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Immunomodulatory and regenerative potential of amniotic fluid stem cells as a treatment strategy for pulmonary fibrosis
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Immunomodulatory and regenerative potential of amniotic fluid stem cells as a treatment strategy for pulmonary fibrosis
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IMMUNOMODULATORY AND REGENERATIVE POTENTIAL OF AMNIOTIC
FLUID STEM CELLS AS A TREATMENT STRATEGY FOR PULMONARY
FIBROSIS
by
Orquidea Helen Garcia
A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(INTEGRATIVE BIOLOGY OF DISEASE)
May 2013
Copyright 2013 Orquidea Helen Garcia
ii
DEDICATION
This dissertation is dedicated to my best friend and biggest supporter: Tony. Thank you
for supporting me as I chased my dream. You kept me grounded and kept me going, and
in all honesty, probably deserve a PhD yourself for all of the late nights and weekends
you spent in the lab with me. Without you I could not have achieved this
accomplishment.
iii
ACKNOWLEDGEMENTS
First and foremost I would like to acknowledge my mentors: Dr. David Warburton and
Dr. Barbara Driscoll. Dr. Warburton, thank you for challenging me by allowing me to
create my own project and set my own trajectory in the sciences by treating my graduate
career as a mini-postdoc. This type of environment, a calculated risk for a graduate
student and mentor, has served as a launch pad for my personal and professional
development as well as for my scientific career. Dr. Driscoll, your passion for
mechanistic science and gift for teaching are qualities I hope to emulate in my own
career. Thank you for all of the advice and guidance you provided while I navigated my
graduate career. I would also like to thank Dr. Laura Perin who has become invaluable to
my development as a scientist as my graduate project has progressed. Your creativeness,
ingenuity, honest advice and support have been invaluable to the direction and success of
this project.
I would like to thank my dissertation committee members, Dr. Wei Shi and Dr. Tom
Keens, along with program director Dr. Alicia McDonough for your critiques and advice
throughout my graduate education.
Finally, I would like to thank those that have been invaluable to the development and
execution and this project: Sue Buckley, Astgik Petrosyan, Gianni Carraro, Gianluca
Turcatel, Jooeun Lee, Ragu Reddy, Sargis Sedrakyan, Stefano Da Sacco, Sarah Utley,
iv
Zoe Ly and last but not least Dawn Burke whose tireless behind the scenes efforts have
not gone unnoticed or unappreciated.
v
TABLE OF CONTENTS
DEDICATION .................................................................................................................... ii
ACKNOWLEDGEMENTS ............................................................................................... iii
ABSTRACT ....................................................................................................................... xi
CHAPTER ONE: CELL BASED THERAPIES FOR LUNG DISEASE
1.1 Introduction: What are cell-based therapies? .................................................... 1
1.2 Cellular therapeutic strategies for the lung: challenges and possibilities ......... 1
1.3 What is a stem/progenitor cell? ......................................................................... 4
1.4 Endogenous progenitors and their niches in the lung ....................................... 6
1.5 Exogenous stem and progenitor cell candidates for cell therapies ................. 12
1.5.1 Hematopoietic stem cells and mesenchymal stem cells .................. 12
1.5.2 induced Pluripotent Stem Cells (iPS cells) and Embryonic Stem cells
(ES cells) ................................................................................................... 13
1.5.3 Fetal associated stem cells: Amniotic Fluid Stem Cells (AFSC), cord
blood stem cells and placental stem cells ................................................. 14
1.6 Engineering whole lung regeneration ............................................................. 15
1.7 Production and delivery of cellular therapies ................................................. 16
1.8 Cellular therapy versus whole lung engineering ............................................. 17
1.9 Areas of agreement ......................................................................................... 19
1.10 Areas of controversy ..................................................................................... 19
1.11 Future challenges .......................................................................................... 20
CHAPTER TWO: AMNIOTIC FLUID STEM CELLS
2.1 Introduction ..................................................................................................... 21
2.2 The amniotic cavity ......................................................................................... 22
2.3 The amniotic fluid ........................................................................................... 24
2.4 Amniotic Fluid-Derived Stem Cells (AFDSCs) ............................................. 27
2.5 Collection of amniotic fluid for Amniotic Fluid Stem Cell isolation ............. 29
2.6 Tissue engineering from Amniotic Fluid-Derived Stem Cells ....................... 30
2.7 Collection, isolation and culture of c-kit
+
Amniotic Fluid Stem Cells ........... 31
2.8 Why use AFSC in Regenerative Medicine? ................................................... 32
2.9 Differentiation of Amniotic Fluid Stem Cells for regenerative medicine ...... 33
2.9.1 Hematopoietic System ..................................................................... 34
2.9.2 Brain ................................................................................................. 35
2.9.3 Bone ................................................................................................. 35
2.9.4 Kidney .............................................................................................. 35
2.9.5 Lung ................................................................................................. 38
2.9.6 Heart ................................................................................................ 40
2.10 Conclusion .................................................................................................... 41
vi
CHAPTER THREE: MODULATING THE ALVEOLAR MILIEU TO ENHANCE
RESOLUTION OF FIBROTIC LUNG INJURY
3.1 Introduction ..................................................................................................... 44
3.2 Response of the alveolar epithelium to injury/disease .................................... 45
3.3 The distal alveolar epithelial milieu is the key to alveolar epithelial
homeostasis ........................................................................................................... 47
3.4 Proteomic analysis of bronchoalveolar lavage (BAL) can be used to
characterize the alveolar milieu ............................................................................ 49
3.5 Cell based therapies have the potential to limit progression of fibrotic lung
disease ................................................................................................................... 49
3.6 Amniotic fluid stem cells augment endogenous alveolar epithelial healing ... 50
3.7 Conclusion ...................................................................................................... 51
CHAPTER FOUR: AMNIOTIC FLUID STEM CELLS INHIBIT THE PROGRESSION
OF BLEOMYCIN INDUCED PULMONARY FIBROSIS VIA CCL2 MODULATION
IN BRONCHOALVEOLAR LAVAGE.
