hour)
96
72
Time (hour)
96
14
12
Proliferation fold
10
8
Ctrl
RRPG 0.005
RRPG 0.01
RRPG 0.1
RRPG 0.2
6
4
2
0
24
Figure C.1 Proliferation of (A) PG and (B) RRPG treated hAFSCs.
188
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