Clemson University
TigerPrints
All Theses
5-2008
Tendon and ligament repair: regeneration and
maturation
Jing Zhao
Clemson University, [email protected]
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Zhao, Jing, "Tendon and ligament repair: regeneration and maturation" (2008). All Theses. Paper 335.
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Theses
TENDON AND LIGAMENT REPAIR: REGENERATION AND MATURATION
A Thesis
Presented to
the Graduate School of
Clemson University
In Partial Fulfillment
of the Requirements for the Degree
Master of Science
Bioengineering
by
Jing Zhao
May 2008
Accepted by:
Dr. Xuejun Wen, Committee Chair
Dr. Zhi Gao
Dr. Ken Webb
ABSTRACT
The thesis constitutes the studies about the two aspects of tendon and ligament
tissue engineering: regeneration and maturation. Injuries to tendon and ligament are
among the most common injuries to the body, particularly in the young and physically
active population. Associated with the problems of incomplete healing and recurrent
injury, these injuries are not only responsible for large health care cost, but also result
in lost work time and individual morbidity. Tissue engineering holds promise in
treating these conditions by replacing the injured tissue with engineered tissue
exhibited similar mechanical and functional characteristics. Collagen plays a central
role in tendon and ligament regeneration, as collagen type I is responsible for more
than 60% and 80% of the dry weigh of tendon and ligament structures, respectively.
The hierarchical organization of collagen type I in bundles confers most of the
mechanical properties of tendons and ligaments. Consequently, tendon or ligament
tissue engineering studies are mainly focused on seeding cells into collagen gels.
However, up to now, no cell-collagen constructs have been able to achieve sufficient
mechanical properties and the complex architecture of the tendon and ligament is
never fully reproduced.
A major cause for low mechanical property of regenerating tendon or ligament
is the slow maturation. The maturation of the engineered tissue is dominated by the
maturation degree of extracellular matrix, such as collagen crosslink density.
ii
To
overcome the slow growth and maturation of tissue engineering grafts, one goal of
this project is to accelerate the tissue maturation using gene therapeutics approach.
Therefore, in the first part of this thesis, two different genes, lysyl oxidase
(LOX) and decorin, were transfected into fibroblasts by retrovirus infection. LOX
initiates the covalent cross-linking of collagen and elastin in the extracellular space by
oxidizing
specific
lysine
residues
in
these
proteins
to
peptidyl
α-aminoadipic-δ-semialdehyde (AAS). These aldehyde residues can spontaneously
condense with vicinal peptidyl aldehydes or with ε-amino groups of peptidyl lysine to
generate the covalent crosslinkages, which stabilizes and insolubilizes polymeric
collagen or elastin fibers in the extracellular matrix. Decorin is considered a key
regulator of matrix assembly because it limits collagen fibril formation and thus
directs tendon and ligament remodeling due to tensile forces. In this study, we found
that the mechanical property of the tissue-engineered grafts was significantly
increased by over-expressing LOX or decorin gene. And decorin over-expression
uniformed the mean diameter of the collagen fibers.
In the second part of this thesis, ECM-based hydrogel system was used to
control the delivery of chemotaxic growth factors, such as hepatocyte growth factor
(HGF), for recruiting endogenous stem cells. This approach was used to attract
endogenous stem cells to the lesion site for tendon or ligament regeneration. In this
study, we found that stem cells could be recruited effectively to the local site where
HGF were released by chemically modified hyaluronic acid (HA) and gelatin (Gtn)
based hydrogels both in vitro and in vivo.
iii
DEDICATION
This thesis is dedicated to my family and friends for their generous support.
iv
ACKNOWLEDGEMENTS
I would like to express my gratitude to all those who gave me the possibility to
complete this thesis. I thank Dr. Xuejun Wen for his continuous guidance and support.
I thank my other committee members, Dr. Zhi Gao and Dr. Ken Webb for their helpful
suggestions and comments throughout my studies. I thank Dr. Ning Zhang, Dr. Hai
Yao, Dr. Qian Kang, Jane Jourdan, and Yuhua Zhang for their help in my experiments.
I thank the Clemson-MUSC Bioengineering department’s faculty and students for
their support, especially our lab members, Xiaowei Li, Yongzhi Qiu, Vince Beachely,
and Xiaoyan Liu.
v
TABLE OF CONTENTS
Page
TITLE PAGE .............................................................................................................
i
ABSTRACT ...............................................................................................................
ii
DEDICATION ...........................................................................................................
iv
ACKNOWLEDGEMENTS .......................................................................................
v
LIST OF FIGURES ...................................................................................................
viii
CHAPTER
1.
THESIS ROAD MAP ................................................................................
1
2.
BACKGROUND .......................................................................................
3
Tendon Structure ..................................................................................
Ligament Structure ...............................................................................
Tendon and Ligament Injuries .............................................................
Tendon and Ligament Healing .............................................................
Traditional Treatments For Tendon And Ligament Injuries ................
Tissue Engineering.................................................................................
Tissue Engineering Treatment For Tendon and Ligament Repair .........
Tendon and Ligament Maturation ........................................................
Gene Therapy .......................................................................................
Stem Cells Recruitment .......................................................................
References ............................................................................................
3
7
9
10
13
15
17
24
32
35
38
RESEARCH OBJECTIVES ......................................................................
51
3.
Table of Contents (Continued)
Page
4.
ACCELERATION OF THE MATURATION OF
TISSUE ENGINEERED GRAFTS BY GENE
THERAPY FOR TENDON OR LIGAMENT
REPAIR ...................................................................................................
53
Abstract ................................................................................................
Introduction ..........................................................................................
Materials and Methods .........................................................................
Results and Discussion ........................................................................
Conclusion .........................................................................................
References ............................................................................................
53
54
58
68
78
78
5 RECRUITMENT OF ENDOGENOUS STEM CELLS
FOR TISSUE REPAIR ................................................................................. 81
Abstract.................................................................................................. 81
Introduction .......................................................................................... 82
Materials and Methods ......................................................................... 85
Results .................................................................................................. 92
Discussion ............................................................................................ 102
Conclusion ........................................................................................... 105
References ............................................................................................ 106
6.
FUTURE DIRECTIONS ...........................................................................
vii
110
LIST OF FIGURES
Figure
Page
2.1
Tendon structure from collagen fibril to the entire tendon ......................
5
2.2
Tendon stress-strain curve..........................................................................
7
2.3
Tensile strength of the injured tendons during wound healing process ...
12
2.4
Concept of tissue engineering ..................................................................
17
2.5
Schematic diagram showing the different posttranslational
modifications and assembly of type I collagen into fibrils ..................
26
Structure of the carbonyl cofactor of lysyl oxidase,
lysyl tyrosine quinone ............................................................................
30
2.7
Mechanism of action of lysyl oxidase .....................................................
30
4.1
Plasmid contruction .................................................................................
60
4.2
Retrovirus production and MEF cells infection .......................................
62
4.3
Silicon molds ...........................................................................................
65
4.4
LOX and decorin expression in MEF ........................................................
69
4.5
Force-displacement curves of the collagen-cell grafts ............................
70
4.6
Young’s Modulus of collagen-MEF constructs .........................................
70
4.7
TEM result for in vitro cell culture ..........................................................
72
4.8
Cells aligned after culture on biodegradable polyurethane nanofibers ....
74
4.9
Force-displacement curves of the nanofiber-cell grafts ...........................
75
4.10
Young’s Modulus of the grafts retrieved from the animals .....................
75
2.6
viii
4.11
Sirius red staining of the grafts retrieved from the animals .....................
78
5.1
Cumulative in vitro HGF release from HA:Gtn
and HA:Gtn:HP hydrogels......................................................................
93
5.2
8μm pore size transwell ...........................................................................
96
5.3
MSCs across the 8μm pores to the lower surface of the filter ...................
96
5.4
The number of MSCs across the 8μm pores
to the lower surface of the filter .............................................................
97
5.5
MSCs migrated into the hydrogels on the lower compartment. ................
98
5.6
The number of MSCs migrated into the hydrogels on the
lower compartment ...............................................................................
99
5.7
SEM images showing morphology of porous scaffold
with pore sizes 100-200μm in diameter ............................................... 100
5.8
Cells migrated into the scaffolds after 1-week implantation ..................... 102
ix
CHAPTER 1
THESIS ROADMAP
The thesis constitutes the studies about the two aspects of tendon and ligament
tissue engineering: regeneration and maturation. Injuries to tendon and ligament are
among the most common injuries to the body, particularly in the young and physically
active population. Associated with the problems of incomplete healing and recurrent
injury, these injuries are not only responsible for large health care cost, but also result
in lost work time and individual morbidity. Tissue engineering holds promise in
treating these conditions by replacing the injured tissue with engineered tissue
exhibited similar mechanical and functional characteristics. Collagen plays a central
role in tendon and ligament regeneration, as collagen type I is responsible for more
than 60% and 80% of the dry weigh of tendon and ligament structures, respectively.
The hierarchical organization of collagen type I in bundles confers most of the
mechanical properties of tendons and ligaments. Consequently, tendon or ligament
tissue engineering studies are mainly focused on seeding cells into collagen gels.
However, up to now, no cell-collagen constructs have been able to achieve sufficient
mechanical properties and the complex architecture of the tendon and ligament is
never fully reproduced.
A major cause for low mechanical property of regenerating tendon or ligament
is the slow maturation. The maturation of the engineered tissue is dominated by the
maturation degree of extracellular matrix, such as collagen crosslink density.
To
overcome the slow growth and maturation of tissue engineering grafts, one goal of
this project is to accelerate the tissue maturation using gene therapeutics approach.
Therefore, in the first part of this thesis, two different genes, lysyl oxidase
(LOX) and decorin, were transfected into fibroblasts by retrovirus infection. LOX
initiates the covalent cross-linking of collagen and elastin in the extracellular space by
oxidizing
specific
lysine
residues
in
these
proteins
to
peptidyl
α-aminoadipic-δ-semialdehyde (AAS). These aldehyde residues can spontaneously
condense with vicinal peptidyl aldehydes or with ε-amino groups of peptidyl lysine to
generate the covalent crosslinkages, which stabilizes and insolubilizes polymeric
collagen or elastin fibers in the extracellular matrix. Decorin is considered a key
regulator of matrix assembly because it limits collagen fibril formation and thus
directs tendon and ligament remodeling due to tensile forces. In this study, we found
that the mechanical property of the tissue-engineered grafts was significantly
increased by over-expressing LOX or decorin gene. And decorin over-expression
uniformed the mean diameter of the collagen fibers.
In the second part of this thesis, ECM-based hydrogel system was used to
control the delivery of chemotaxic growth factors, such as hepatocyte growth factor
(HGF), for recruiting endogenous stem cells. This approach was used to attract
endogenous stem cells to the lesion site for tendon or ligament regeneration. In this
study, we found that stem cells could be recruited effectively to the local site where
HGF were released by chemically modified hyaluronic acid (HA) and gelatin (Gtn)
based hydrogels both in vitro and in vivo.
2
CHAPTER 2
BACKGROUND
Tendon Structure
Tendons are anatomic structures that connect muscles to bones and transmit
the forces created in the muscles to bones making joint movements possible. Healthy
tendons, which exhibit high mechanical strength and good flexibility, are brilliant
white in color and fibro-elastic in texture. Tendons vary in form. They can be rounded
cords, strap-like bands or flattened ribbons [1]. Tendons consist of collagens,
proteoglycans, glycoproteins, water and cells.
Tendon cells
Tendon fibroblasts, including tenoblasts and tenocytes, comprise about
90-95% of the cellular elements of the tendon. The other 5-10% includes the
chondrocytes at the pressure and insertion sites, the synovial cells of the tendon sheath
on the tendon surface, the vascular cells, such as capillary endothelial cells and
smooth muscle cells of the arterioles [2, 3].
The newborn tendon has a very high cell-to-matrix ratio. Tenoblasts are
immature tendon cells, and they are spindle-shaped, with numerous cytoplasmic
organelles reflecting their high metabolic activity [1]. When the cell-to-matrix ratio
gradually decreases with aging, the tenoblasts become elongated and transform into
tenocytes. Tenoblasts and tenocytes align in rows between collagen fiber bundles, and
they are responsible for synthesizing extracellular matrix proteins, producing an
organized collagen matrix and remodeling it during tendon healing [4].
Tendon extracellular matrix
Tendons are rich in collagens, with the most abundant tendon component
being type I collagen, which constitutes about 60% of the dry mass of the tendon and
about 95% of the total collagen [4-6]. The remaining 5% consists of type III and V
collagen. Type III collagen has major role in tendon early repair [7] and type V
collagen copolymerizes with type I collagen to regulate tendon fibril diameter during
fibrillogenesis [8] and aging [9].
Collagen is arranged in hierarchical levels of increasing complexity, beginning
with tropocollagen (a triple-helix polypeptide chain), which unites into fibrils; fibers
(primary bundles); fascicles (secondary bundles); tertiary bundles; and the tendon
itself [10] (Figure 2.1). Soluble tropocollagen molecules form cross-links to create
insoluble collagen molecules, which aggregate to form collagen fibrils. A bunch of
collagen fibrils form a collagen fiber, which is the basic unit of a tendon. Collagen
fibers are bound by endotenons, a thin layer of connective tissue that contains blood
vessels, lymphatics and nerves [11]. Fascicles are composed of fiber bundles, and
bundles of fascicles are enclosed by the epitenon, a fine, loose connective-tissue
sheath containing the vascular, lymphatic, and nerve supply to the tendon [11].
4
Tendon
Primary
fiber bundle
Collagen (subfascicle)
Collagen fiber
fibril
Tertiary fiber
bundle
Secondary
fiber bundle
(fascicle)
Endotenon
Epitenon
Figure 2.1 Tendon structure from collagen fibril to the entire tendon.
Besides collagens, tendons also contain proteoglycans and glycoproteins.
There are many proteoglycans, including aggrecan and decorin [12]. Aggrecan holds
water within the fibrocartilage and resists compression [13]. Decorin, which located in
the tropocollagen interstices, is believed to regulate fibril diameter and to provide
fibril-to-fibril and fibril-to-proteoglycan binding[14]. Glycoproteins present in tendon
extracellular matrix include tenascin-C and fibronectin. Tenascin-C contributes to the
mechanical stability of the extracellular matrix through its interaction with collagen
fibrils [15]. Fibronectin has role in wound healing, its synthesis increases at wound
healing [16].
Elastin, which composes about 2% of the dry weight of the tendon, is another
component of tendon extracellular matrix. Elastic fibers are scarcely present in human
tendons, and it is reported that elastic fibers were actually demonstrable in only 10%
5
of healthy human tendons [17]. The function of elastic fiber is not entirely clear, but
they may contribute primarily to the recovery of the wavy collagen fiber configuration
after tendinous stretch [18].