4.1 Introduction ..................................................................................................... 53
4.2 Methods ........................................................................................................... 56
4.2.1 Ethics Statement ............................................................................... 56
4.2.2 Isolation and Culture of AFSC ........................................................ 56
4.2.3 Bleomycin Induced Lung Injury and AFSC Treatment ................... 57
4.2.4 Histology .......................................................................................... 58
4.2.5 Measurements of Lung Mechanics and Collagen Quantification .... 58
4.2.6 Collection of Bronchoalveolar Lavage (BAL) and Lung Tissue ..... 59
4.2.7 Proteomic Cytokine Analysis .......................................................... 60
4.2.8 CCL2 ELISA ................................................................................... 60
4.2.9 In Vitro Collagen Assay ................................................................... 60
4.2.10 Western blotting and Zymography ................................................ 61
4.2.11 Migration Assay ............................................................................. 61
4.2.12 In Vitro AFSC Rescue ................................................................... 62
4.2.13 In Vitro MMP2 inhibition .............................................................. 63
4.2.14 Data Presentation and Statistical Analysis ..................................... 63
4.3 Results ............................................................................................................. 63
4.3.1 AFSC treatment inhibits fibrotic parenchymal destruction 28 days
post-bleomycin injury in vivo ................................................................... 64
4.3.2 AFSC treatment inhibits loss of pulmonary function associated with
the development of pulmonary fibrosis 28 days post-bleomycin injury in
vivo ............................................................................................................ 67
4.3.3 AFSC modulate acute inflammatory cytokine expression in BAL and
lung tissue in vivo ...................................................................................... 70
4.3.4 AFSC modulate CCL2 through MMP2 mediated proteolytic
cleavage ..................................................................................................... 74
4.3.5 AFSC chemotactically respond to increased CCL2 gradients ......... 76
vii
4.3.6 AFSC co-cultured with bleomycin injured AECII inhibit increased
CCL2 expression in vitro .......................................................................... 80
4.3.7 Inhibition of MMP2 in vitro attenuates the ability of AFSC to reduce
CCL2 expression ....................................................................................... 82
4.4 Discussion ....................................................................................................... 84
CHAPTER FIVE: REPOPULATION OF DECELLULARIZED LUNG MATRIX
USING AMNIOTIC FLUID STEM CELLS
5.1 Introduction ..................................................................................................... 88
5.2 Methods ........................................................................................................... 90
5.2.1 Isolation and characterization of human AFSC .............................. 90
5.2.2 Characterization of human AFSC via qPCR for subpopulations with
lung specific markers ................................................................................ 91
5.2.3 Western blot analysis of lung lineage markers ................................ 92
5.2.4 Decellularization of lung matrix ...................................................... 92
5.2.5 Seeding and Culture of AFSC on decellularized lung ..................... 93
5.2.6 Histology .......................................................................................... 94
5.2.7 Analysis of secreted cytokines, growth factors and pluripotential
characteristics ............................................................................................ 94
5.3 Results ............................................................................................................. 94
5.3.1 AFSC express Type I and Type II alveolar epithelial lineage markers
prior to culture on decellularized lung matrices ....................................... 94
5.3.2 Characterization of secreted cytokines, growth factors and
interleukins prior to culture on lung matrix .............................................. 99
5.3.3 Decellularization removes cellular material from extracellular matrix
without disrupting alveolar structure ...................................................... 101
5.3.4 AFSC seeded on decellularized matrices can be cultured for up to 35
days ......................................................................................................... 102
5.4 Conclusion .................................................................................................... 104
CHAPTER SIX: CONCLUDING REMARKS AND FUTURE DIRECTIONS ........... 106
REFERENCES ............................................................................................................... 110
BIBLIOGRAPHY ........................................................................................................... 124
APPENDICES ................................................................................................................ 138
viii
LIST OF TABLES
Table 1.1 Lung Diseases Potentially Treatable with Cellular Therapies ............................ 4
Table 3.1 Key signaling molecules within the pro-fibrotic alveolar milieu ..................... 48
Table 5.1 Identification of the 5 human AFSC lines tested .............................................. 91
Table 5.2 Gene, lineage and primer sequences for lung lineage markers tested .............. 92
Table 5.3 Gene, expression and lineage of lung markers tested ....................................... 96
ix
LIST OF FIGURES
Figure 1.1 Illustration of putative stem cell niches in the adult mouse lung ...................... 9
Figure 1.2 Cellular Therapy Decision Tree ...................................................................... 18
Figure 2.1 Amniotic cavity formation .............................................................................. 23
Figure 2.2 Extra-embryonic membranes ........................................................................... 24
Figure 2.3 Kidney amniotic fluid cells ............................................................................. 27
Figure 2.4 Diagram for Amniotic Fluid-Derived Stem Cell isolation .............................. 30
Figure 2.5 Amniotic Fluid Stem Cells .............................................................................. 32
Figure 3.1 Response of the alveolar epithelium to injury/disease .................................... 47
Figure 4.1 IV administration of murine AFSC inhibits fibrotic alveolar and parenchymal
remodeling when injected during either acute or chronic periods following bleomycin
induced lung injury ........................................................................................................... 66
Figure 4.2 IV administration of AFSC attenuates loss of pulmonary function when
injected during either acute or chronic periods following bleomycin induced lung injury69
Figure 4.3 IV AFSC treatment modulates the acute inflammatory cytokine milieu in both
BAL and tissue following bleomycin induced lung injury ............................................... 73
Figure 4.4 AFSC modulate AECII secreted CCL2 in BAL through proteolytic cleavage
by transient MMP2 expression ......................................................................................... 75
Figure 4.5 AFSC are retained within fibrotic lesions and migrate toward increased CCL2
concentrations ................................................................................................................... 79
Figure 4.6 In vitro AFSC co-culture with in vivo injured AECII recapitulates in vivo
CCL2 regulation ................................................................................................................ 82
Figure 4.7 Inhibition of MMP2 in vitro inhibits the ability of AFSC to reduce CCL2
levels following AECII bleomycin injury ......................................................................... 84
Figure 5.1 qPCR analysis expressed as relative mRNA expression for lineage markers in
all 5 hAFSC lines .............................................................................................................. 97
Figure 5.2 Western blot analysis of lung lineage markers ................................................ 98
x
Figure 5.3 Immunofluorescent staining of lung lineage markers within all 5 hAFSC lines99
Figure 5.4 Detection of secreted interleukins, cytokines, growth factors and intracellular
pluripotency markers within AFSC prior to culture on decellularized lung matrices .... 101
Figure 5.5 Decellularization of lung specimens produces acellular matrices ................. 102
Figure 5.6 Acellular matrices provide a culture system for AFSC for up to 28 days ..... 103
Figure 5.7 Detection of secreted interleukins, cytokines, growth factors and intracellular
pluripotency markers within AFSC during culture on decellularized lung matrices ...... 104
Appendix 1 AFSC modulation of the acute inflammatory cytokine milieu in both BAL
and tissue following bleomycin induced lung injury ...................................................... 138
Appendix 2 AFSC modulation of the acute inflammatory cellular populations in BAL
following bleomycin induced lung injury ....................................................................... 139
xi
ABSTRACT
Idiopathic Pulmonary Fibrosis (IPF) is a chronic, progressive and fatal lung disease, with
no effective or definitive treatment other than lung transplantation. Stem cell therapy has
the potential to treat IPF through the specific targeting of therapeutic properties or
compounds to inhibit or delay disease progression and aid in repair at specific sites of
injury. Furthermore, as is the case with most progressive diseases, once the progression
of fibrosis has diminished organ function to the point where transplantation is necessary,
the use of stem cells to create bioengineered organoid units provides a therapeutic
strategy reliant only upon organ reengineering time, and not on donor availability. This
dissertation investigates the use of Amniotic Fluid Stem Cells as a therapeutic
intervention strategy and a candidate cell population for organ engineering.
The potential for amniotic fluid stem cell (AFSC) treatment to inhibit the progression of
fibrotic lung injury has not been described. We have previously demonstrated that AFSC
can attenuate both acute and chronic-fibrotic kidney injury through modification of the
cytokine environment. Fibrotic lung injury, such as in Idiopathic Pulmonary Fibrosis
(IPF), is mediated through pro-fibrotic and pro-inflammatory cytokine activity. Thus, we
hypothesized that AFSC treatment might inhibit the progression of bleomycin-induced
pulmonary fibrosis through cytokine modulation. In particular, we aimed to modulate the
pro-fibrotic cytokine CCL2, which is increased in human IPF patients and is correlated
with poor prognoses, advanced disease states and worse fibrotic outcomes. The impact
of intravenous AFSC given at acute (day 0) or chronic (day 14) intervention time points
xii
after bleomycin injury were analyzed at either day 3 or day 28 post-injury. AFSC
treatment at either day 0 or day 14 post-bleomycin injury significantly inhibited collagen
deposition and preserved pulmonary function. CCL2 expression increased in bleomycin-
injured bronchoalveolar lavage (BAL), but significantly decreased following AFSC
treatment at either day 0 or at day 14. AFSC were observed to localize within fibrotic
lesions in the lung, showing specific targeting of AFSC to the area of fibrosis. We also
observed that MMP2 was transiently increased following AFSC treatment. Increased
MMP2 activity was further associated with cleavage of CCL2, rendering it a putative
receptor antagonist for CCR2, which we surmise is a potential mechanism for CCL2
reduction in BAL following AFSC treatment.
When fibrotic lung injury has progressed beyond the point that a therapeutic intervention
would be effective, organ transplant remains the only viable option. With the scarcity of
donor lungs, researchers have begun investigating the potential for whole lung
engineering. The complexity of the architecture and cell populations contained within
the lung, and specifically the alveolus, present a challenge. Thus, in vitro engineering of
bioartificial lungs require the selection of an appropriate candidate cell population and a
physiologically relevant scaffold. Our preliminary data indicates that AFSC are a viable
candidate cell population that express type I and type II alveolar epithelial lineage
markers as well as stem cell markers. Furthermore, our data demonstrates that we have
developed detergent/enzymatic protocols that efficiently remove all cellular material
from lung extracellular matrices, leaving behind biological scaffolds that AFSC can be
xiii
cultured on. Finally, we demonstrate that throughout culture on these scaffolds, AFSC
secrete growth and angiogenic factors, as well as anti-inflammatory, proliferative and
pluripotent factors, all critical for bioengineering functional organoid units.
Based on the entirety of this data, we have concluded that AFSC have the potential to
inhibit the development or progression of fibrosis in a bleomycin injury model during
both acute and chronic remodeling events and have the potential to repopulate
decellularized lung matrix as an in vitro bioarticifical engineering strategy.
1
CHAPTER ONE: CELL BASED THERAPIES FOR LUNG DISEASE
1.1 Introduction: What are cell-based therapies?
The term cell-based therapy encompasses a large array of therapeutic strategies based
upon a simple principle: that cells, whether endogenous (residing within the organ) or
exogenous (isolated from outside the organ) can be stimulated, manipulated or
transplanted to regenerate or repair a disease state. Cell based therapies have become the
new frontier in regenerative medicine as focus has shifted from ameliorating or restricting
the progression of a disease state to preventing, repairing and recellularizing injured,
damaged and diseased organs.
The lung is a prime target for cellular therapies as it, unlike other organs such as the liver,
has shown little capability for regenerative capacity. Compounding the problems posed
by focusing on the lung as a regenerative target is the complex architecture and diversity
of cell types found within the lung and its varying niches. The target cells for these
therapies, the source of transplanted cells, and the potential therapeutic applications for
these cells are the focus cell based therapeutic research.
1.2 Cellular therapeutic strategies for the lung: challenges and possibilities
The adult human lung has 23 generations of airways, beginning with the trachea and
terminating with the alveolar ducts. When examining the lung as a whole, from trachea
to alveoli, there are distinguishable functional regions, each of which has a distinct
2
anatomy, with characteristic tissue and cell types tailored to perform a specific function.
At present as many as 40 or more phenotypically distinct cell lineages have been
identified in the lung [1]. The larger conducting airways are lined by respiratory
epithelium composed of ciliated columnar epithelium, goblet cells, basal cells, brush
cells, serous cells and neuroendocrine (PNE) cells. Traveling down the airways further,
respiratory bronchioles distribute air to alveoli, which provide a large surface area for gas
exchange and the maintenance of normal arterial oxygen, carbon dioxide and pH.
Respiratory bronchioles are lined by Clara cells and a few ciliated cells while alveoli are
lined by type I and II pneumocytes.
The enormity of cell lineages present within the lungs means that multiple cell types are
affected for any one disease or injury, and these cell types vary depending upon the
location of the affected region. Because of the complex geography of lung tissue and
because the number of different cell types that contribute to each functional compartment
is extensive, it is not logical to assume that a single stem cell type can be used as an
endogenous or exogenous “catch all” therapeutic. Thus there are multiple cellular niches
responsible for supporting these different populations and thus multiple populations of
progenitors must be required to respond to region specific lung injury. At present, each
niche and its role in homeostasis, injury and repair is still not fully characterized.
Therefore, the challenge of deciding which cellular therapy to employ in the complex,
fractally-branched anatomy of the lung without a full understanding of cellular niches
within the lung makes targeting of cellular therapy a major challenge.
3
In addition to the enormity of the different cell types contained within the lung, their
highly diverse functions and abilities to regenerate the litany of various lung diseases and
injuries further complicates the decision to utilize certain types of cellular therapies. For
example, in acute alveolar injury, where closure of breaches in the alveolar surface is
urgent, targeting and supporting the proliferative healing potential of endogenous
progenitor populations makes sense. Thus, disease states resulting from apoptosis and
damage to epithelia such as acute inhalational injury or blast injury could be managed
through increased proliferation and repair by endogenous cells as well as transplantation
of cells endogenous to the lung that can quickly repopulate the damaged area, without
having to be “reprogrammed." However, for other disease states resulting from
developmental deficiencies or chronic depletion of endogenous stem cell pools such as
old age and chronic obstructive pulmonary disease (COPD), a disorder marked by
chronic bronchitis and emphysema, exogenous stem cell therapies are, theoretically at
least, the better approach [2]. Progressive diseases such as pulmonary fibrosis, which
affects the regenerative capacity of alveolar epithelium and results in its replacement with
fibrotic tissue, might benefit from exogenous cellular therapies that may not only
repopulate various cellular niches, but may aid in slowing the progression of tissue
destruction (Table 1.1).