Tendon mechanical properties
Tendons transmit force generated by muscle to bone, and act as a buffer by
absorbing external forces to limit muscle damage. Tendons exhibit high mechanical
strength, good flexibility, and an optimal level of elasticity to perform their unique
role [19]. Tendons are viscoelastic tissues, and they display stress relaxation and creep
[1].
A typical tendon stress-strain curve is shown to demonstrate the behavior of
tendon (Figure 2.2). The mechanical behavior of tendon collagen is dependent on the
number and types of intra- and inter- molecular bonds [20]. At rest, collagen fibers
and fibrils display a crimped configuration. The initial concave portion of the curve is
called toe region. This toe region, where tendon is strained up to 2%, represents the
stretching-out the crimp pattern [18]. In the linear region of the stress-strain curve,
where the tendon is stretched less than 4%, collagen fibers lose their crimp pattern and
the fibers become more parallel [21]. In this region, the tendon behaves in an elastic
fashion, and returns to its original length when unloaded. The slope of this linear
region is referred to as the Young’s module of the tendon [4]. When the tendon is
stretched over 4%, microscopic failure occurs. When the tendon is stretched over
8-10%, macroscopic failure occurs. And further stretch causes tendon rupture [18].
6
Stress
(N/mm2)
Rupture
Physiological
range
n
gio
re
r
ea
Lin
‘toe’
region
0
Strain
(%)
2
Crimped Straighten
fibers
fibers
4
6
8
Microscopic
failure
Macroscopic
failure
Figure 2.2 Tendon stress-strain curve showing the initial non-linear toe region followed by a linear
region up to a failure region.
Tendons are viscoelastic and sensitive to different strain rates. Tendons are
more deformable at low strain rates. Therefore, the tendons absorb more energy, but
are less effective in transferring loads. At high strain rates, tendons become less
deformable with a high degree of stiffness and are more effective in moving large
loads [10]. Furthermore, tendons are at the highest risk for rupture if tension is applied
quickly and obliquely, and highest forces are seen during eccentric muscle contraction
[20].
Ligament Structure
Ligament is fibrous tissue that connects bones to other bones. Ligament is
composed of closely packed collagen fiber bundles organized mostly in a parallel
7
configuration along the length of the tissue to resist tensile loads [22]. So, ligament
and tendon have a similar hierarchical structure that affects their mechanical behavior.
Ligament cells
Fibroblast is the major cell type of ligament. The cells are relatively few in
number and represent a small percentage of the total ligament volume. Fibroblasts are
not only responsible for collagen synthesis but also for collagen renewal process,
which includes enzymatically breaking down and removing old collagen [23].
Ligament extracellular matrix
Ligaments are approximately two-third water and one-third solid. Water is
responsible for contributing to cellular function and viscoelastic behavior. The solid
components of ligaments are principally collagen. The most abundant ligament
component is type I collagen, which constitutes about 80% of the dry mass of the
ligament and about 85% of the total collagen [24]. The rest collagens in ligament
include collagen type III, V, VI, XI, and XIV. Type III and V collagen make up 8% of
and 12% of the dry weight of the ligament, respectively, and they have been shown to
be involved in regulating collagen fibril diameter [25]. Besides collagen, ligaments
also contain proteoglycans, elastin, glycoproteins such as actin, laminin and integrins
[24].
Collagen is arranged in hierarchical levels in ligament, which is similar as in
tendon. Briefly, collagen molecules assemble sequentially into microfibrils, subfibrils,
8
and fibrils (20-150nm in diameter) before forming fibers, which are 1-20μm in
diameter [23]. Crosslinked fibers further make up a subfascicular unit, which is
surrounded by a loose band of connective tissue called endotenon. The diameter of
subfascicular units is 100-250μm. Three to twenty subfasciculi subsequently form a
fasciculus (from 250μm to several millimeters in diameter)[23]. The fasciculus are
surrounded by epitenon, which supports the neurovascular elements of the ligament
[23].
Tendon and ligament injuries
Tendon injuries are among the most common injuries to the body, particularly
in the young and physically active population. The most common tendon disorders are
observed in the Achilles tendon (100,000 per year) [26], in the rotator cuff in the
shoulder (51,000 per year) [27], and patellar tendon donor autograft injuries in the
knee (42,000 per year) [22].
Tendon injuries can be acute or chronic, and are caused by intrinsic or
extrinsic factors, either alone or in combination. Acute injuries typically arise from
trauma, whereas chronic injuries often occur after repetitive subfailure mechanical
events followed by inflammation [28, 29]. Chronic injuries can be associated with
inflammation (tendonitis), without inflammation (tendinosis), or involve the
surrounding tissue (peritendinitis) [19].
9
Acute tears of rotator cuff tendons commonly occur in athletes participating in
overhead sports [30], whereas chronic degeneration and tearing tend to occur in older
more sedentary subjects [27]. The Achilles tendon is often injured traumatically, but it
is also frequently involved in chronic injuries. Acute Achilles tendon injury and pain
usually occur in active individuals, when the tendon is subjected to high or unusual
loads. In contrast to acute injuries, Achilles tendinopathies are caused by repeated
microinjuries coupled with a degenerative or failed healing response [31].
Ligaments are most often torn in traumatic joint injuries that can result in
either partial or complete ligament discontinuities. For example, over 100,000 anterior
cruciate ligament (ACL) ruptures estimated to occur each year in the United
States[32]. All of these tendon and ligament disorders can bring significant morbidity
and decrease the quality of life.
Tendon and ligament healing
When tendons or ligaments are injured, the body initiates a process of healing.
Tendon and ligament healing can be largely divided into three overlapping phases: the
inflammatory, repairing, and remodeling phases [33].
The initial inflammatory phase lasts about 24 hours. In this phase, erythrocytes,
platelets and inflammatory cells (neutrophils, monocytes, macrophages) migrate to the
wound site and clean the site of necrotic materials by phagocytosis. In the mean time,
10
these cells release vasoactive and chemotactic factors, which recruit fibroblasts to
begin collagen synthesis and deposition [4]. During this phase, there is an increase in
fibronectin, glycosaminoglycan, water, and collagen type III content, which
collectively stabilize the newly formed extracellular matrix [10]. A few days after the
injury, the repairing phase begins. This phase lasts a few weeks. In this phase, the
predominant cell types are fibroblasts along with a smaller number of macrophages
and mast cells [34]. Fibroblasts synthesize abundant collagen and other extracellular
matrix components such as proteoglycans and deposit them to the wound site.
Synthesis of type III collagen peaks during this stage, water content and
glycosaminoglycan concentrations remain high [1]. About 6 weeks later, the
remodeling phase begins. In this phase, fibroblasts decrease in size and slow their
matrix synthesis, and collagen fibers begin to orient themselves longitudinally along
the long axis of the tendon or ligament. The repair tissue changes from cellular to
fibrous, which again changes to scar-like tendon tissue after 10 weeks [1]. When the
scar enters maturation, type III to type I collagen ratio, collagen crosslinks, and
glycosaminoglycan, and water concentrations approach normal levels. During the
later remodeling phase, covalent bonding between collagen fibers increases, which
results in repaired tissue with higher stiffness and tensile strength [4]. Although the
tensile strength of the healing tissue improves over time, it does not reach the levels of
uninjured, normal tissue (Figure 2.3) [34]. Despite remodeling, the biochemical and
mechanical properties of healed tissue never match those of intact tendon. In normal
tendon or ligament, the collagen fibers are all arranged in a vertical alignment to best
11
withstand the force that the tendon or ligament would be loaded; while in repaired
scar tissue, the collagen fibers are randomly arranged in many directions. Also the
healed tissue is unable to recover normal crosslink density, it may be because of the
alteration in the proportion of collagen types, proteoglycans and other unknown
factors within the wound environment [33]. Mechanically, healed tendons or ligament
do not achieve normal tissue failure force in the long term. Butler et al reviewed that
1.5 months after creating a defect injury in the patellar tendon, the repair tissue
achieves 7-9% of normal failure force and by 3 and 6 months after injury, repair tissue
failure force changed to 11% and 8% of normal respectively [31].
Inflammation, Repairing,
hemostasis proliferation
Remodeling,
maturation
Healthy tissue
Tensile
strength
Injured tissue
Time
Days
Weeks
Months
Years
Figure 2.3 Tensile strength of the injured tendons during wound healing process.
12
Traditional Treatments for tendon and ligament injuries
Because tendons and ligaments heal poorly, there are many different types of
treatment used in tendon and ligament disorders, such as physical therapy, reduction
of inflammation, restoration of flexibility and surgical repair [27, 35-37]. Some
current treatments and some therapies in development will be introduced in the
following paragraphs.
Clinical used treatments
Sutures There are a variety of different suture techniques used clinically, each with
their preferred suture material that may be employed at the discretion of the surgeon.
Some of these techniques include Lin locking [38], Savage [39], Kessler [40],
modified Kessler [39], and epitenon suture [41] method. It has been shown that the
repair strength is directly proportional to the number of strands crossing a repair site,
ruptures usually occur at the knots in the suture, and epitenon sutures increase repair
strength over core sutures alone [34, 41].
Eccentric exercise therapy Eccentric strength training is particularly effective in
treating tendinopathies and helps promote the formation of new collagen [42].
Eccentric contraction involves the lengthening of muscle fibers as the muscle
contracts, preferentially loading the tendon [43]. Eccentric exercise has proved
beneficial in Achilles tendinosus and patellar tendinosus and may be helpful in other
tendinopathies [43].
13
NSAIDs Nonsteroidal anti-inflammatory drugs (NSAIDs) effectively relieve
tendinopathy pain and may offer additional benefit in acute inflammatory tendonitis
because of their anti-inflammatory properties. However, because the majority of
chronic tendinopathies are not inflammatory, few data exist to support the use of
NSAIDs over analgesics without anti-inflammatory effects [43].
Other therapies include cryotherapy (reduction of acute inflammation and decrease in
cell metabolism [44]), heat treatment (stimulation of cell activity and increase of
blood flow [44]), and physiotherapy (same as heat treatment).
Treatments under development
Laser treatment Low-level laser therapy (LLLT) is using of low-power lasers and
superluminous diodes for the treatment of a variety of medical conditions, and it now
has been known as a possible treatment for tendon and ligament injuries. LLLT can
enhance ATP production, enhance cell function and increase protein synthesis [45].
LLLT has also been shown to have positive effects on reduction of inflammation,
increase of collagen synthesis, and angiogenesis [45]. There have been a few studies
regarding effects of LLLT on tendon and ligament repair [45, 46].
Corticosteroid injection Corticosteroids may reduce inflammation and inhibit protein
synthesis [44]. Injected corticosteroids may be more effective than oral NSAIDs for
relief in the acute phase of tendon pain, but they do not tend to alter long-term
outcomes. The effects of peritendinous corticosteroid injections are unknown, but they
should be used with some caution [43]. Because the role of inflammation in
14
tendinopathies is unclear, corticosteroids may serve only to inhibit healing and reduce
the tensile strength of the tissue, predisposing to spontaneous rupture [43].
Other treatments under development include shock-wave therapy (stimulate
tenocytes for repair), nitroglycerin patches (enhance collagen synthesis), and
sclerosant injections (block tendon blood flow) [44].
There are many drawbacks of existing treatments. One principal drawback is
that the characteristic response to injury is for fibroplasias to occur, which inevitable
leads to scar tissue in the tendon or ligament. Although remodeling of the scar tissue
occurs over time, the subsequent tissue is not normal and, in particular, has less
compliance and functionality than the original tendon and ligament matrix [47].
As a result of the deficiencies of current treatment, there is great interest in
investigating the potential for tissue engineering in tendon and ligament injuries.
Tissue Engineering
Many people suffer from tendon and ligament diseases each year. For example,
100,000 people suffer from Achilles tendon disorders every year, 51,000 people suffer
from rotator cuff tendon disorder each year, and over 100,000 anterior cruciate
ligament (ACL) ruptures estimated to occur each year in the United States [32].
Associated with the problems of incomplete healing and recurrent injury, these
injuries are not only responsible for large health care costs, but also result in lost work
15
time and individual morbidity [48]. The rate at which these injuries have increased
and the degree to which costs have as well. For example, between 1992 and 1999,
total knee replacement increased over 77.8%, at a compounded annual rate of 15.5%
[49]. Surgical repair, artificial prostheses, mechanical devices, and both autographs
and allografts continue to be important in medical treatment of certain tendon
disorders. However, there are drawbacks of these traditional treatments for tendon
repair. For example, prosthetic devices have complications such as durability and poor
long-term performance, and allografts have drawbacks such as donor scarcity,
donor-site morbidity, tissue rejection, and disease transmission. Tissue engineering
holds promise in treating these conditions by replacing the injured tissue with
engineered tissue with similar mechanical and functional characteristics [50].
A commonly applied definition of tissue engineering, as stated by Langer and
Vacanti, is “an interdisciplinary field that applies the principles of engineering and life
sciences toward the development of biological substitutes that restore, maintain, or
improve tissue function or a whole organ” [50]. In other words, tissue engineering is
the use of a combination of cells, engineering and materials methods, and suitable
biochemical and physio-chemical factors to improve or replace biological functions.
The basic concept of tissue engineering includes a scaffold that provides an
architecture on which seeded cells can organize and develop into the desired organ or
tissue prior to implantation. The scaffold provides an initial biomechanical profile for
the replacement tissue until the cells produce an adequate extracellular matrix. During
the formation, deposition, and organization of the newly generated matrix, the
16
scaffold is either degraded or metabolized, eventually leaving a vital organ or tissue
that restores, maintains, or improves tissue function [51] (Figure 2.4).
Biomaterials
and scaffolds
d
g
f
c
Bioreactor
e
a
h
Cell source
b
Cell sources
(allogenic or xenogenic)
(autologous)
Figure 2.4 Concept of tissue engineering.
a, Cells are harvested from autologous; b, cells are harvested from allogenic or xenogenic sources; c,
cell proliferation in vitro; d,cells are seeded to a scaffold; e, expanded cells reinjected to the patient
without scaffold material; f, expanded cells reinjected to the patient with scaffold; g, cells are cultured
in bioreactor; h, cultured cells reinjected to the patient.
Tissue Engineering Treatment For Tendon and Ligament Repair
Biomaterials, cell sources, growth factors and bioreactors are four components
for tissue engineering. In this part, these four components are reviewed for tendon and
ligament tissue engineering.