4
Lung Disease Pathology Affected Regions Therapeutic Target
Adult RDS
Inflammation, hypoxemia and impaired
gas exchange
Alveolar epithelium, Capillary
endothelium
Regeneration of epithelia and
endothelium
Asthma
Inflammation, bronchospasm and
airflow obstruction
Airway epithelium,
myofibroblasts, airway smooth
muscle
Reduce inflammatory milieu,
inhibit airway wall remodeling,
inhibit smooth muscle hypertrophy
and hyperplasia
Bronchiolitis obliterans Inflammation and fibrosis of bronchioles Airway epithelium Regeneration of epithelia
Bronchopulmonary
Dysplasia
Necrotizing bronchiolitis and alveolar
septal injury
Alveolar epithelium, interstitial
fibroblasts, capillary endothelium
Reduce inflammatory milieu,
regeneration of alveolar septa and
epithelium
Congenital lung
hypoplasia
Incomplete development of lung
resulting in reduced number or size of
bronchopulmonary segments or alveoli
Alveolar epithelium, interstitial
fibroblasts, capillary endothelium
Generate alveolar septa and 3-
Dimensional alveolar structure
Cystic fibrosis
CFTR mutation resulting in decreased
mucociliary clearance and inflammation
Airway epithelium Delivery of functional CFTR
Neonatal RDS
Insufficient surfactant production in the
lungs
Alveolar epithelium, Capillary
endothelium
Generation of surfactant
production, regenerate epithelia
and endothelia
Pulmonary emphysema
(COPD)
Loss of alveolar integrity and reduction
of ventilation
Alveolar epithelium, interstitial
fibroblasts, capillary endothelium
Generate alveolar septa and 3-
Dimensional alveolar structure
Pulmonary fibrosis
Inflammation and fibrosis of alveolar
tissue
Alveolar epithelium, Interstitial
fibroblasts, endothelium
Reduce inflammation, reduce
alveolar epithelia loss, inhibit
fibroblast proliferation
Sarcoidosis
Inflammation accompanied by
granuloma formation
Epithelium
Reduction of inflammation,
regeneration of epithelia
RDS=Respiratory Distress Syndrome, COPD=Chronic Obstructive Pulmonary Disease, CFTR=cystic fibrosis transmembrane conductance regulator
Table 1.1 Lung Diseases Potentially Treatable with Cellular Therapies.
1.3 What is a stem/progenitor cell?
Following injury or disease, the ultimate goal of any cellular therapy is to restore the
microenvironment, tissue and organ to what is considered a “normal” state. The
restoration to normal physiological function can be achieved through the stimulation of
endogenous stem or progenitor cells, transplantation of stem or progenitor cells or
through the complete decellularization of the lung and recellularization with stem or
progenitor cells. The question then becomes, what type of cells, stem or progenitor cells,
should be used for regeneration?
Stem cells are primitive, undifferentiated cells, which are able to self-renew, while giving
rise to descendants with the distinct characteristics of differentiated cell lineages. In
5
adults, tissue specific stem cells are capable of regenerating normal and damaged tissues
throughout life. Stem cells are also critical in normal development and growth; however,
with age it is hypothesized that a depleted and/or dysfunctional pool of adult stem cells
underlies most degenerative disease states and the aging process [3]. Thus stem cells that
can no longer self-renew are thought to be the underlying cause of various degenerative
diseases characterized by a lack of tissue regeneration following injury. The prevailing
theory for this lack of regenerative capacity is that in adult stem cells, telomere length
decreases with increasing age, which suggests a mechanism for the decreased
proliferative potential of these populations [4]. In contrast, lung stem cells that proliferate
out of control are hypothesized to play a key role in lung cancer [5].
Progenitor cells are cells that are committed to a specific lineage and differentiate into a
specific cell or cell type. Progenitor cells, like stem cells can self renew, however it is
surmised that these have a finite capacity for self-renewal, unlike their stem cell
counterparts. Thus, progenitors are an “intermediate” cell type that is not fully a stem
cell and not terminally differentiated either. These progenitors are thought to occupy
various niches within an organ and repopulate tissue for various reasons (normal growth,
injury or disease). Progenitor cells are also susceptible to injury and disease and are
surmised to be the cause of various disorders [6].
6
1.4 Endogenous progenitors and their niches in the lung
There are a handful of niches within the lung that have been surmised to contain stem or
progenitor cells. Stem or progenitor cell “niches” are highly specialized
microenvironments that maintain the undifferentiated state of stem or progenitor cells
while also specifying their fate. The spatial localization and properties of lung stem cell
niches and the cells residing within them are a major focus of research in lung
regenerative medicine. Although there is still controversy over the existence and
characterization of stem and progenitor cells within the lung, there are various cell
populations that have been described. Most importantly however, is to be mindful of the
fact that most of these characterizations have occurred in the mouse lung, and although
various homologous genes between mice and humans have been described, the extent of
the characterization of human endogenous progenitors is far behind that of their murine
counterparts.
At least 5 putative niches containing progenitor cells have been postulated in the adult
mouse lung (Figure 1.1) [7].
1. Airway submucosal glands (SMG) occur in the trachea. Transplantation
of decellularized trachea into nude mice showed reconstitution of surface
tracheal epithelia from the remaining gland duct and acinar cells found in
airway submucosal glands.
2. In the distal trachea and proximal bronchi, Keratin-14 (K14)-expressing
tracheal basal cells are considered a second nidus of stem/progenitors.
7
When Clara cells are injured with naphthalene in mice, tracheal basal
cells can repopulate tracheal epithelia, including columnar secretory and
ciliated cells, which is indicative of a progenitor-progeny response to
injury [8]. Recently, human trachea has been engineered from the two
main autologous cell types in the trachea, mesenchymal stem cell-derived
cartilage-like cells and respiratory epithelial cells [9]. Additionally, whole
tissue engineered bronchus has been transplanted successfully into a
woman to replace a tubercular, stenotic human mainstem bronchus with a
successfully engineered bronchus made from these stem/progenitor cells
and decellularized matrix [10].
3. & 4. Variant Clara cells (Clara
v
) are believed to be the responsible population
for cell renewal in the distal airways. Two subpopulations of Clara
v
cells
have been identified as being associated either with neuroendocrine
bodies (NEB) or bronchoalveolar duct junctions (BADJ) [11]. Clara
v
cells associated with NEB are in close contact with sensory nerve fibers
and are structurally suggestive of some type of endocrine function. These
cells are long lived and reside in a specialized microenvironment or niche
that allows for their differentiation and proliferation following
naphthalene injury [12]. Naphthalene, which typically ablates Clara cells,
seemingly has no effect on Clara
v
cells, which can repopulate distal
airways following injury. Another population of Clara
v
cells found at the
BADJ has been designated a bronchoalveolar stem cell (BASC). These
8
BASC characteristically express both alveolar surfactant protein C (Sp-
C), crucial in the reduction of surface tension in alveoli, as well as Clara
cell 10kDa secretory protein (CC10), whose dysfunction in implicated in
asthma [6]. Like their NEB counterparts, they are highly resistant to
injury and can readily proliferate and repopulate the distal parts of the
airways following injury.
5. Alveolar epithelial type II cells (AECII) are responsible for generating
fully differentiated type I pneumocytes (AECI) in the alveolus. AECII
produce surfactant proteins and are relatively resistant to apoptosis,
suggesting that this subpopulation of cells could be available to re-
epithelialize injured alveoli [13]. Expression of telomerase, a
stem/progenitor cell marker, is elevated within these cells following acute
oxygen injury [14].
9
Figure 1.1 Illustration of putative stem cell niches in the adult mouse lung. Epithelia
of the adult mouse lung can be divided into four major, biologically distinct trophic units
(trachea, bronchi, bronchioles, and alveoli), each of which encompasses unique types of
airway epithelial cells (epithelia relevant to each unit are shown inside circles). Five
potential stem cell niches for these various trophic units are shown on the right, with
locations of candidate stem cells marked by arrowheads (cells are in red). Stem cells and
niches include the following: 1) an unknown cell type in the SMG ducts of the proximal
trachea; 2) basal cells in the intercartilaginous zones of the lower trachea and bronchi
(these structures may also be associated with innervated NEBs; 3) variant Clara cells
associated with NEBs in bronchioles; 4) Clara cell associated with BADJ; and 5) alveolar
type II cells of the alveoli. Abbreviations: BADJ, bronchiolar alveolar duct junctions;
Clara
v
, variant Clara cells; NEB, neuroendocrine body; SMG, submucosal gland.
Reprinted with permission of the American Thoracic Society. Copyright © American
Thoracic Society. From Liu X, Engelhardt JF. The glandular stem/progenitor cell niche in
airway development and repair. Proc Am Thorac Soc 2008;5:682-688. OFFICIAL
JOURNAL OF THE AMERICAN THORACIC SOCIETY DIANE GERN, Publisher
10
In conjunction with identification of resident lung progenitor cells, are attempts at
isolating and characterizing these cells to better understand their mechanisms of action
both in vivo and in vitro. Successful isolation, purification, and maintenance of a
population termed multipotent lung stem cells (MLSCs) was accomplished recently.
Multipotent cells are capable of giving rise to multiple but limited numbers of cell
lineages. This cell population was shown to generate endothelial, Clara, AECI and
AECII cells, as well as differentiate into mesenchymal cells in vitro [15]. Furthermore,
while these cells showed minimal engraftment in elastase-injured lungs, transplant of
these cells significantly improved survival when compared to non-treated mice. These
data suggest that MLSCs are an excellent target for future research and translational
strategies in human lung disease.
Another recently identified multipotent epithelial stem/progenitor cell population in the
adult mouse lung has been characterized [16]. When cultured in a system mimicking the
normal physiological air-liquid interface of the lung, in this stem/progenitor epithelial
population gives rise to colonies of airway, alveolar or mixed lung epithelial cell
lineages. Furthermore, these multipotent epithelial colony-forming cells self-renew, and
give rise to airway and alveolar epithelial lineage restricted progenitor cells in culture,
and can be resolved from endogenous lung mesenchymal stromal progenitor cells [16].
Finally, a recently published study in the New England Journal of Medicine described the
existence of stem cells within the human lung. Unlike previously identified
11
stem/progenitor phenotypes in the lung, these cells were described as multipotent,
clonogenic and self-renewing. In essence, these cells are closer to the idea of a “true
stem cell” than any other cells identified in the lung to date. These cells, isolated on the
basis of the well-known stem cell marker c-kit, were shown to differentiate into
phenotypes characteristic only of their organ of origin, the lung [17]. Furthermore,
following thoracotomy in mice, transplanted human lung stem cells were reported to
form bronchioles, alveoli and pulmonary vessels. While the initial prospect described in
this study is exciting, caution must be exercised before more rigorous testing including
appropriate controls and characterization have been performed.
The likelihood of needing multiple lung stem and progenitor cell sources for tissue
generation and regeneration highlights the importance of establishing the developmental
potential and regenerative capacity of candidate stem and progenitor cells distributed
along the proximal-distal axis of the lung; the properties of the regional
microenvironmental niches in which they reside and which specify their fate; and, how
they are affected in different lung diseases [18].
The importance of the role of these
progenitor cells during injury and disease cannot be overlooked if treatment strategies
aimed at reinvigorating the regenerative potential of these cells is to be effective. With
the majority of recent research efforts directed at inhibiting aberrant cells from
proliferating, investigations focused on rejuvenating non-proliferating cells demands
further study. Recent studies using gene transfer vectors have shown the potential for
specific targeting within distinct populations in the lung. Rhinoviral infection of cultured
12
tracheal epithelium showed that basal cells were selectively infected [19]. Thus novel
viral targeting of stem cell populations with vectors that provide increased expression
within them could prove an effective strategy for “awakening” resident stem cell
populations.
1.5 Exogenous stem and progenitor cell candidates for cell therapies
1.5.1 Hematopoietic stem cells (HSC) and mesenchymal stem cells (MSC): Stem cells
originating within the bone marrow have garnered much attention in the stem cell field as
these progenitors are not just a phenomenon of early development, but continue
throughout life. These cells, known as hematopoietic and mesenchymal stem cells are
thought to circulate through the blood stream and potentially respond to sites of disease
or injury and participate in repair. Furthermore, HSC and MSC are considered good
candidates for use in cell based regenerative therapies as they have been well
characterized and have shown healing potential when genetically manipulated. However
their ability to efficiently and fully differentiate into lung epithelia appears doubtful
[20,21]. A certain amount of lung repair and regeneration is hypothesized be attributed to
circulating progenitor cells. For example, the introduction of these cells into diseased
tissue has aided in the amelioration of injury and the augmentation of endogenous injury
responses [22-24]. This suggests that these cells may be acting "pharmacologically,"
apparently by modulating the immune or inflammatory homeostatic milieu within the
lung. A logical target for this ameliorating, protective and/or activating effect may be the
endogenous progenitor populations previously described, which could therefore respond
13
to exogenous stem cell administration by effecting repair of damage. The potential for
the use of such progenitor cells is exciting when considering cases such as COPD, in
which patients have experienced destruction of much normal respiratory tissue.
The role that circulating progenitors, HSC and MSC, may play in lung repair has until
recently been largely controversial as many lung diseases are attributed to mesenchymal
abnormalities. As a caveat to this potential benefit of circulating progenitors, it must be
noted that circulating fibrocytes have been identified as a potential major contributor to
the progression of pulmonary fibrosis [25]. Thus the potential of these cells to facilitate
rather than inhibit fibrosis must be considered when balancing the need for therapy
against possibly harmful side effects. These conflicting roles for circulating progenitor
cells, of either mesenchymal or hematopoietic origin, thus require further study and
characterization in order to determine which specific populations contribute beneficial,
versus deleterious, outcomes following administration.