17
Biomaterial and Scaffold
There are several requirements govern the choice of materials for tendon and
ligament tissue engineering. Firstly, the material must be biocompatible and
non-toxicity [52]. The material must not cause abnormal responses in local tissues
surrounding the implant and should not produce toxic or carcinogenic effects.
Secondly, the material should avoid infection, so that the material should be
sterile-able before seeding the cells onto the scaffold, and should remain sterile prior
to surgical implantation. The sterilization technique should be carefully chosen to not
to alter the mechanical and physical properties of the material. Finally, the scaffold
material should provide some mechanical integrity initially to replace the function of
the lost tendon or ligament before the scaffold degrades and is replaced with new
extracellular matrix. Importantly, tendon and ligament have highly hierarchal
organized collagen structure, so the further requirement of the scaffolds for tendon
and ligament repair is that the scaffolds need the similar micro and macro morphology
as tendon and ligament.
As mentioned above, collagen type I constitutes about 60% of the dry mass
of the tendon and 80% of the dry weight of ligament, its hierarchical organization in
bundles confers most of the tendon and ligament mechanical properties, collagen
plays a central role in tendon and ligament tissue engineering. Collagen gels have
been used as model system for in vitro tissue engineering [53, 54], and could be used
to assess the effects of different stimuli on cell proliferation. Also, the collagen gels
have been used as a vehicle to deliver mesenchymal stem cells in defects of the
18
rabbits in vivo [55]. It is also reported that preliminary alignment of the cells in the
gel increases the efficiency of the method [56]. But the collagen constructs are not
able to achieve sufficient mechanical properties, and the complex architecture of the
tendon is never fully reproduced.
Although collagen type I play a central role in tendon and ligament tissue
engineering, collagen type I does not account for all the tissue properties. The
proteoglycans and small leucine rich protein (SLRP) also play important roles in the
organization and mechanical properties of the tendon and ligament structure [57].
Collagen cross-linking has been proposed to enhance early mechanical properties, and
this will be discussed in detail later. Besides collagen, chitin-based [58] and
chitosan-based [59, 60] scaffolds also have been reported used for tendon tissue
engineering.
Except the materials from nature, some synthetic materials have been reported
used for tendon repair. Knitted poly (D,L-lactide-co-glycolide) (PLGA) scaffolds have
been shown to possess good mechanical strength and internal communicating spaces
and have been effectively used for rabbit Achilles tendon repair [61]. Recently,
electrospin technique has been employed to modify the polymers to facilitate cell
attachment, new ECM deposition, and tissue formation for better tendon repair [62].
Cell
Fibroblast: There are two types of tendon fibroblast cells in tendon, tenocytes and
tenoblasts. And fibroblast is major cell type in ligament. Some studies have been done
using fibroblasts for tendon and ligament repair in animal models [63, 64].
19
Stem cells: Stem cells are very attractive cell sources for tendon tissue engineering.
Stem cells are undifferentiated cells and have the ability to self-renew and
differentiate to one or more types of specialized cells. Stem cells can be classed to two
types: embryonic stem (ES) cells and adult stem cells. ES cells are from blastocysts
and fetal tissue. And they have potential in the field of tissue engineering and
regenerative medicine because they have the potential to produce most types of cells
in the body [65-67]. Although the potential of ES cells in tissue engineering is vast,
there are many problems must be overcome. For example, human ES cells may be
contaminated by animal cells or proteins. It is reported that nonhuman protein
expressed by human ES cell lines grown on animal feeder layers [68]. Other problems
such as immunorejection and tumorigenesis are also obstacles for the clinical
applications of ES cells. Currently, no embryonic stem cells-base therapies are
approved for human use.
Adult stem cells are undifferentiated cells, which are found among
differentiated cells in a tissue or organ. Adult stem cells can renew themselves and they
can give rise to mature cell types that have characteristic morphologies and specialized
functions. Adult stem cells are rare, but they are located in various tissue niches
throughout the body, including bone marrow, brain, liver, skin, blood, etc. [69].
Mesenchymal stem cells (MSCs) have attracted much attention because of
their multipotential properties with regard to differentiation, and their possible use for
cell and gene therapy. MSCs reside in diverse host tissues, bone marrow was considered
to be the most accessible and enriched source of MSCs [70]. Also, MSCs have being
20
isolated from various tissues, such as cartilage [71], periostium [72], synovium [73],
synovial fluid [74], muscle [75] and tendons [76]. Fetal tissue [77], placenta, umbilical
blood and vasculature [78] also have been reported to contain MSCs. MSCs retain the
ability to differentiate into a variety of cell types, including adipocytes [79],
chondrocytes [80], osteoblasts [79], skeletal muscle cells [81], and so on. Several
methods are currently available for isolation of MSC based on their physical and
physicochemical characteristics, for example, adherence to plastic or to other
extracellular matrix components. Their wide extend resources, multipotency of
differentiation, ease of isolation and their highly reduced immunoreactivity after
allogenic transfer make MSCs an ideal cell type for tissue repair and regeneration in cell
therapy applications. Several studies have demonstrated the possible use of MSCs in
transplantation for systemic diseases, or implantation to repair local tissue defects.
Several in vivo and in vitro experiments have demonstrated the potential of MSCs
isolated from bone marrow [82] or from adipose tissue[83] for tendon engineering.
Under mechanical stimulation, MSCs differentiated in fibroblasts, aligned themselves in
collagen gel, and helped in vivo tendon regeneration.
Although using MSCs for tissue repair is very promising, there are still issues
related to the application of MSCs for human use. For example, the pluripotency of
MSCs would decrease during in vitro propagation [84, 85]. The cost of in vitro process
for cell propagation and transplantation is high. To overcome this problem, we are
developing a new strategy to use endogenous stem cells for tissue repair. The detail will
be discussed later.
21
Growth Factor
The main growth factors that affect growth and differentiation of ligament and
tendon tissue include basic fibroblast growth factor (bFGF), platelet-derived growth
factor (PDGF), epidermal growth factor (EGF), insulin-like growth factor (IGF)-1,
and members of the family of TGF-β/bone morphogenetic proteins (TGF-β/BMPs).
bFGF is the major mitogenic agent for fibroblasts, and it increases cell
proliferation of rat tenocytes [86]. Injection of bFGF results in a dose-dependent
increase of cell proliferation and collagen type III production [87]. PDGF is also an
important mitogenic agent for fibroblasts [86]. Researches found that enhanced
expression of PDGF-BB in healing ligament led to an initial promotion of
angiogenesis and subsequent enhanced and accelerated matrix deposition[88]. And it
is also reported that PDGF-BB could augment tendon repair in a rotator cuff injury
model [89]. IGF-1 provides some anti-inflammatory functions and also acts as a
chemotactic attractant for endothelial cells[90]. It was reported that via a possible
anti-inflammatory mechanism, IGF-1 could reduce maximum functional deficit and
accelerate recovery after Achilles tendon injury[90].
The growth factors can be delivered locally, but they are easy to be over
loaded. It is important to understand the right time for the administration of growth
factors and their dosage for effective therapies.
Bioreactor
Bioreactors, which provide dynamic loading, are essential for meeting the
complex requirements of in vitro engineering of functional skeletal tissues. It is
22
reported that an increase in cell proliferation, migration and collagen synthesis has
been observed in lacerated chicken tendon cultured in vitro undergoing cyclic tension
after 14 days [91]. Some studies demonstrated that cyclic strain (5%, 1Hz, for 15 or
minutes) could increase TGF-β, bFGF, and PDGF production in human tendon
fibroblasts [92], cyclic mechanical stretching (8%, 0.5HZ, for 4 hours) slightly but
significantly increases tendon proliferation on a microgrooved substrate, and increases
collagen type I and TGF-β expression [93]. And there are some bioreactors designed
specifically for ligament tissue engineering. For example, Altman and his colleagues
designed an advanced bioreactor, which provided the application of multidimensional
mechanical strains (axial tension/compression and torsion) to MSC cultured in a
hydrogel or on a fibrous scaffold [94]. The mechanical stimulation induced cell
alignment in the direction of the resulting force, and fostered the expression of
collagen type I and type III and tenascin C [94].
In conclusion, well combination of the four components (cell, scaffold, growth
factor and bioreactor) is very important for the success of tendon and ligament tissue
engineering.
Two major strategies may be applied for tendon tissue engineering. One is the
in vivo approach of tendon engineering involving in situ delivery of cells,
biomolecules, hydrogels or scaffolds to boost the regeneration and healing process to
deal with the repair of small defects and the induction of tissue self-regeneration [88].
The other is the in vitro tissue engineering involving the growth of tendon-like tissue
23
structure outside of the body and the implantation into the defect. This strategy is
more concerned with larger defects or tissue replacement.
Tendon and Ligament Maturation
The overall shape and function, in terms of flexibility and locomotion, of the
human skeletal system depend on a basic framework of collagen fibers [95]. The
collagen fibers are essentially in extensible and therefore provide mechanical strength
and through that strength confer and maintain form whilst allowing flexibility
between various organs of the body. The collagen fibers of tendons are aligned in
parallel and therefore loaded instantly, permitting maximum transfer of the energy of
muscle contraction to the bone.
Collagen
The collagen molecule, also called tropocollagen, is composed of three
intertwined polypeptide chains. Each of the polypeptides that constitute the collagen
molecule is designated as an α-chain. They intertwine, forming a right-handed triple
helix. Except at the ends of the α-chain, every third amino acid in the chain is a
glycine molecule because glycine is the smallest amino acid. A hydroxyproline
frequently precedes each glycine in the chain, and a proline frequently follows each
glycine in the chain. The glycine, along with proline and hydroxyproline, is essential
24
for the triple-helix conformation. Sugar groups, which are joined to hydroxylysyl
residues, are associated with the helix. Because of these sugar groups, collagen is
properly described as a glycoprotein [96]. The α-chains that constitute the helix are
not all alike, and according to differences in α-chains and differences in molecules
organization and arrangement, as many as 28 different types of collagen have been
described in literature. Tendon and ligament are predominantly fibrous Type I
collagen.
The production of fibrillar collagen involves a series of events within the cells
that leads to the production of procollagen, the precursor of the collagen molecule.
The production of the actual fibril occurs outside of the cell and involves enzymatic
activity at the plasma membrane to produce the collagen molecule, followed by the
assembly of the molecules into fibrils under guidance of the cell. Most of the
synthesis of procollagen is in the rough endoplasmic reticulum (RE). Soluble
procollagens are “packaged” in the Golgi apparatus to be excreted to outside the cell.
Then the procollagen stays close to the cell surface where most of maturation occurs.
The procollagen moves to the exterior of the cell by means of exocytosis of secretory
vesicles. Then the procollagen, on secretion from the cell, is converted to collagen
molecules by procollagen peptidase, which cleaves the uncoiled ends of the molecule.
The aggregated collagen molecules then align to form the final collagen fibrils (Figure
2.5).
25
Figure 2.5 Schematic diagram showing the different posttranslational modifications and assembly of
type I collagen into fibrils [96]. Procollagen consists of a 300-nm-long triple-helical domain flanked by
a trimeric globular C-propeptide domain and a trimeric N-propeptide domain. Procollagen is secreted
from cells and is converted into collagen by the removal of the N- and C-propeptides by procollagen
N-proteinase and procollagen C-proteinase respectively. The collagen generated in the reaction
spontaneously self-assembles into cross-striated fibrils that occur in the extracellular matrix of
connective tissues. The fibrils are stabilized by covalent cross-linking, which is initiated by oxidative
deamination of specific lysine and hydroxylysine residues in collagen by lysyl ocidase.
Collagen maturation
Collagen maturation occurs outside the cell. Once outside the cell, an
enzyme-mediated cross-linking takes place and the triple helical collagen molecules
line up and begin to form fibrils and then fibers. The cross-linking mechanism in the
fibrillar collagens is based upon aldehyde formation from the single telopeptide lysine
or hydroxylysine residue. This step is initiated by a specialized enzyme called lysyl
26
oxidase (LOX), which oxidatively deaminates these residues to promote crosslink
formation. Once LOX deaminates the residues to initiate the cross-linking, the
subsequent reactions of the aldehydes are spontaneous and governed by
post-translational modifications and the structural organization of the collagen. Two
major cross-links are aldimines which are formed by telopeptide lysine-aldehydes
condense with either lysine or hydroxylysine residues in the conserved sequence of
the triple helix and keto-imines which are formed by hydroxylated telopeptide
lysine-aldehyde reacts with the ε-amino group of a helical hydroxylysine [97]. Other
forms of cross-links such as aldol-derived cross-link that is formed by reaction of the
intramolecular aldol condensation product with histidine also exist. The formation of
cross-links in mature collagen creates stability and gives definition to the structure.
The majority of tendons and ligaments contain both the aldimine and the keto-imine
corss-links. The ratio depends upon the particular function of the tissue, in particular
whether there is tension on the tissue.
Besides aldehyde formation, another kind of intermolecular cross-linking of
collagen which will increase with age is glycation. Glycation can affect the properties
of collagen in a number of ways, for example, its ability to form precise
supramolecular aggregates, the alteration of its charge profile and hence its interaction
with cells, and, additionally, glycated collagen can act as an oxidizing agent [97].
Increasing collagen cross-linking of engineered tissue may enhance
mechanical stability. Some strategies have been investigated to increase collagen
cross-linking. Besides enzymatic cross-linking (initiated by LOX) and glycation,
27
which mimic the physiological cross-linking, chemical processing, is another method
for cross-linking. But the chemically cross-linked engineered tissue may subject to
calcification in vivo. And glycation increases a tissue’s platelet aggregation potency
and increases the potential for thrombosis formation. And glycation is widely known
because of the side effect of diabetes when too much glycation occurs [98, 99].
Collagen cross-links formed by the lysyl oxidase (LOX) have no known negative side
effects. In contrast, too few LOX-generated cross-links can lead to decreased tissue
strength.
Lysyl Oxidase
The LOX family consists of five members: LOX and four known family
members called Lysyl oxidase-like (LOXL [100], LOXL2 [101], LOXL3 [102], and
LOXL4 [103]). The LOXLs are isoenzymes: enzymes that have the same catalytic
activity as LOX but different construction. They have related by different functions
including cell growth control, tumor suppression, senescence and chemotaxis [104].
LOXL2, LOXL3, LOXL4 create a subfamily characterized by the presence of four
scavenger receptor cysteine-rich (SRCR) domains [103].