1.5.2 induced Pluripotent Stem Cells (iPS cells) and Embryonic Stem cells (ES cells)
have shown promise as therapeutic agents in other organs, however full exploration of
their potential in lung disease is yet to be tested [26]. Researchers have been successful
in generating airway epithelial cells as well as cells that express epithelial type II cell
markers from ES cells. More recently, AECII cell lines, were established from human
ES cell lines and shown to differentiate in vitro into AECI cells. Furthermore, when
transplanted intratracheally into mice following Bleomycin injury, which induces
14
pulmonary fibrosis, these cells expressed phenotypic markers consistent with AECII and
AECI cells [27]. Such lung cell directed differentiation is a rare event and the
functionality of these cells in vivo has yet to be elucidated [28]. Human ES cells have
not been fully investigated in the lung largely due to ethical considerations and
governmental regulations. Attempts to use iPS cells instead of the more controversial ES
cells has also raised concerns after tumorgenicity was observed in mouse models [29].
Refinement of the iPS procedure to inactivate or remove the oncogenes used to drive de-
differentiation of these lines is underway, and the potential for these cells as a therapeutic
source remains high.
1.5.3 Fetal associated stem cells: Amniotic fluid stem cells (AFSC), cord blood stem cells
and placental stem cells are also an attractive candidate for use in lung repair. Their use is
not constrained by the ethical considerations and potential hazards associated with
embryonic stem cells, and they have been shown to express lung epithelial markers in
vivo [30]. Human amniotic fluid stem cells (hAFSCs) can be derived from discarded
amniocentesis specimens and thus circumvent the ethically charged arguments about ES
cells. Similarly, placental and cord blood derived stem cells can be collected at birth and
stored for potential future use. These fetal associated cells, are traditionally characterized
as mesenchymal and have shown immunomodulatory and regenerative effects in various
injury models.
15
hAFSCs in particular, are pluripotent, giving rise to all three germ layers in mouse
chimeras as well as in vitro under carefully controlled conditions [31]. hAFSCs have
also shown the ability to differentiate into various tissue types, such as the embryonic
kidney or lung, when placed into the correct microenvironment, and have not been shown
to produce teratomas as ES cells do [32]. In preliminary studies, it has been found that
hAFSCs can incorporate into mouse embryonic lung and express human lung epithelial
cell markers. Following lung injury in nude mice, it has been shown that after
intravenous injection of the cells, they become trapped in the lung and remain at sites of
injury [30]. hAFSCs persist in the lung after injury, but decrease over time and their true
efficacy and potential in lung repair has yet to be elucidated.
1.6 Engineering whole lung regeneration
When the prospect of repairing the lung is out of the question, researchers have begun
investigating the potential for whole organ regeneration. This type of regeneration is
based on the theory that scaffolding, whether obtained from decellularized donor lungs or
bioengineered materials, can be seeded with various cell types and cultured to make an
entire functional organ, such as the lung. Recently, a whole lung decellularization method
and subsequent tissue engineering using neonatal lung epithelia has been reported [33]. It
has long been known that seeded epithelial cells can repopulate alveolar structures within
denuded lung matrix. Seeding of pulmonary endothelium and epithelium was
successfully accomplished on a decellularized rat lung. Following orthotopic
transplantation, the re-engineered lung was capable of gas exchange as shown through
16
blood gas analysis [34]. Bioengineered scaffolds on the other hand have been utilized to
create larger sections of the lung, such as the trachea and bronchus, however, the finite
architecture of alveoli and terminal bronchioles remains a challenge for engineers. While
decellularized lung matrix seems to currently be the most promising scaffold for whole
lung regeneration, artificial matrix constructs need further investigation and may
eventually serve to maximize regeneration of alveolar integrity.
1.7 Production and delivery of cellular therapies
The primary delivery choices for cell therapy into the lung are either down the airway or
through the pulmonary arterial circulation. Intravenous injection of therapeutic cells has
been used in numerous studies as the lung acts as a cellular strainer and thus traps various
cells that are too big to continue on through the pulmonary circulation. A caveat may be
that once entrapped in the lung microvasculature, regenerative cells may not be able to
efficiently cross the endothelial barrier to home and lodge at the site of injury. Also
described is the aerosolization or discrete placement of cells within the lung, either as a
bolus of cells or embedded within a dissolving matrix. As with intravenous delivery,
cells may become trapped in the alveoli and may not be able to migrate easily to
mesenchymal or endothelial sites of injury. Furthermore, while both routes offer
relatively efficient initial deposition of cells, in the absence of ongoing injury, long term
tissue uptake, retention and differentiation is as yet relatively inefficient in vivo. By both
routes of administration, the bulk of exogenous cells are cleared either into the local
17
lymphatic tissue or via the circulation to the liver and spleen thus, using these approaches
to achieve significant replacement of damaged lung or airway remains impractical..
There are also a number of practical considerations, which must be addressed when
devising cellular therapies for lung regeneration and repair. Robust cell separation
protocols for the prospective isolation of highly enriched stem and progenitor cell targets,
capable of repairing and regenerating affected lung cell lineages will need to be devised.
Ex vivo cell culture systems able to expand stem cells without compromising their
regenerative potential will need to be developed; because stem cell fate is dictated by
extrinsic signals provided by their tissue microenvironment, preconditioning strategies
and routes and mode of delivery will need to be devised to optimize their engraftment
and harness their regenerative capacity.
1.8 Cellular therapy versus whole lung engineering
The decision to use the approach of cellular therapy versus whole lung engineering when
approaching lung disease models is as complex as trying to categorize various lung
diseases in comprehensive review. The timing of a diagnosis coupled with the extent of
the injury or disease is essential when a clinician is tasked with determining an
appropriate course of treatment (Figure 1.2). Cellular therapies are advantageous when
minimally invasive procedures are favored. Additionally, cellular therapies are more
attractive than whole lung engineering when repair of injury can be achieved by utilizing
remaining lung tissue or the patient has sufficient remaining lung function to withstand
18
prolonged therapy that may require the growth of cells and secretion of specific
therapeutic factors by the transplanted cells. Whole lung engineering with a patient’s own
scaffolding, or donor scaffold, and self-progenitor cells is likely to be less immunogenic
for the patient than transplanting non-self lungs. Furthermore, patient survival rates
would be improved as exact donor matches would be unnecessary, and transplantation
rates would only be limited by donor lung scaffold availability and organ re-engineering
time.
Figure 1.2 Cellular Therapy Decision Tree.