Protein-lysine 6-oxidase (lysyl oxidase, LOX; EC 1.4.3.13), which is
expressed and secreted by fibrogenic cells, is a copper amine oxidase. LOX initiates
the covalent cross-linking of collagen and elastin in the extracellular space by
oxidizing
specific
lysine
residues
in
these
proteins
to
peptidyl
α-aminoadipic-δ-semialdehyde (AAS). These aldehyde residues can spontaneously
28
condense with vicinal peptidyl aldehydes or with ε-amino groups of peptidyl lysine to
generate the covalent crosslinkages, which stabilize and insolubilize polymeric
collagen or elastin fibers in the extracellular matrix [105]. LOX plays a central role in
the morphogenesis and repair of connective tissues of the cardiovascular, respiratory,
skeletal, and other systems of the body.
Lysyl oxidase is synthesized in a precursor form as the 50kDa, N-glycosylated
prolysyl oxidase (pro-LOX). Pro-LOX is secreted to the extracellular space where it
undergoes activation to the 32 kDa, functional catalyst by proteolytic cleavage at a
specific Gly-Asp bond [106].
Lysyl oxidase contains two cosfactors, Cu (II) and a peptide-linked carbonyl
residue, at its active site essential to its catalytic function. The carbonyl cofactor
contains an ortho-quinone function and has been identified as lysine tyrosylquinone
(LTQ), derived by oxidative post-translational modification of a specific tyrosine
residue (tyrosine 349 in rat pro-LOX) to a dihydroxyphenylalanine (DOPA) quinone
residue. Subsequently, the amino group of a specific lysine residue (lysine 314 in rat
pro-LOX) covalently bonds to a ring carbon of peptidely DOPA quinone [107] (Figure
2.6). Copper in lysyl oxidase appears to be involved in the transfer of electrons to and
from oxygen to facilitate the oxidative deamination of targeted peptidyl lysyl groups
in tropocollagen or tropoelastin and to internally catalyze quinone cofactor formation
[108]. LOX catalyzes primary amine oxidation through a ping pong bi ter kinetic
mechanism (Figure 2.7) [109].
29
Figure 2.6 Structure of the carbonyl cofactor of lysyl oxidase, lysyl tyrosine quinone[105].
Figure 2.7 Mechanism of action of lysyl oxidase[105].
30
LOX for many years was unable to be studied because LOX aggregates in
most solutions. When it was found that when urea is added to the solution LOX
became soluble and that urea can be easily separated, LOX studies began intearnest. It
was found that LOX is essential in the functioning of embryonic cardiovascular
system [110], and lungs [110]. It also is important in wound healing and has possible
chemotaxic effects especially to monocytes. LOX has also been shown to enhance
mechanical properties in vascular tissue scaffolds [112].
Decorin
Proteoglycans (PGs) play a crucial role in collagen fibrillogenesis, and
therefore in tendon function. PGs are composed of a core protein to which one or
more GAG chains are covalently attached.
Decorin is the most abundant tendon PG. Decorin is a member of the small
leucine-rich proteoglycan (SLRP) family, a group of secreted proteins that includes
biglycan, fibromodulin, lumican, and keratocan, among others [113-115]. SLRPs play
major roles in collagen fibrillogenesis, growth factor modulation, and direct
regulation of cellular growth [116-118].
Fibroblast, chondrocytes, endothelial cells and smooth muscle cells are
decorin-producing cells. Decorin is composed of three domains: an N-terminal region
which possesses a single chondoitin/dermatan sulfate side chain and a distinct pattern
of Cys residues (CX3CXCX6C), a central region composed of ten leucine-rich repeats
which are believed to be the prime sites of interaction with other proteins, and another
31
Cys-rich C-terminal region [119]. Positioning of collagen in the model is consistent
with other evidence that the central leucine-rich repeats comprise the high-affinity
collagen-binding site [120, 121]. It is believed that decorin likely maps to a narrow
region near the C-terminus of collagen type I, very close to none of its major
intermolecular cross-linking sites [119]. Decorin is considered a key regulator of
matrix assembly because it limits collagen fibril formation and thus directs tendon
remodeling due to tensile forces [122, 123]. The assumed location of decorin on
collagen fibrils in vivo and its ability to retard collagen fibrillogenesis in vitro suggest
that it has a role in the collagen network organization and in the maintenance of tissue
integrity [124]. Thus absence of decorin might affect not only fibril formation but also
fibril stability within tissues.
Gene Therapy
Although the direct application of human recombinant proteins can result in
beneficial effects on the healing process, high dosages and repeated injections of these
proteins are often required due to their relatively short biological half-lives [125]. In
addition, the use of growth factor proteins to promote healing is severely hindered by
the difficulty of ensuring their delivery to a specific injured site [125]. Combined
tissue engineering, gene therapy is a very promising strategy to overcome the
problems of protein therapy. For example, cell isolated from the patients could be
32
genetically engineered to express a given therapeutic gene, and then seeded into a
particular scaffold. After in vitro proliferation, the cells can be transplanted back to
the patient to promote tissue regeneration.
Gene therapy is the science of the transfer of genetic material into individuals
for therapeutic purposes by altering cellular function or structure at the molecular
level. By employing these techniques, genes can be used therapeutically to produce
proteins to treat and potentially cure diseases [126].
Vectors
In order for target cells to produce the protein products of the introduced gene,
the exogenous genetic material must be delivered to the cell’s nucleus. This process of
transfection exists in two classes of vectors: viral and non-viral.
The non-viral vectors are usually easier to produce and result in lower toxicity
and immunogenicity. The use of non-viral vectors can be in the form of injections of
naked DNA (usually plasmids), liposomes, or particle-mediated gene transfer. The
genetic material can be placed in liposomes in order to increase DNA uptake in tissue
culture. But non-viral gene delivery efficiency is hindered by a low transfection rate,
the success associated with non-viral vector usage remains limited.
Currently, viral vectors represent a more efficient method of gene delivery
[127, 128]. The most commonly used viruses are adenovirus, retrovirus,
adeno-associated virus, and herpes simplex virus.
33
Retroviruses are RNA viruses that carry a gene for a reverse transcriptase that
transcribes the viral genetic material into a double-stranded DNA intermediate. This
DNA intermediate is then incorporated into the host DNA allowing the host cell
machinery to produce all the necessary viral components. Because the viral genome is
stably integrated into the host DNA, any modification that has been made will be
passed to all daughter cells that are derived from the transfected cells [129].
In contrast to retroviruses, adenovirus does not integrate its genome into the
host genome. Instead, the adenoviral genome remains in the nucleus as an episomal
element after infection of the host cell. Adenoviral vectors are ease of purification and
concentration, and they could infect various cell types, dividing or non-dividing, at
high efficiency rate [129].
Adeno-associated virus requires a helper virus such as adenovirus or herpes
simplex virus for replication. Adeno-associated virus is actually a member of the
parvovirus family of single-stranded DNA viruses. The small size of the genome
allows for easy manipulation such that shuttle vectors carrying the entire genome have
been constructed. This virus is non-pathogenic and not associated with any known
disease but can infect a wide variety of cells, dividing or non-dividing, although with
varying levels of efficiency [129].
Herpes simplex virus vectors can infect most cells including non-dividing cells,
and they have large carrying capacity and ability to insert expression cassettes into
specific loci of the genome that allows for the construction of vectors able to express
multiple transgenes [130].
34
Strategies
There are two general ways that gene therapy can be performed: a direct in
vivo method and an indirect ex vivo method. The direct method involves transferring
the genetic material into the target somatic cells in vivo. The indirect technique
involves removal of cells from the patient followed by genetic modification of the
cells ex vivo and return of the cells to the patient. The direct method is technically
simpler, while the indirect gene delivery technique is safer because the gene
manipulation takes place under controlled conditions outside the body [32].
Furthermore, this method allows for selection of the cells that express the therapeutic
gene at higher levels [131]. The choice of the strategy depends on the disease to be
treated, the gene to be delivered to treat the disease and the vector used to deliver the
gene.
In the study presented in this thesis, LOX or decorin transfected retrovirus
were used to infect mouse embryonic fibroblasts (MEF) to accelerate collagen
maturation for tendon or ligament regeneration through an indirect ex vivo strategy.
Stem Cells Recruitment
A classic concept of tissue repair holds that inflammatory cells enter the
damaged tissue or damaged tissue secret signaling molecules and signal resident cells
mitosis. Several studies suggest that multipotent stem cells can also contribute to
35
tissue repair after mobilization, migration and engraftment into the damaged tissue
[132]. In addition, circulating immature cells seem to participate in regeneration of
many different tissues [133, 134]. Some models show the role of stem cells in tissue
remodeling, such as for hepatic regeneration [135, 136], muscle regeneration [137,
138] and infracted myocardium [139, 140]. It is not surprising that using endogenous
stem cells have attracted lots of attention because they hold great therapeutic potential
for endogenous tissue repair and tissue engineering.
Stem cell mobilization
Stem cell mobilization is to release of stem cells from their local niche into the
circulation in response to chemotherapy or cytokine stimulation, which was first
documented in the late 1970s and 1980s [141]. Mobilizing cells is a preferable
strategy for clinical use since this may allow higher yield of these cells, faster
engraftment and lower procedural risks compared with traditional stem cell
transplantation approach. Several molecules, such as granulocyte colony-stimulating
factor (G-CSF), granulocyte-macrophage colony-stimulating factor (GM-CSF), stem
cell factor (SCF), and IL-8, IL-12, and stromal cell derived factor-1 (SDF-1)[142] are
shown to be able to mobilize endogenous stem cells into circulation. These molecules
differ in their time frame to achieve mobilization, the type of cells mobilized and
efficiency. G-CSF is the most commonly used agent, usually administrated daily at a
dose of 5-10μg/kg for 5-10 days [141]. It has been shown that G-CSF could mobilize
more CD34+ cells and have less toxicity than any other single agent. GM-CSF as a
36
single agent is used less often today for mobilization than G-CSF, because it
mobilizes somewhat less well than G-CSF and because of relatively higher incidence
of both mild and severe side effects [143]. SDF-1 is an α-chemokine that strongly
chemoattracts hematopoietic stem/progenitor cells (HSPCs) through interaction with
their unique receptor CXCR4. It has become evident that the SDF-1-CXCR4
signaling axis plays an important role in the homing and engraftment of HSPCs in the
bone marrow [144].
Stem cells also migrate to nonhematopoietic organs, such as the liver,
especially during liver injury/inflammation which creates an alarm situation and
transmitting stress signals that mobilize and recruit stem cells as part of organ repair.
Hepatocyte growth factor (HGF)
Hepatocyte growth factor (HGF) is a pleiotropic cytokine of mesenchymal
origin, promoting motility, proliferation, invasion, morphogenesis and survival of a
wide spectrum of cells, namely epithelial and endothelial cells [145]. The coordinated
integration of these processes plays a pivotal role in organ formation during
embryogenesis and tissue homeostasis in adults. In adults, HGF displays in vivo
cytoprotective activity in different cell types of injured organs. It is reported that HGF
is able to enhance mesenchymal stem cells engraftment in injured heart [146]. Many
researches have demonstrated that HGF could activate mesenchymal stem cells
migration in vitro [144, 147, 148].
37
In the study presented in this thesis, HGF was loaded into extracellular matrix
based hydrogels to attract mesenchymal stem cells to the local site for tissue repair.
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50
CHAPTER 3
RESEARCH OBJECTIVES
The preceding literature review has identified the necessity of new strategies
to treat tendon or ligament injuries. Majority of research on tendon and ligament
tissue engineering recently were focused on the cell seeded collagen gels. However,
up to now, no cell-collagen construct has been able to achieve sufficient mechanical
properties. On the other hand, a better cell source need to be identified for tendon and
ligament tissue engineering, due to taking tenocytes out would cause donor-site
morbidity.
According to the current problems in tendon and ligament tissue engineering,
our goal in this study is to enhance tissue-engineered grafts maturation and use
endogenous stem cells for de novo tendon or ligament regeneration and repair. The
specific objectives of this study are:
(1)
To engineer a functional graft with the similar histological
structures and mechanical properties as native tendon or ligament
tissue. Gene therapy would be used as model approach, by which
cells can over-express genes, such as LOX or decorin, to increase
collagen cross-linking, which can enhance collagen maturation, and
optimize collagen fibril formation.
(2)
To attract endogenous stem cells to the lesion site for tendon
regeneration. ECM-based hydrogel system would be used to
control the delivery of chemotaxic growth factors, such as
hepatocyte growth factor (HGF), for recruiting mesenchymal stem
cells into the lesion site.
52
PART I: CHAPTER 4
ACCELERATION OF THE MATURATIONOF TISSUE ENGINEERED GRAFTS
BY GENE THERAPY FOR TENDON OR LIGAMENT REPAIR
Abstract
Despite significant progress of tissue engineering technologies has been
achieved both in academic labs and in industries, a number of difficulties are still
present which slow down the commercial process of tissue engineered products. For
tendon and ligament repair by means of tissue engineering, one major difficulty is
how to gain functional grafts with the similar mechanical properties as the native
tissues. To overcome the poor performance of engineered tendon tissue grafts on
histological structures and mechanical properties, we try to accelerate the maturation
of engineered grafts and optimize the graft structure using gene therapy. The
maturation of tendon tissue is dominated by the amount and the crosslinkage of the
extracellular matrices, such as collagen, elastin, glycosaminoglycans, and so on.
Maturation of collagen and elastin is controlled by the degree of crosslinking in these
molecules. Crosslinking can occur in various ways, but in vivo most crosslinkings are
mediated by lysyl oxidase (LOX), which recognizes different binding sites that are
similar on the type I, II, and III collagens and elastin. On the other hand, decorin is
considered a key regulator of matrix assembly, and it plays a major role in the
collagen network organization and in the maintenance of tissue integrity. In this study,
Infected MEF cells that expressed LOX or decorin were utilized as a model system to
test our strategies. The results demonstrated that LOX and decorin significantly
promote tissue maturation indicated by showing the significant increase in the
strength of the engineered grafts in vitro and in vivo.
Introduction
Tendons and ligaments are anatomic structures that connect muscles to bones
or bone to bones and transmit the forces to bones making joint movements possible.
Healthy tendons, which exhibit high mechanical strength and good flexibility, are
brilliant white in color and fibro-elastic in texture. Tendon and ligament injuries are
among the most common injuries to the body, particularly in the young and physically
active population. The most common tendon disorders are observed in the Achilles
tendon (100,000 per year) [1], in the rotator cuff in the shoulder (51,000 per year) [2],
and patellar tendon donor autograft injuries in the knee (42,000 per year) [3].
Ligaments are most often torn in traumatic joint injuries that can result in either
partial or complete ligament discontinuities. For example, over 100,000 anterior
cruciate ligament (ACL) ruptures estimated to occur each year in the United States[4].
All of these tendon and ligament disorders can bring significant morbidity and
decrease the quality of life.
Treatment of tendon and ligament disorders has traditionally involved
reduction of inflammation, restoration of flexibility and surgical repair. When
inflammation (tendinitis) and degeneration (teninosis) cannot be resolved, primary
54
repairs, autografts [5], allografts [6], and xenografts [7], and resorbable synthetic
biomaterials [7-9] have been attempted, but with varying success. Recently, tissue
engineering approaches have been evaluated to treat tendon disorders, based on the
concept that an improved repair outcome will achieved by combing principles of
engineering and biology Thus the focus of developing new treatment strategies for
tissue repair has shifted from transplantation and replacement to biological
regeneration.
Recent advances in sophisticated scaffold fabrication, controlled release of
biomolecules, stem cell biology, and bioreactor development, have raised extensive
interests in tissue engineering and regenerative medicine-based tissue regeneration
strategies in academic researchers, clinicians, and biotechnology industries. To date,
fundamental techniques for tissue engineering both in vitro and in vivo, such as
scaffold fabrication, controlled release of growth factors and genes, manipulation of
stem cells and precursors cells, and dynamic bioreactors, have been established for
clinical translation of tissue engineering strategies.
Despite significant progress has been achieved both in academic research and
in industry, a number of difficulties slow down the commercial process. One major
difficulty is how to gain a functional graft the similar mechanical properties as native
tissues. To overcome the poor performance of engineered tendon or ligament tissue
grafts on histological structures and mechanical properties, we try to accelerate the
maturation of engineered grafts and optimize the graft structure using gene therapy.
The maturation of tendon tissue is dominated by the maturation degree of
55
extracellular matrix. The maturation of extracellular matrix is proceeded by the
cross-linking of various components in extracellular matrix, such as collagen, elastin,
glycosaminoglycans, and so on. Cross-linking of ECM is also an effective approach to
improve the strength of constructs for tissue-engineering applications. Since collagen,
which constitutes more than 60% of the dry mass of tendon, is the major component
of ECM of tendon. The most abundant ligament component is type I collagen as well,
which constitutes about 80% of the dry mass of the ligament and about 85% of the
total collagen [10]. The rest collagens in ligament include collagen type III, V, VI, XI,
and XIV. Type III and V collagen make up 8% of and 12% of the dry weight of the
ligament, respectively, and they have been shown to be involved in regulating
collagen fibril diameter [11]. In our study, for tendon or ligament tissue engineering,
the maturation of collagen type I was focused on.
The collagen fibers of tendons and ligaments are aligned in a parallel and
therefore loaded instantly, permitting maximum transfer of the energy of muscle
contraction to the bone or bone to bone. Collagen has triple-helix structure. Collagen
synthesis occurs in the endoplasmic reticulum, packaged in the Golgi apparatus and
excreted to extracellular environment. Maturation/cross-linking of collagen occurs
outside the cell. The formation of cross-links in mature collagen creates stability and
gives definition to the structure. The major ways to form cross-links in vitro are
nonenzymatic cross-linking and enzyme cross-linking. Nonenzymatic cross-linking
includes glutaraldehyde fixation, sugar-mediated cross-linking and so on. The main
drawback to glutaradehyde derived crosslinks is that glutaraldehyde-fixed tissues are
56
subject to calcification in vivo. Sugar-mediated cross-link formation, also called
glycation can form covalent bonds between collagen molecules and increase tissue
stiffness. Glycation increases a tissue’s platelet aggregation potency and increase the
potential for thrombosis formation. Collagen cross-links can be mediated by secretory
enzymes such as lysyl oxidase (LOX). In fact, too few LOX-generated cross-links is
reported to associate with decreased tissue strength. Protein-lysine 6-oxidase (lysyl
oxidase, LOX; EC 1.4.3.13), which is expressed and secreted by fibrogenic cells, is a
copper amine oxidase. LOX initiates the covalent cross-linking of collagen and elastin
in the extracellular space by oxidizing specific lysine residues in these proteins to
peptidyl α-aminoadipic-δ-semialdehyde (AAS). These aldehyde residues can
spontaneously condense with vicinal peptidyl aldehydes or with ε-amino groups of
peptidyl lysine to generate the covalent crosslinkages, which stabilize and insolubilize
polymeric collagen or elastin fibres in the extracellular matrix [12]. LOX plays a
central role in the morphogenesis and repair of connective tissues of the
cardiovascular, respiratory, skeletal, and other systems of the body.
Besides collagen, proteoglycans (PGs) play a crucial role in collagen
fibrillogenesis, and therefore in tendon function. Decorin is the most abundant tendon
PG. Decorin is considered a key regulator of matrix assembly because it limits
collagen fibril formation and thus directs tendon remodeling due to tensile forces [13,
14]. The assumed location of decorin on collagen fibrils in vivo and its ability to
retard collagen fibrillogenesis in vitro suggest that it has a role in the collagen
network organization and in the maintenance of tissue integrity[15]. Thus absence of
57
decorin might affect not only fibril formation but also fibril stability within tissues.
In our study, we used LOX or decorin transfected retrovirus to infect mouse
embryonic fibroblasts (MEF) to accelerate collagen maturation for tendon repair.
Materials and Methods
Materials
LOX plasmid was a gift from Dr. Ben Fogelgren. Decorin plasmid was
purchased from ATCC. DMEM media, FBS, L-glutamine solution, non-essential
amino acids solution were purchased from Invitrogen. Restriction endonucleases used
for plasmid construction were purchased from New England Biolabs.
Derivation of mouse embryonic fibroblasts (MEF)
CF-1 strain mice at 12.5 days gestation were anesthetized. Mouse was placed
belly up in sterile tissue culture hood. The belly skin was cut using sterilized
instruments to expose the peritoneum. And then the peritoneal wall was cut and the
uterine horns were exposed. Uterine horns were removed and put to a new petri dish
after washing. Then embryonic sacs were open and embryos were released using
forceps and scissors. Visceral tissue was separated from embryos, and the embryos
were put in a new plate. Mince tissue with dissecting scissors into grain sized pieces.
Add 2ml trypsin, and mince for an additional few minutes until pieces are further
reduced in size. Add an additional 5ml trypsin and place dish into incubator for 20-30
58
minutes. And then vigorously pipet mixture up and down. Add about 20ml MEF
culture media (DMEM media with 10% FBS, 1% 200mM L-glutamine solution and
1% non-essential amino acids solution) and transfer contents to a sterile 50ml plastic
conical tube. Divide the mixture to several T75 flasks. Incubate the flasks containing
the minced tissue mixture in a 37oC tissue culture incubator overnight. Continue to
culture the flask until at least 90% of the flask surface is covered with cells.
Plasmid construction
Plasmid pENTR 1A (Invitrogen) was used to get the entry clones containing
LOX or Decorin genes. To construct pENTR-LOX plasmid, EcoRI/PmeI double
digested LOX fragment was ligated with EcoRI/EcoR1V digested pENTR 1A
fragment. To construct pENTR-Decorin plasmid, BamH1/BglII digested Decorin
fragment was ligated with BamH1/EcoRV digested pENTR 1A fragment.
Recombination reactions were then carried out to transfer the LOX or Decorin gene
into
expression
vector.
Briefly,
entry
clone
was
mixed
with
MSCV-IRES-GFP-GATEWAY expression vector and LR clonase II enzyme mix
(Invitrogen). The mixture was incubated at 25oC for 1 hour, and proteinase K was
then added to the reaction and incubated for 10 min at 37oC. The recombination
product could be used to transform competent cells. (As show in Figure 4.1)
59
Recombination by LR ClonaseTM
Growth on Ampicillin plates
after transformation
(-)
(-)
(-)
(+)
Figure 4.1 Plasmid contruction.
Retrovirus production
HEK 293 cells were transfected with effectene transfection system (Qiagen)
with
the
expression
plasmids
MSCV-LOX-IRES-GFP
or
MSCV-DECORIN-IRES-GFP together with packaging plasmid EcoPAC. The amount
of DNA used for transfection was 4.5μg expression plasmid and 5.5 packaging
plasmid (molecular ratio 1:1) per 150mm dish. 24 hours after transfection, medium
was discarded and 10ml fresh medium was added. Medium containing viral particles
was collected at 48 hours after transfection. Medium was then centrifuged at 3000rpm
for 15min at 4oC to pellet debris. Supernatant was stored at -80oC until use.
60
Cell infection
MEF cells were trypsinized and seeded into 6-well plate to grow overnight at
37oC in a humidified 5% CO2 incubator. 1.5ml viral stock was mixed with 1.5ml fresh
medium and then added to cells with 30-50% confluence after old medium was
removed. Polybrene was added to each well to a final concentration of 10 μg/ml. Cells
were incubated for 24 hours at 37oC in a humidified 5% CO2 incubator. The following
day, medium was removed and 2ml fresh medium was added to incubate for another
24 hours. Cells were checked for the efficiency of infection under a fluorescence
microscope by the expression of the GFP protein.
61
Generate viral expression
constructs containing genes
of interest (LOX or Decorin)
EcoPAC packaging
plasmid
293T cell line
Cotransfect the 293T cell line
with viral expression construct
and packaging plasmid
Harvest viral supernatant
Add the viral supernatant
to MEF cells
MEF cells
Assay for protein expression
of interest
promoter
Figure 4.2 Retrovirus production and MEF cells infection.
62
Reverse transcriptase polymerase chain reaction (RT-PCR)
Total RNA was extracted using Versagene RNA kit (Invitrogen). Reverse
transcription was accomplished with 5μg of total RNA using the TranscriptTM kit
(BioRad). PCR was carried out under the following conditions: denaturation at 95oC
for 1 min, annealing at 55oC for 1 min, extension at 72oC for 1 min (30 cycles), and a
final extension at 72oC for 10 min. PCR production were analyzed by electrophoresis
on a 1% agarose gel and visualized with ethidium bromide. The sequences of
oligonucleotide primers were as follows:
LOX:
Forward sequence:
5’-GACCTGGGGGCAGGTACCGG-3’
Reverse sequence:
5’-TGGGTGTTGGCATCAAGCAGG-3’
Decorin:
Forward sequence:
5’- CTTAACTCGAGATGAAGGCCACTATCATCCTCCT-3’
Reverse sequence:
5’-AATATGGATCCTTACTTATAGTTTCCGAGTTGAATGGC-3’
63
Western blotting
MEF cells infected with LOX or Decorin expressing retrovirus were lysed
with lysis buffer, and protein concentration was determined using the BCA kit. 10μg
of protein lysate and protein marker were run in 10% SDS-PAGE followed by transfer
to a PVDF membrane. Western analysis was carried out using the following primary
antibodies: goat polyclonal anti-LOX (E-19) antibody (Santa Cruz), and goat
polyclonal anti-human decorin antibody (R&D systems). After extensive washing,
immunocomplexes were detected with horseradish peroxidase-conjugated appropriate
secondary antibodies followed by enhanced chemiluminescence reaction (ECLTM,
Amersham).
Preparation of MEF-collagen constructs
First, collagen type I (rat tail collagen type I, BD Biosciences) gel was
prepared. Briefly, determine the final volume of collagen solution to be used and the
desired final collagen concentration (1.6mg/ml). 10X PBS with 1/10 final volume was
added to the collagen type I. Then 0.023 final volume of 1N NaOH was added into the
solution. Then appropriate DMEM culture media was added to the mixture to adjust
the collagen type I to final concentration 1.6mg/ml. Left the mixture on ice until ready
for use.
105 cells were mixed with collagen type I gel. Then the cell and collagen gel
mixture were pipetted into specially designed silicone dishes that permitted
contraction around posts (Figure 4.3). For each acellular sample, only the collagen
64
(1.6mg/ml) was pipetted into the silicone dish. All constructs remained in an incubator
(37oC, 5% CO2) for four weeks and were fed high glucose DMEM with ascorbic acid
and 10% FBS twice weekly. 4 weeks after cell culture, some of the samples were for
the mechanical test and the rest of them were for histology study.
Figure 4.3 Silicon mode. Each silicone dish consists of eight wells into which cells and collagen
gels were pipetted. The cells contracted the gels around two posts located in the base of each well.
Aligned Polyurethane fibers fabrication (electrospinning)
Fibers were electrospun from polymer solutions containing a degradable PU
(synthesized in our lab) based on lysine diisocyanate (LDI, Kyowa Hakko Koygo Co.,
Ltd, Tokyo, Japan).
5-7 wt/v% and degradable PU was dissolved in
dichloromethane:dimethylformamide (3:1) (DCM:DMF, Sigma) at concentrations of
13-17 wt/v%. Polymer solution was feed by a syringe pump (Medfusion 2010i;
Medex Inc.) at adjustable feed rates of 0.010- 0.020 ml/min through 1/16”
65
polyethylene tubing into a 30blunt tipped needle (Smallparts).
A voltage of 10 kV
was applied to the needle tip with a high voltage power supply (Gamma High Voltage
ES40P-10W).
The needle tip was held at 10 cm above the level of the collecting device.
The collecting devices were grounded. Polymer fibers deposited across the parallel plates
were collected with a home-made polycarbonate frame.
Cell culture in aligned nano-polyurethane fibers
The aligned polyurethane fiber frames were sterilized using 1N HCl and
washed with 1X PBS for several times. 106 normal MEF cells were seeded on the
nanofibers first. Two days after seeding, the MEF cells were confluent on the fibers,
and then the frames were turn over for the second time cell seeding. LOX or Decorin
expressing cells were seeded on the other side of the frames. The cells were cultured
with high glucose DMEM culture media with ascorbic acid and 10% FBS in 37oC,
5% CO2 incubator. One week after second time cell seeding, the frames were taken
out and the nano-fibers with cells were rolled over for animal surgery. One of the
frames was fixed with 4% Paraformaldehyde, and then stained with Alexa
546-phalloidin.
Briefly, the sample was blocked in TBSA-BSAT (10mM Tris-HCl, 150mM
NaCl, 0.02% sodium azide, 1% Triton X-100) for 1 hour at room temperature, and
then washed with wash buffer (PBS, with 0.05% sodium azide, 0.05% Triton X-100)
for 3 times, 10 minutes each. Then the sample was incubated in appropriate diluted
546-phalloidin solution for 2 hours at room temperature. The sample could be stored
66
in the wash buffer at 4oC and for later use. The stained sample was taken pictures
under Confocal microscope.
In vivo animal surgery
Nude rats were anesthetized and shaved to facilitate the incision. Incisions
were made on the back with scissors, and pockets adjacent to the incision site were
created with the aids of curved forceps. The nano-polyruethane fibers cultured with
different cells were implanted and the incision was closed. 4 weeks after surgery, the
samples were taken out from the pockets of rats, and some of them were for the
mechanical test, and some of them were fixed for histology study.