Lung Disease/Injury
Acute
Extent of
Injury
Enough
functional lung
tissue remaining
to participate in
repair
Endogenous or
exogenous
cellular therapy
Not enough
functional lung
tissue to
participate in
repair
Exogenous cell
therapy
Whole lung
engineering
Chronic
Timing of
Diagnosis
Enough
functional lung
tissue remaining
to participate in
repair
Type of
Disease/
Injury
Genetic
Exogenous
cellular
therapy
Exposure
Endogenous
cell therapy
Progressive
Endogenous
or
exogenous
cell therapy
Viral/
Bacterial
Endogenous
or
exogenous
cell therapy
Not enough
functional lung
tissue to
participate in
repair
Whole lung
engineering
Exogenous
cell therapy
19
1.9 Areas of agreement
Lung researchers have expended considerable effort in attempting to come to a consensus
regarding the potential for cellular therapies within the lung. It is fair to say that
researchers and clinicians alike agree that current treatment strategies are simply not
satisfactory when dealing with progressive lung diseases. Thus, cellular therapies have
been agreed upon as one important new frontier in lung research and disease treatment.
Furthermore, it has been agreed that various different cell types will need to be utilized to
contribute to the population and repopulation of the respiratory architecture.
1.10 Areas of controversy
Attempts at characterizing individual progenitor populations, their designation as “stem
cells” versus “progenitor cells”, the identification of “niches”, the potential of
engraftment of exogenous populations and their efficacy is where many lung researchers
disagree. The existence of stem cells as they are classically defined has recently been
called into question, with some scholars asserting that stem cells do not exist independent
of their niches, and are in fact driven to behave as stem cells only when they are within
these niches. This calls into question the identification of the endogenous stem cells
within the lung discussed herein as being cells that are induced to act only in response to
injury, disease or the need to repopulate surfaces, rather than cells that can be
characterized by the intrinsic properties of "stemness." In addition to these technical and
biological questions, a layer of ethical controversy can be added when one considers the
possibility of exogenous stem cells differentiating into unwanted and potentially
20
deleterious tissue. Finally, there has been much disagreement over the definition of what
constitutes “engraftment” when relating to transplantation of endogenous and exogenous
stem cells.
1.11 Future challenges
The first hurdle that must be overcome in the area of endogenous stem/progenitor cells
within the lung is the identification of consensus markers that can be used to identify
these cells. Secondly, robust functional assays are needed that test the ability of these
cells to either self renew or proliferate and differentiate. Finally, in vivo assays are
needed to measure how the functional ability of endogenous stem cells is impaired
following injury and in the diseased lung, and how endogenous and engrafted stem cells
are recruited to repair the lung. While those in the field recognize the exciting potential of
cellular therapies for lung diseases, further work to characterize the optimal cell
populations and therapeutic strategies and the safety and efficacy of the approaches will
be required.
21
CHAPTER TWO: AMNIOTIC FLUID STEM CELLS
2.1 Introduction
The amniotic fluid or liquor amnii, was first isolated and studied during the beginning of
the 20
th
century [35]. More recently, in the 1960s and 1970s there was an increased
interest in characterization and culture of the cells contained in the amniotic fluid [36,37].
Nevertheless, most all of these studies were directed at using amniotic fluid, and the cells
contained within, for determining the health of the fetus during development, or to
provide a general characterization of the amniotic fluid. Although the discovery of stem
cells, in particular bone marrow stem cells, occurred in the 1960’s, it was not until
recently that the possibility of isolating stem cells from the amniotic fluid was
investigated. Amniotic fluid stem cell isolation and characterization is therefore fairly
recent, dating back to the early 1990’s [38].
The study of amniotic fluid-derived stem cells (AFDSCs) has captured the attention of
researchers and clinicians for several reasons. First, AFDSCs can be collected during
amniocentesis and isolated from material that would be otherwise discarded. Therefore,
their use is not subject to the ethical debate that surrounds the use of embryonic stem
cells. Second, like other fetal derived stem cells, storage of AFDSCs is easy and achieved
at minimal costs. AFDSC populations can be easily expanded, and have shown the
capability of being stored over long periods of time with no adverse effects [39].
Furthermore, the “banking” of AFDSCs from developing fetuses, may guarantee a source
22
of stem cells with a matching immune profile to that of the recipient. Most importantly,
the extensive characterization of a specific subset of AFDSCs positive for the marker c-
kit
+
[31], have displayed no tumor formation following transplant into an animal model,
even after several months [30]. These cells, known simply as amniotic fluid stem cells
(AFSC) have been at the forefront of AFDSC research and will be discussed in depth
later. Finally, as a source of stem cells collected before birth AFDSCs may become an
invaluable source of stem cells for direct treatment of various genetic disorders treatable
in utero [40].
The potential applications and implications of AFDSCs in regenerative medicine and
therapeutic treatments are significant, however; AFDSC research is still in its infancy and
much work is required to properly characterize AFDSCs and determine their
effectiveness. In this chapter, we describe the different AFDSCs that have been isolated
to date, list their characteristics, and provide an overview of the different organs in which
AFDSCs have been used in vitro or in vivo to develop this stem cell population into a
viable therapeutic strategy.
2.2 The amniotic cavity
The amniotic fluid is contained in the amniotic cavity that, in humans, starts forming as
early as seven days post fertilization, and is delimited by a membrane called amnion
(Figure 2.1). The formation of the amniotic cavity is a result of the cavitation of the
epiblast. The amnion is formed by the cells of the epiblast, by the side facing the
23
cytotrophoblast. This is the first appearance of the amniotic ectoderm, and at this stage it
is still a continuum of the portion of the epiblast that will form the embryo. The amnion
formation is completed at fourteen days post fertilization and is constituted of two layers:
the amniotic ectoderm (inner layer facing the amniotic fluid) and the amniotic mesoderm
(outer layer). The amnion has the important function of protecting the embryo and
controlling the composition and the volume of the amniotic fluid. In humans, after
seventeen weeks of gestation the amnion becomes surrounded and fused with another
membrane, the chorion, and is therefore incorporated into the placenta. At the beginning
of the formation of the amniotic cavity, active transport of solutes from the amnion,
followed by passive movement of water, comprise the amniotic fluid.
Figure 2.1 Amniotic cavity formation. Twelve days post fertilization the human
amniotic cavity is delimitated by the amnion (that at this stage is composed by the
amniotic ectoderm) and the embryonic ectoderm (left). In the 7.5-day mouse embryo
(right) the amnion is formed by the amniotic mesoderm and the amniotic ectoderm.
In mice the amniotic cavity starts forming at embryonic stage E0.5 as a result of apoptotic
events in the epiblast. At this stage, there is still the presence of a proamniotic cavity and
24
the amnion that will start forming during gastrulation, is not yet defined. At
approximately day E7.5 the amniotic cavity is formed and one day later the embryo starts
the rotation process. At the end of the rotation, the embryo will be surrounded by the
amnion. Surrounding the amnion are two more membranes, the visceral yolk sac and
most externally the parietal yolk sac [41] (Figure 2.2). These membranes represent three
distinct layers surrounding the mouse embryo. Differently from humans, in mice, the
amnion does not fuse with the chorion and is not included in the placenta.