Histology
For histology study, the samples were fixed in 10% neutral buffered formalin.
All samples were then processed through a gradient of alcohols embedded in paraffin
blocks, sectioned, and stained with hematoxylin and eosin (HE staining).
For Sirius
red collagen staining, the sections were deparaffinized and rehydrated first. Then the
sections were dipped in saturated picro-sirius red solution for one hour and washed
with 0.01 N HCL solution twice for 0.5 minute each. Dehydrate the sample with 100
% ethanol and clear in xylene. The sections were mounted and ready for microscope
analysis.
67
Mechanical test
The retrieved samples were tested for mechanical properties using tensile test
(Bose Inc). The stress strain curves were obtained through a mechanical testing
software provided by the manufacturer.
Statistics analysis
Data will be presented as the mean +/- the standard error of mean for each
group. One-way analysis of variance (ANOVA) will be performed to determine the
effect of LOX or Decorin on the properties of regenerating tissue. Statistical
significance is accepted at p< 0.05.
Results and Discussion
LOX and decorin expression in MEF
Our goal in this study was to implement gene therapy as a tool to optimize the
performance of mechanically challenged engineered tissue. Mouse embryonic
fibroblasts were infected with LOX or decorin-expressing retrovirus. RT-PCR
confirmed the transcription of LOX or decorin genes compared to control group that
infected with retrovirus with empty vector (Figure 4.4A). LOX or decorin protein
expression was confirmed by western blotting (Figure 4.4B).
68
d
d
ct e
ct e
ct e
sfe
n
fe
fe
s
a
s
n
tr
ran
tra
rin
rt
o
X
o
c
t
O
c
De
ve
hL
d
OX
hL
tra
c
fe
ns
c
De
ted
i
or
ra
nt
c
fe
ns
c
ve
tor
t
ted
ct e
sfe
n
a
r
d
Lox
hLOX
Decorin
Decorin
GAPDH
Tubulin
A
B
Figure 4.4 LOX and decorin expression in MEF. A) Reverse transcription-polymerase chain reaction
(RT-PCR) analysis of RNA isolated from control MEF cells and LOX or Decorin expressing retrovirus
infected MEF cells. B) Western immunoblotting analysis of LOX or Decorin expression in MEF.
Mechanical test for in vitro experiment
After four weeks cell culture, there was significant difference in the strength
among the groups (Figure 4.5). The maximum load for the control MEF group was
0.2 ±0.04 N, and the maximum load for the LOX and decorin group was 0.8 ±0.13 N
and 0.5±0.09N, respectively. The maximum load for the collagen-cell graft
significantly increased after the cells transfected with LOX or decorin. The collagencell constructs were 1mm in width and 20mm in length. Young’s modulus of the
control grafts was 780 ±80KPa, young’s modulus of decorin-expressing group and
LOX-expressing group was 1980 ± 21.8KPa and 2110 ± 23.2KPa, respectively
(Figure 4.6).
69
0.8
Control
Lox Transfected
Decorin Treansfected
0.7
0.6
Force (N)
0.5
0.4
0.3
0.2
0.1
0
0
1
2
3
4
5
6
7
Displacement (mm)
Figure 4.5 Force-displacement curves of the collagen-cell grafts. (A) Control group. (B) LOX
transfected group. (C) decorin transfected group.
Young's Modulus (MPa)
2.5
2
1.5
1
0.5
0
Control group
Decorin-expressing group
LOX-expressing group
Figure 4.6 Young’s Modulus of collagen-MEF constructs.
70
Collagen plays a central role in tendon and ligament regeneration, as collagen
type I is responsible for major composition of the tissue, and its hierarchical
organization in bundles confers most of the tendon mechanical properties.
Consequently, tendon engineering studies mainly cell seeded collagen gels.
Contraction of the gel is related to the cell seeding density, generally followed by
alignment and reorganization of the matrix [16, 17]. But, up to now, no
tenocytes-collagen constructs have been able to achieve sufficient mechanical
properties, and the complex architecture of the tendon is never fully reproduced [18].
Also, collagen type I does not account for all the tissue properties. The role of
proteoglycans and small leucine rich protein (SLRP) in the organization and
mechanical properties of the tissue structure has been recently outlined [19]. In our
study, in order to enhance mechanical property of cell-collagen grafts, cross-linking
strategy was used by over-expressing LOX gene in MEF cells. Meanwhile,
over-expressing decorin gene in MEF cells was also used to enhance the mechanical
property of the tissue-engineered grafts. Our data showed that the LOX or decorin
expressing MEF-collagen grafts had much stronger mechanical property than the
control grafts.
TEM result for in vitro cell culture
TEM images of collagen fibril diameters are shown in Figure 4.7. Majority
collagen fibrils in the control cells (blank vector transfected) were 15 ~ 20 nm, in
LOX-expressing group were 30-35 nm, and in decorin-expressing group were 10~ 15
71
nm. Mean diameters of collagen fibrils in the decroin-expressing group were smaller
than control group and LOX-expressing group.
A
45
D
40
Percentage (%)
C
B
35
Control
LOX
Decorin
30
25
20
15
10
5
~6
5
60
60
55
~
~5
5
50
~5
0
45
45
40
~
30
~3
5
35
~4
0
~3
0
25
15
~2
0
20
~2
5
15
10
~
10
5~
0~
5
0
Size (nm)
Figure 4.7 TEM result for in vitro cell culture. A) Control group, B) LOX-expressing group, C) decorin
-expressing group, and D) The diameter distribution of collagen fibrils.
The major proteoglycan in the tendon or ligament is decorin, which anchors to
d-bands with several collagen fibrils by non-covalent binding [20]. Decorin is
composed of dermatan sulfate, which is a glycosaminoglycan that affects the
72
morphology and formation of collagen fibrils and core protein by covalent binding
[21]. It is reported that tendons of animals lacking decorin had collagen fibrils with
irregular diameters [14]. Down-regulation of decorin has also been shown to be
correlated with the development of collagen fibrils with large diameters [22]. Our data
showed that there were significant differences among the diameters of the control
group, decorin-expressing group, and LOX-expressing group.
Cells aligned after culture in nano-polyurethane fibers
Cells in all three groups are aligned well with the direction of oriented nanofibers
as shown in Figure 4.8. Mouse embryonic fibroblasts attached to scaffolds over a 24-h
period, after 3 days culture, the cells were aligned well on the oriented nanofibers
compared to the cells seeded on the normal culture flask (data not shown). One of the
considerable challenges remaining in the field of tendon and ligament tissue
engineering is to produce a construct with an anatomically correct architectural
framework, both in terms of cell morphology and matrix deposition. No cell-collagen
constructs are able to achieve sufficient mechanical properties, and the complex
architecture of the tendon is never fully reproduced. It is therefore hypothesized that
by creating an initial architecture, newly formed ECM will be deposited along the
prescribed fiber direction in these nanofibrous scaffolds, expediting the formation of a
neo-tissue with enhanced functional characteristics.
73
Figure 4.8 Cells aligned after culture on biodegradable polyurethane nanofibers. Blue color showed cell nuclei
stained with draq5 and green color showed transfected cells expressing green fluorescence protein (GFP).
Mechanical test for in vivo experiment
Four weeks after surgery, the grafts were taken out for mechanical test. Our
data showed that there was significant difference in linear stiffness between the
groups (Figure 4.9). The maximum load for the control group was 1.8 ± 0.3 N, the
maximum load for the LOX group and decorin group were 3.5 ± 0.6 N and 3.1 ± 0.5
N, respectively. The scaffolds were 1mm in diameter and 20mm in length. Young’s
modulus of the control grafts was 3200 ± 36.1KPa, young’s modulus of
decorin-expressing group and LOX-expressing group was 4670 ± 52.8KPa and 6130
± 69KPa, respectively (Figure 4.10). The strength of the graft was significantly
increased when the cells over-express LOX or decorin.
74
4
Control
Lox
Decorin
3.5
Force (N)
3
2.5
2
1.5
1
0.5
0
0
1
2
3
4
5
Displacement (mm)
6
7
8
Figure 4.9 Force-displacement curves of the nanofiber-cell grafts. (A) Control MEF cells. (B) Infected
MEF cells with LOX-expressing retrovirus. (C) Infected MEF cells with decorin-expressing retrovirus.
8
Young's Modulus (MPa)
7
6
5
4
3
2
1
0
Control group
Decorin-expressing group
LOX-expressing group
Figure 4.10 Young’s Modulus of the grafts retrieved from the animals.
75
There are some inconsistent results about the effect of decorin on collagen
fibrillogenesis and its biomechanical properties. Decorin-deficient mice were reported
to have fragile skin that is not able to withstand sudden tensile strain. Electron
microscopic examination of the skin in decorin knockout mice showed coarser,
irregular and haphazardly arranged collagen fibrils compared with wild type mice[14].
Overexperssion of decorin by rat arterial smooth muscle cells enhances contraction of
type I collagen in vitro[23]. In our study, overexpression of decorin in MEF cells was
also proved to increase the mechanical property of MEF-collagen graft although the
mean diameter of collagen fibrils was reduced. While Nakamura et al demonstrated
that in vivo treatment of injured rabbit ligament by antisense decorin
oligodeoxynucleotides led to an increased development of larger collagen fibrils in
early scar and a significant improvement in both scar failure strength and scar creep
elongation under loading which suggested that down-regulation of decorin helps to
improve the healing of various soft tissues including tendon and ligament[22]. One
explanation for that paradox is that decorin uniforms the collagen fibrils leading to
better-organized collagen structure that has improved mechanical property. It is
noticeable that the mean diameter of collagen fibrils in decorin-expressing group
(10-15nm) only showed slight decrease compared with control group (15-20nm). And
the distribution of diameters of collagen fibrils is more confined in decorin-expressing
group than in control group. We hypothesize that moderate overexpression of decorin
can help well organize collagen structure and improve its mechanical property but too
much decorin may induce the opposite effects by inhibiting collagen fibril formation.
76
In later situation, down-regulation of decorin may be necessary to improve the
collagen structure. Another explanation is that in our study decorin, which is essential
for the regulation of newly synthesized collagen, was secreted to regulate the fibril
formation of soluble collagen molecules while in Nakamura’s work decorin worked
on existing mature tendon. And the difference of animals used in the experiments
might also be counted.
Histology
Sirius red staining was used to examine the maturation of collagen fibrils. When
examined through crossed polars, the matured collagen fibers are bright yellow or
light red, and the immature collagen fibers are green. Figure 4.11 shows the Sirius red
staining of the grafts retrieved from the animals. Immature collagen fibrils were
dominated in control group (Figure 4.11 A) and decorin-expressing group (Figure 4.11
C) with overall green staining profile.
Significant more mature collagen fibrils were
found in LOX-expressing group (Figure 4.11 B) with more yellow and light red
staining profile.
77
A
B
C
50 μm
Figure 4.11: Sirius red staining of the grafts retrieved from the animals. (A) Control group. (B)
Lox-transfected group. (C) Decorin transfected group.
Conclusions
Our in vitro and in vivo experiments both showed that the mechanical property of
tissue-engineered graft could be significantly increased by over-expressing LOX or
decorin gene in fibroblast cells. The maturity of the collagen fibrils could be improved
or accelerated by over expressing LOX. The electrospinning technique can help cells
aligned on the nanofibers.
References
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Liu SH, N.T., Ankle sprains and other soft tissue injuries. Curr Opin
Rheumatol., 1999. 11(2): p. 132-7.
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Cofield RH, P.J., Hoffmeyer PJ, Lanzer WL, Ilstrup DM, Rowland CM.,
Surgical repair of chronic rotator cuff tears. A prospective long-term study. J
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Salehpour A, B.D., Proch FS, Schwartz HE, Feder SM, Doxey CM, Ratcliffe
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Badylak SF, T.R., Kokini K, Shelbourne KD, Klootwyk T, Voytik SL, Kraine
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Ouyang HW, G.J., Thambyah A, Teoh SH, Lee EH., Knitted
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CB., F., Ligament structure, physiology and function. J Musculoskelet
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Woo SL, J.F., Zou L, Gabriel MT., Functional tissue engineering for ligament
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McCormick RJ., Extracellular modifications to muscle collagen: implications
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Danielson KG, B.H., Holmes DF, Graham H, Kadler KE, Iozzo RV., Targeted
disruption of decorin leads to abnormal collagen fibril morphology and skin
fragility. J Cell Biol., 1997. 136(3): p. 729-43.
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Sini P, D.A., Tira ME, Balduini C., Role of decorin on in vitro fibrillogenesis
of type I collagen. Glycoconj J., 1997. 14(7): p. 871-4.
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Redaelli A, V.S., Soncini M, Vena P, Mantero S, Montevecchi FM., Possible
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80
PART II: CHAPTER 5
RECRUITMENT OF ENDOGENOUS STEM CELLS FOR TISSUE REPAIR
Abstract
The traditional concept of stem cell therapy envisions the isolation of stem
cells from patients, propagation and differentiation in vitro, and subsequent
re-injection of autologous cells to the patient. There are many problems associated
with this paradigm, particularly during the in vitro manipulation process and the
delivery and local retention of re-injected cells. An alternative paradigm that could be
easier, safer, and more efficient, would involve attracting endogenous stem cells and
precursor cells to the defect site for de novo tissue regeneration. Hepatocyte growth
factor (HGF), a pleiotropic cytokine of mesenchymal origin, exerts strong
chemoattractive effect on mesenchymal stem cells (MSCs) and neural stem cells
(NSCs), inducing migration of MSCs in vitro. However, HGF undergoes rapid
proteolysis in vivo, resulting in a very short lifetime of the bioactive cytokine. To
maintain the therapeutic level of HGF at the defect site necessary for endogenous
stem cell recruitment, sustained, long-term, and localized delivery of HGF is required.
Thiol-modified glycosaminoglycans hyaluronan (HA) and heparin (HP), combined
with modified gelatin (Gtn), were crosslinked with poly (ethylene glycol) diacrylate
(PEGDA) afford semi-synthetic ECM-like (sECM) hydrogels that can both provide
controlled growth factor release and permit cell infiltration and proliferation. Herein
we compare the use of different sECM compositions, for controlled release of HGF
and concomitant recruitment of human bone marrow MSCs into the scaffold in vitro.
Also, the stem cells recruitment ability by HGF-loaded scaffolds was demonstrated in
a subcutaneous implantation model in mouse.
Introduction
The traditional concept of cell therapy is based upon several basic steps. The
first step is cell sourcing and cells may be isolated from autologous, allogenic, or
xenogenic sources. Second, the isolated cells are expanded in vitro to a cell population
sufficient for effective treatment. The expanded cells can also be seeded on a scaffold
and cultured in a bioreactor. Finally, the expanded cells are re-implanted into the
patient. However, this final process is associated with ethical, economic, regulatory
and clinical problems.
Clinically, allogenic and xenogenic sources face the greatest likelihood if
immune rejection by the patient. Ethical and regulatory issues must also be resolved
for this to be a routine clinical treatment. Thus, autologous cells would seem to be the
best choice, but cell isolation from patients in need of treatment can cause additional
normal tissue morbidity. In order to obtain sufficient number for transplantation, in
vitro proliferation is essential, which may cause undesirable phenotype change [1].
Pluripotency of stem cells may decrease during in vitro culture [2, 3]. Allogenic and
xenogenic components used in culture may cause host immune rejection. In addition,
82
the cost for in vitro expansion of stem cells is very high, since a battery of growth
factors is needed for the propagation procedures. The economic and multi-week
expansion period present important challenges to these clinical procedures. Finally,
while mesenchymal stem cells attract much attention due to their pluripotency, the
pluripotency of mesenchymal stem cells decreases during in vitro culture [2, 3] using
conventional 2-D culture conditions.
An alternate cell source could be endogenous stem cells. Indeed, a
regenerative medicine approach for tissue repair focused on the direct manipulation of
endogenous adult stem cells is very appealing. There are several advantages to the use
of endogenous stem cells for tissue repair. First, using endogenous stem cells avoids
the immunocompatibility issues that accompany the use of allogenic and xenogenic
cells. Second, it is easier, safer, and more efficient to use endogenous stem cells for
tissue repair to expand and re-implant autologous cells. Third, only a single surgical
intervention is required, rather than two surgeries several weeks apart. Finally, the
process of recruiting endogenous stem cells offers both regulatory and economic
advantages relative to ex vivo approaches.
The utilization of endogenous stem cells may be enhanced in two ways. One
strategy is to mobilize the endogenous stem cells into the circulation. For example, it
is reported that granulocyte colony-stimulating factor (G-CSF) mobilizes stem and
progenitor cells from the bone marrow into the peripheral blood, from which they can
“home” into the lesion site in the brain and have a protective or restorative effect [4].
Also, the mobilized endogenous stem cells are showing promising outcome for
83
cardiac repair [5]. A second strategy is to enhance the recruitment of endogenous stem
cells into the lesion site for tissue regeneration. Several factors, such as hepatocyte
growth factor (HGF) [6] and stromal cell-derived factor-1 (SDF-1) [7] have shown
chemotaxic effects on stem cells. In this study, we used HGF as a model factor to
attract bone marrow mesenchymal stem cells into an ECM-like hydrogel in vitro.
Hepatocyte growth factor (HGF) has been shown to be a strong chemotactic factor the
mobilization and migration of mesenchymal stem cells (MSCs) [6]. Exposure of
MSCs to HGF induced migration of MSCs in vitro. However, like most proteins, HGF
undergoes rapid proteolysis in vivo, resulting in a very short lifetime of the bioactive
growth factor. The half-life of HGF that was delivered in a soluble form in vivo is 3-5
minutes.
In contrast, the time required to recruit a sufficient number of stem cells for
tissue repair is usually days to weeks. Therefore, direct injection of growth factors to
the repair site has limited success.
To maintain the therapeutic level of HGF at the repair site necessary for
endogenous stem cell recruitment, sustained, long-term, and localized delivery of
HGF is essential. Several polymer delivery systems are being developed for proteins
and growth factors delivery. Reservoir devices, solid implants, polymeric micro- and
nano-particles, and hydrogels are the most commonly used. Polymer systems have
many advantages; for example, they can stabilize proteins, provide localized delivery,
and produce diffusion-limited concentration gradients in tissues. ECM-derived
polymers with a wide array of physiological functions represent ideal substrates for
HGF delivery and stem cell recruitment, since ECM based materials may provide
84
adhesion sites for migrating stem cells to grow in. Recently, an injectable, in
situ-crosslinkable semi-synthetc ECM-like hydrogel, or sECM, was created by
crosslinking thiol-modified hyaluronan (HA) and gelatin (Gtn) using poly(ethylene
glycol) diacrylate (PEGDA) [8] This material offers a flexible composition and
compliance, simplicity of use, and was developed for ease of translation from in vitro
to preclinical in vivo as well as clinical uses [9]. In this study, we compared different
compositions of the sECM hydrogel for controlled release of HGF to recruit human
bone marrow mesenchymal stem cells (hMSCs) to the scaffold in vitro. Also, the stem
cells recruitment ability by HGF-loaded scaffold was evaluated in a subcutaneous
implantation model in mouse.
Materials and Methods
Materials
Thiolated chemically-modified HA (CMHA-S), thiol-modified gelatin
(Gtn-DTPH), thiol-modified heparin (HP-DTPH) and PEGDA were kindly provided
by Glycosan BioSystems Inc. (Salt Lake City, UT). Bovine serum albumin (BSA),
4’,6-diamidino-2-phenylindole (DAPI) and heparin were purchased from Sigma
Chemical Co. (St. Louis, MO). Recombinant human hepatocyte growth factor
(HuHGF) was purchased from PeproTech (Rock Hill, NJ). Draq 5 was purchased
from Alexis Corporation. Alexa 546-phalloidin was purchased from Molecular Probes
Inc. (Eugene, OR).
Recombinant mouse hepatocyte growth factor (mHGF) and
85
Recombinant mouse granulocyte colony stimulating factor (mG-CSF) were purchased
from R&D Systems, Inc. (Minneapolis, MN). Human HGF ELISA kit was purchased
from Biosource International, Inc. (Camarillo, CA). 8μm pore size transwells were
purchased from Corning Costar Inc. (Coring, NY). Rabbit polyclonal CD34 antibody
and monoclonal Stro-1 antibody were purchased from Santa Cruz Biotechnology, Inc.
(Santa Cruz, CA). Rat anti-mouse Sca-1/Ly6 antibody was purchased from R&D
Systems, Inc. (Minneapolis, MN). Secondary antibodies, Texas Red F(ab’) Fragment
Goat Anti-Rat IgG, Cy2 Goat anti-rabbit IgG (H+L) and Cy2 Goat anti-mouse IgG
(H+L), were purchased from Jackson ImmunoResearch Laboratories, Inc.(West Grove,
PA).
Hydrogel preparation for in vitro experiments
Two different sets of hydrogels were prepared. For HA:Gtn gels, CMHA-S
and Gtn-DTPH in DI water solution (pH 7.4) were mixed in a volume ratio of 1:1, and
then HA:Gtn solution was cross-linked by mixing it with 2 wt% PEGDA in DI water
in a volume ration of 4:1. For HA:Gtn:HP gels, HP-DTPH in DI water solution (pH
7.4) were mixed with the HA:Gtn solution mentioned above: the concentration of HP
was 0.3%. Hepatocyte growth factor (HGF) was incorporated non-covalently in the
gels by pre-mixing 500 ng HGF with 200 μL of the modified HA and gelatin solution
prior to cross-linking. Gels formed within 20 minutes.
86
In vitro release kinetics of hHGF
HGF was allowed to release from these hydrogels at 37 oC into a PBS buffer
solution supplemented with 1% BSA, 1 mM EDTA, and 10 μg/mL heparin. At days 1,
2, 3, 5, 9, 14, 20, and 26, 100μl of conditioned medium was sampled, and an equal
volume of fresh release medium was added back to maintain the total volume.
Samples were then immediately frozen at -20oC until measurement. Each release
condition was performed in triplicate.
HGF measurement with enzyme-linked immunosorbent assay (ELISA)
Released amount of HGF was quantified with sandwich enzymelinked
immunosorbent assay (ELISA) using an assay from a commercially available kit
(Biosource). Briefly, 100 μLof the sample was added into the designated ELISA wells
and incubated at 4 oC overnight. After the incubation, the plate was washed 4 times
with a wash buffer. Then, 100 μL of biotin antibody was added into each well, and the
plate was incubated at room temperature for 1h. Then, the plate was washed 4 times,
and 100 μL Streptavidin solution was added and incubated for 45 min at room
temperature. Then, the substrate was added to the wells and the absorbance was
measured at 450 nm. Total accumulated released HGF was calculated by integration
of the individual measurements over the cumulative time of the experiment.
87
Human bone marrow derived mesenchymal stem cells culture
Human bone marrow–derived mesenchymal stem cells (hMSCs) were
obtained from Lonza (Allendale, NJ). The expression of CD105 and CD 44
characterized before use for this study. The expression of CD 105 and CD 44 are more
than 92%. hMSCs were maintained in culture in MSCGM™ - Mesenchymal Stem
Cell Medium (Lonza, Allendale, NJ).
In vitro experimental procedure for stem cell recruitment
The migration capacity of hMSCs was analyzed using Costar Transwell
invasion chambers with polycarbonate membrane filters of 6.5 mm diameter and 8μm
pore size to form dual compartments in a 24-well tissue culture plate. Samples each
containing 104 cells in 200 μL of media were added to the upper compartments. The
lower compartments were covered with 200 μL hydrogels, with or without HGF as a
source of chemoattractants, and 400 μL culture media were added. The migration
chambers were incubated for 8 hours at 37 oC and 5% CO2. After incubation, cells on
the top surface of the filters were wiped off with cotton swabs. Cells that had migrated
into the lower compartment and attached to the lower surface of the filter were
counted after being stained with Alexa 546-phalloidin and Draq 5. Briefly, the
samples were blocked in TBSA-BSAT (10mM Tris-HCl, 150mM NaCl, 0.02%
sodium azide, 1% Triton X-100) for 1 hour at room temperature, and then washed
with wash buffer (PBS, with 0.05% sodium azide, 0.05% Triton X-100) for 3 times,
10 minutes each. Then the samples were incubated in appropriate diluted
88
546-phalloidin solution for 2 hours at room temperature. After 3 times of wash in
wash buffer, samples were incubated in appropriate diluted Draq 5 for 15 minutes at
room temperature. Finally, after 2 times of wash in wash buffer, samples were
incubated in appropriate diluted DAPI solution for 15 minutes at room temperature.
The samples could be stored in wash buffer at 4oC for later use. The stained samples
were taken pictures under Confocal microscope.
Polyurethane (PU) scaffolds fabrication
Three-dimensional, porous, scaffolds were fabricated using a particulate
leaching technique.
Briefly, 16 wt% of polyurethane (SG80-A, Noveon Inc,
Cleveland, OH, USA) was dissolved in dimethylacetamide (DMAC, Sigma-Aldrich
Inc., St. Louis, MO, USA) at 60°C for 12 hours. Grounded sodium chloride particles
at 100 ~ 150 μm in diameter were then added into the SG80-A in DMAC solution at
45 wt% and mixed thoroughly. This was then poured into stainless steel molds, which
were placed in a dry ice/ethanol bath for 15 minutes. Finally, the molds were dropped
into distilled water until the scaffolds released from the molds.
The scaffolds were
then placed into another beaker of distilled for one week to allow the salt to dissolve.
The scaffolds were frozen at -20 °C for several hours before they were cut into
rectangular slices (5 mm×3 mm×3 mm). The slices were then washed in distilled
water for 12 hours before sterilized using ethylene oxide (EtO). The scaffolds were
immersed into HA-Gtn gel with or without mHGF at 4oC overnight. The next day, the
scaffolds, which were loaded gel, were picked out and ready for animal surgery.
89
Morphological analysis of the scaffolds was conducted using scanning
electron microscopy (SEM, JEOL 5410). Scaffolds were allowed to vacuum dry for
24 hours prior to SEM analysis. The samples were sputter coated with gold at a
thickness of 50-70 nm using a Cressington 108 AUTO sputter-coater with a current of
30 mA for 2 minutes. The coated scaffolds were analyzed on a SEM with an
acceleration voltage of 10 kV. Representative images of the scaffolds were obtained.
Scaffold implantation
The stem cells recruitment ability was evaluated in a mouse subcutaneous
implantation model. Male DBA/2 mice aged 6-8 weeks were injected with MG-CSF
(5μg/100g body weight) for 5 consecutive days through intraperitoneal route. The
mice were anesthetized and shaved to facilitate the incision. Incisions were made on
the back with surgical blade, and pockets adjacent to the incision site were created
with the aids of curved forceps. Six scaffolds (with or without mHGF) were implanted
per mouse.
Histology
Seven days after surgery, the mice were anesthetized and perfused. The
scaffolds were fixed in 10% formalin. The scaffolds were then embedded in paraffin,
section and stained with stem cell markers (Stro-1), DAPI and Draq 5. Briefly, after
embedded in paraffin and cut into 10μm sections using microtome. The sections were
washed with xylene for 5 minutes each, and were incubated in two washes of 100%,
90
95%, 85%, 75%, and 50% ethanol for 10 minutes each. The sections were rinsed
twice in DI water for 5 minutes each. Incubated the sections in 0.03% H2O2, 80%
methanol for 30 minutes and rinsed twice in DI water for 5 minutes each. Sections
were incubated in 10mM sodium citrate buffer pH6.0 at room temperature for 1 hour
and then boiled using water bath or microwave and maintain at 95-99oC for 10
minutes. Sections were cooled to room temperature, rinsed in DI water three times for
5 minutes each, and rinsed in PBS for 5 minutes. Sections were blocked in 4% goat
serum in PBS for 1 hour. Primary antibodies (Stro-1) were prepared as indicated on
manufacturer datasheet in 1XPBS with 4% goat serum. After blocking, primary
antibody was applied. Sections were incubated at 4oC overnight. The specimens were
rinsed three times in 1XPBS with 0.3% Triton X-100 for 5 minutes each. Then the
specimens were incubated in appropriate secondary antibodies for 2 hours. After 3
times washing in wash buffer, samples were incubated in 1:10,000 diluted Draq 5 for
15 minutes at room temperature. Finally, after 2 times washing in 1XPBS, sections
were incubated in appropriate diluted DAPI solution for 15 minutes at room
temperature. The sections were mounted for visualization using confocal laser
microscope.
Statistical analysis
Data will be presented as the mean +/- the standard error of mean for each
group. One-way analysis of variance (ANOVA) will be performed to determine the
effect of HGF loading on the recruitment of stem cells. Statistical significance is
91
accepted at p< 0.05.
Results
Cumulative in vitro HGF release from ECM-based hydrogel
HGF release profiles from the hydrogel with or without heparin were in vitro.
Immobilization of heparin in HA-Gtn based hydrogel may protect growth factors from
enzymatic
degradation
and
thermal
denaturation
[10].
These
crosslinked
heparin-containing hydrogel behave in an analogous fashion to a heparan sulfate
proteoglycan, extending the release times of growth factors in vitro and in vivo while
retaining their bioactivity [11, 12]. Figure 5.1 presents the time course of total
cumulative release for HGF. Covalently cross-linked heparin slows down the HGF
release significantly in HA: Gtn based gels, but also reduces the net quantity released
at a given time point. Without heparin, HA: Gtn gels released a total of 35% of the
initially loaded HGF in 26 days. In contrast, HA: Gtn: HP gels released a total of 18%
of their loaded HGF at the end of 26 days.
92
300
Accumulated Release Amount (ng)
A
250
200
HA-Gtn-DTPH
150
100
HA-Gtn-HP-DTPH
50
0
0
5
10
15
20
25
30
Time (Days in culture)
60
Percentage of Release Amount (%)
B
50
40
HA-Gtn-DTPH
30
20
HA-Gtn-HP-DTPH
10
0
0
5
10
15
20
25
30
Time (Days in culture)
Figure 5.1 Cumulative in vitro HGF release from HA: Gtn and HA: Gtn: HP hydrogels. (A) Release
amount in nano grams and (B) Percentage of release.
93
Bone marrow mesenchymal stem cells migration in vitro
In this experiment, four groups were assigned. In the negative control group, no
HGF was present in either the lower or upper compartment of Transwells. In the
positive control group, 20 ng/mL HGF was present in the culture media of the lower
compartment of Transwell. In the third group, 500 ng of HGF encapsulated in an HA:
Gtn gel was placed in each lower compartment. In the fourth group, 500 ng of HGF
encapsulated in HA: Gtn: HP gel was placed in each lower compartment.
In order to analyze the migration ability of hMSCs in response to released
HGF, two locations were observed under a confocal microscope: the lower surface of
the filter and the hydrogel in each lower compartment (Figure 5.2). Figure 5.3 shows
the number of hMSCs that migrated to the lower surface of the filter. All four groups
(Figure 5.3B, 5.3C, 5.3D) showed significantly greater number of MSCs on the lower
surface of the filter than the negative control group (Figure 5.3A). Figure 5.4 presents
the cell density in cells per mm2 on the lower surface of the filter. Significantly more
MSCs migrated across the filter than for each group relative to the negative control.
The numbers of cells recruited into the hydrogels in the lower compartments
were also determined. Large numbers of cells were found inside the HA: Gtn gel
loaded with 500 ng of HGF (Figure 5.5C). Indeed, significantly more cells were found
in the HA: Gtn gel loaded with 500 ng of HGF (Figure 5.5C) than in any of the other
three groups: negative control group without HGF loading (Figure 5.5A), positive
control group with 20 ng/mL of soluble HGF in medium (Figure 5.5B), and the HA:
Gtn: HP hydrogel loaded with 500 ng of HGF (Figure 5.5D). Paradoxically, no cells
94
were found in HA: Gtn: HP gel loaded with 500 ng HGF Figure (5.5D), similar to the
cell density in the negative control group Figure 5.5A.
95
Transwell
insert
Upper
compartment
Microporous
membrane
Lower
compartment
Cells
Hydrogel
1 mm
Figure 5.2: 8μm pore size transwell. The device includes two compartments: the lower
compartment and the upper compartment. The insert has a polycarbonate membrane with 8.0µm Pores.
The cells were seeded on the membrane and the hydrogels were coated on the surface of the lower
compartment.
A
B
C
D
300 um
Figure 5.3: MSCs across the 8 μm pores to the lower surface of the filter. (A) Negative control (no
HGF), (B) positive control (20 ng/mL HGF in culture media), (C) 500 ng HGF encapsulated in HA:
Gtn hydrogel, and (D) 500 ng HGF encapsulated in HA: Gtn: HP hydrogel.
96
45
Number of hMSCs across the 8 um filter
after 8 h in culture (#/mm2)
40
35
30
25
20
15
10
5
0
No HGF
20ng/ml HGF in
culture media
500 ng HGF in HAGtn gel
500 ng HGF in HAGtn-HP gel
Figure 5.4: The number of MSCs across the 8 μm pores to the lower surface of the filter.
97
A
B
C
D
300 um
Figure 5.5: MSCs migrated into the hydrogels on the lower compartment . (A) Negative control (no
HGF), (B) positive control (20 ng/ml HGF in culture media), (C) 500 ng HGF encapsulated in HA: Gtn
hydrogel, and (D) 500 ng HGF encapsulated in HA: Gtn: HP hydrogel.
98
100
Number of hMSCs migrated into the gel
3
after 8 h in culture (#/mm )
90
80
70
60
50
40
30
20
10
0
No HGF
20ng/ml HGF in
culture media
500ng HGF in HA- 500ng HGF in HAGtn gel
Gtn-HP gel
Figure 5.6: The number of MSCs migrated into the hydrogels on the lower compartment.
SEM for porous polyurethane (PU) scaffolds and hydrogel scaffolds
Figure 5.7 shows the scanning electron microscopy (SEM) images of
Polyurethane (PU) scaffold. Leaching is one popular approach used in tissue
engineering to fabricate porous scaffolds. Pores or channels are created using
porogens, such as salt (NaCl), wax, or sugar. In this study, NaCl was used for pores
making. For particle leaching, pores are regulated by the salt particles and 100μm-150
μm salts were used, so that the pores would be around 100 μm-200μm and larger than
99
the cells size (about 15μm). After the solvent (DMAC) was removed and the salt was
washed, pores can be seen in the interior and on the surface of the scaffold under SEM
(figure 5.7), and the pores are well interconnected throughout the scaffold. There is
enough space for cell migration into the hydrogel.
A
B
Figure 5.7: SEM images showing morphology of porous scaffold with pore sizes 100-200 μm in
diameter.
Mesenchymal stem cells recruitment in a mouse subcutaneous implantation model
In order to evaluate the stem cell recruitment potential of HGF loaded
scaffolds in vivo, a subcutaneous implantation mouse model was used. According to
the profile of the cumulative in vitro HGF release, HA:Gtn hydrogel was chosen for
the in vivo study. In order to identify the implant at the time of scaffold harvesting,
100
HA:Gtn hydrogels were infiltrated into nondegradable polyurethane porous scaffolds
The scaffolds with or without mHGF loading were implanted to the back of the mice,
and 1 week after surgery, the scaffolds were processed for immuno-staining and
visualized under confocal laser microscope.
First, the number of cells infiltrated into the porous scaffold was quantified
and compared among groups (Figure 5.8 A, D, and G). Significantly more cells were
detected in the mHGF releasing scaffolds (Figure 5.8 D) than the control scaffolds
(Figure 5.8 A) without mHGF loading. This data demonstrated that more cells could
be recruited to the mHGF releasing scaffolds. In order to identify whether there are
stem cells among the recruited cells, mesenchymal stem cell marker, Stro-1, was used
to identify non-hematopoietic stromal stem cells [31]. Our data show that about
14.2±3.6 % of recruited cells expressed mesenchymal stem cell marker (Stro-1) in the
scaffolds loaded with mHGF. While in the non-mHGF releasing scaffolds, the
percentage of the cells expressed Stro-1 is significantly lower at 7.9±1.3 %. This
demonstrated that scaffolds loaded with HGF may recruit more stem cells to the local
implantation site (Figure 5.8 H).
101
A
B
C
D
E
F
30 μm
12000
1600
G
H
1400
Number of Cells/mm
2
Number of Stro-1 (+) Cells/mm2
10000
8000
6000
4000
2000
1200
1000
800
600
400
200
0
0
No-HGF
HGF
No-HGF
HGF
Figure 5.8 Cells migrated into the scaffoldss after 1-week implantation. A-C) Scaffold without mHGF
release, D-F) scaffold with mHGF release, A,D) Nuclei staining in blue, B and E) Stro-1 Staining in
green. C and F) The overlay images of nuclei staining and Stro-1 staining. G) The number of cells in
the scaffolds. And (H) The number of stro-1 positive mesenchymal stem cells.
Discussion
Thiolated hyaluronan are biocompatible and can be adapted to produce a variety
of desirable biologic effects [13]. Gelatin is a partially hydrolyzed collagen product,
102
which is an excellent substrate for cell attachment, proliferation, and differentiation.
Both thiolated HA [14, 15] and thiolated gelatin [16, 17] have been used in a number
of tissue engineering applications, including the delivery of basic fibroblast growth
factor (bFGF) for cutaneous wound repair [18] and human demineralized bone matrix
for bone repair [19]. The hydrogels created by co-crosslinking thiol-modified HA with
thiol-modified gelatin produces mechanically robust, bioresorbable scaffolds that can
be implanted to achieve tissue growth in vivo[13, 20-22]. The HA-Gtn hydrogel is
also an effective vehicle for cell delivery and retention in vivo, including the repair of
osteochondral defects using encapsulated autologous MSCs [23]. Growth factors
delivered locally using the HA-Gtn hydrogels elicit a strong angiogenic response,
even when the growth factors are delivered in low, nanogram doses [11-13, 24]. In
addition, it is reported that incorporation of small amounts of heparin in the gels can
prolong and spatiotemporally regulate the rate of growth factor release in vitro [11,
12]. In this study, we demonstrated that through delivery of HGF from
heparin-containing HA: Gtn hydrogels or HA: Gtn hydrogel only, the biologic activity
of HGF can be maintained, and the human mesenchymal stem cells are most
effectively recruited into the HGF releasing hydrogels when the release rate is
maintained to certain threshold.
As commonly occurs with highly bioactive polypeptides in vivo, HGF undergoes
rapid proteolysis and has a half-life in vivo of 3-5 minutes. Figure 5.1 illustrated that
release of HGF from crosslinked thiol-modified HA and gelatin hydrogel in vitro
could be sustained for up to 3 to 6 months based on the extrapolated value from a
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26-day study. This is a dramatic increase in time of availability compared to the short
half-life of free HGF in vivo. Furthermore, the rate of HGF release could be controlled
with heparin. In the absence of heparin, HGF released twice as fast from HA:Gtn
hydrogel than the HA:Gtn:HP hydrogel. This result is consistent with extended release
of bioactive keratinocyte growth factor (KGF), vascular endothelial growth factor
(VEGF), angiopoetin-1 (Ang-1), and bFGF from similar heparan-sulfate mimetic
hydrogels [10-12]
The biological activity of the HGF released from the hydrogels was confirmed
by in vitro hMSCs migration experiments. On the lower surface of each filter, all
groups except the negative controls showed an obvious accumulation of cells (Figures
5.3 and 5.4). In the hydrogels in the lower compartments, many cells migrated into the
HA: Gtn gel loaded with 500ng HGF, while there were few cells attached on the
negative control group or the HA:Gtn :HP gel loaded with 500ng HGF (Figures 5.5
and 5.6). This seemingly paradoxical result seems to indicate that the amount of
loading and the release rate of HGF from the gel are both crucial parameters affecting
the migration of hMSCs. If the release rate or amount of HGF is lower than the
threshold for hMSC to response, there will be no control of hMSCs to migrate, since
when we double the amount of HGF loading into the HA-Gtn-HP gel to 1000ng,
significant amount of hMSCs migrated into the HA-Gtn-HP hydrogels. According to
cumulative release of hHGF from two types of hydrgels (Figure 5.1), HA:Gtn
hydrogel released about 10% of total stored hHGF (about 50ng)at the first two days,
while HA:Gtn:HP hydrogel released less than 5% of total loaded hHGF (less than
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25ng) at the first two days. In the literature, the minimum effect dosage for HGF
activating mesenchymal stem cell migration was 20ng/ml [13, 27]. In our experiment,
the incubation time for the mesenchymal stem cells migration was short (8 hours),
hHGF did not have enough time to release to the amount that can attract stem cell
migration efficiently, especially the HA-Gtn_HP hydrogel.
The other interesting indication is the importance of HGF gradient in the control
of MSC migration. If we compare the 20 ng/mL HGF group (Figure 5B) with the
500ng/mL group (Figure 5C), fewer MSCs are migrated to the hydrogel in 20ng/ml
group (Group II) than that in slow releasing group (Group III). There are sufficient
amount of HGF in group II for MSC to migrate, however, the difference between
group II and III is the establishment of HGF gradient. HGF gradient is well
established in group III due to the constant release of HGF from the HA-Gtn
hydrogel.
In vivo study further verified the observation in vitro. There were significant
more cells observed in the mHGF-loaded scaffolds than that in control scaffolds
(Figure 5.8 G). The total numbers of recruited stro-1 positive mesenchymal stem cells
were also significantly higher in mHGF group (Figure 5.8 H).
Conclusions
The release characteristics of HGF and stem cell recruitment capacity of a
series of HA-Gtn hydrogels loaded with HGF have been investigated in vitro and in
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vivo. These hydrogels provide sustained release of biologically active HGF in vitro
and in vivo, with HGF release sustained for over three weeks. This is a dramatic
increase in time of availability compared to the short half-life of free HGF in vivo.
HGF released from HA: Gtn hydrogel attracted human bone marrow MSCs to migrate
into the hydrogel following the HGF gradient established by HGF loaded HA-Gtn
hydrogels. To evaluate the stem cell recruitment ability in vivo, subcutaneous
implantation mouse model was used. Significantly more cells were detected in the
mHGF releasing scaffolds than the control scaffolds without mHGF loading.
Mesenchymal stem cell marker, Stro-1, can be detected in the recruited cells. Our in
vitro and in vivo studies demonstrated that stem cells could be recruited to the local
site where HGF was released by chemically modified HA and modified gelatin based
hydrogels. And this strategy is promising for tendon and ligament tissue regeneration
applications.
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CHAPTER 6
FUTURE DIRECTIONS
In the tissue maturation part of this thesis, we figured out that
over-expression of LOX or decorin genes in cells could increase the stiffness of
tissue-engineered grafts respectively by different mechanism. LOX cross-links
collagen molecules to enhance collagen maturation, while decorin increases the
stiffness of the grafts by uniforming the diameter of collagen fibers and regulating
collagen fibrogenesis. In our future study, we are going to over-express both LOX and
decorin to figure out whether they can further increase the stiffness of the graft, and to
design a graft that has similar mechanical and functional characteristics of the native
tissue in a short period of time.
The second part of this thesis is to use endogenous stem cells for tendon and
ligament regeneration. Our in vitro and in vivo studies demonstrated that stem cells
could be recruited to the local site where HGF were released by chemically modified
HA and gelatin based hydrogels. In the future work, we plan to optimize heparin
concentration in the hydrogel for growth factor releasing profile that best promotes
stem cell recruitment. Furthermore, we are going to control the differentiation of the
recruited stem cells into desirable cell types by controlling release of differentiation
factors.