Figure 2.2 Extra-embryonic membranes. In mammals the embryo is immersed in the
amniotic fluid contained inside the amniotic cavity. In human (left) the cavity is delimited
by the amniotic ectoderm and the amniotic mesoderm that constitute the amnion, and by
the chorion. The amniotic ectoderm is in direct contact with the amniotic fluid. In mouse
(right) the amnion is surrounded by two extra membranes, the visceral and parietal yolk
sac.
2.3 The amniotic fluid
The amniotic fluid is the liquid present in the amniotic cavity and is constituted of
approximately 98% water. The volume and composition of amniotic fluid changes
continuously during the different gestational stages. The volume of the amniotic fluid at
25
the beginning of the pregnancy is multiple times the volume of the fetus, but at the end of
gestation, at forty weeks, it will represent only a quarter of the volume of the fetus. Early
during development, when the fetus has not yet started urination and deglutition, the
plasma from the mother is surmised to play an important role in the composition of the
amniotic fluid, and even though the mechanism is not completely understood, active
transport of solutes is probably present between the amnion into the amniotic cavity,
therefore creating a gradient for water recruitment [42]. The exchange of fluid through
the skin that occurs until keratinization is also an important contributor to the osmolarity
of the amniotic fluid. After keratinization, urination, swallowing and secretion due to
breathing events also contributes to the composition of the amniotic fluid. Urine starts to
be part of the composition of the amniotic fluid at about eight weeks and its amount will
increase during gestation, reaching a flow rate of up to 900 ml/day at the end of gestation
[43]. Similarly, at approximately eight weeks, the fetus begins swallowing and secreting
material including lung fluid and urine. Secretion of lung fluid is due to an active
transport of chloride through the epithelium of the lung [44]. Sampling of amniotic fluid
at later stages of the pregnancy is used to monitor lung development via the presence or
absence of surfactant lipids and proteins secreted into the amniotic fluid.
The cells present in the amniotic fluid have both embryonic and extraembryonic origins.
Approximately forty years ago, researchers attempted to characterize these cells by
cytological and biochemical parameters [45]. Early characterization distinguished four
epithelial cell types in the amniotic fluid: large eosinophilic cells, large cyanophilic cells,
26
small round cyanophilic cells, and polygonal eosinophilic cells [36]. Today we know that
most of the cells of the amniotic fluid are derived from the skin, digestive, urinary and
pulmonary tracts of the fetus and from the surrounding amnion (Figure 2.3). We also
know that the proportion and type of cells changes continuously during the different
gestational stages. Some cells may also be derived from the mother, passing through the
placenta into the fluid itself. The size of the cells contained in the fluid can range from
6um to 50um and the shape can vary notably from round to squamous in morphology
[46].
27
Figure 2.3 Kidney amniotic fluid cells. Amniotic fluid contains cell populations
derived from several different tissues. Pictured above is a population of cells isolated
using kidney specific markers.
2.4 Amniotic Fluid-Derived Stem Cells (AFDSCs)
AFDSCs belong to the group of stem cells present in extra embryonic tissues; all sharing
the feature of belonging to material that is discarded after birth or that can be collected
28
during amniocentesis. Besides the amniotic fluid, the amnion, umbilical cord and
placenta have shown to contain stem cells that can be isolated at birth [47-50].
The first studies of AFDSCs, were completed using mesenchymal amniocytes isolated
from sheep. These cells showed the ability to expand in vitro and to integrate into a
scaffold [51]. In the following years, the identification of cells expressing the marker
Oct4 [52], or co-expressing Oct4, CD44 and CD105 [53] were discovered in amniotic
fluid. More recently a clonal population of AFDSCs derived from human and mouse
were isolated and characterized [31]. These cells named AFSCs, were isolated through
positive selection for the marker CD117 (or c-kit), and represented 1% of cells derived
from amniocentesis. AFSCs express the marker of “stemness”, Oct4, and the embryonic
stem cell (ESC) marker SSEA-4. Furthermore AFSCs express markers characteristic of
mesenchymal and neural stem cells such as CD29, CD44, CD73, CD90, and CD105.
Interestingly, these cells are negative for markers of hematopoietic stem cells such as
CD34 and CD133.
Recently, a screen for the expression profile of cells present in the amniotic fluid was
reported [39]. This screening analyzed cells obtained from human amniotic fluid
between gestational weeks 15 to 20 and showed that markers such as Oct4 and CD117
are stably expressed during gestation. Furthermore, while markers for ectoderm are stably
expressed during gestation, markers for the early endoderm and mesoderm are more
abundant during early gestation and tend to disappear after 17 to 18 weeks. During the
29
same time, organ specific markers start to become highly expressed. A full proteome
analysis [54] using bi-dimensional gel electrophoresis and mass spectrometry, has
allowed the identification of specific proteins expressed in the cells present in the
amniotic fluid. This analysis has confirmed that amniotic fluid contains a heterogeneous
population of cells, both differentiated and with characteristics of stem cells. In the
following paragraphs a detailed description of the approaches used to differentiate
AFDSCs into various lineages is presented.
2.5 Collection of amniotic fluid for Amniotic Fluid Stem Cell isolation
When considering the use of amniotic fluid stem cells for regenerative medicine and
various therapeutic interventions, clinicians and researchers agree that the ease of
amniotic fluid stem cell isolation and culture make them attractive candidates for further
research and development. As mentioned previously, amniotic fluid stem cells are
isolated from samples of amniotic fluid collected during routine amniocentesis. This
routine procedure occurs during weeks 16-20 of a pregnancy, where approximately 10-20
milliliters of amniotic fluid is collected and split into two samples [55] (Figure 2.4). One
sample serves as the test sample to screen for genetic and gestational abnormalities, while
the other sample serves as a back up. When the back-up sample is no longer needed,
some diagnostic laboratories donate this “medical waste” to research laboratories for
stem cell isolation and further research. Throughout this entire process, neither the
mother, nor the fetus is harmed, making the collection of these cells ethically neutral.
30
Figure 2.4 Diagram for Amniotic Fluid-Derived Stem Cell isolation.
2.6 Tissue engineering from Amniotic Fluid-Derived Stem Cells
The use of AFDSCs for the treatment of congenital anomalies has great potential, but in
most cases is still far from clinical applications. Nerveless there is at least one case in
which cells derived from amniotic fluid have been successfully used for tissue
engineering. Mesenchymal cells isolated from amniotic fluid have been expanded in vitro
using a chondrogenic medium and than seeded into a biodegradable scaffold and
maintained in a rotating bioreactor [56]. The cells used in this report were not specifically
analyzed for pluripotency or selected for specific markers, and were considered
31
progenitor cells by the authors. Being a mixed population of cells they likely contained
both committed lineages and AFDSCs, but most importantly they were able to
differentiate into cartilage in vitro into a three-dimensional scaffold and maintain these
characteristics for as long as fifteen weeks.
2.7 Collection, isolation and culture of c-kit
+
Amniotic Fluid Stem Cells
Within amniotic fluid are a menagerie of cells previously described as AFDSCs, however
approximately 1% of the cells contained within the fluid have been identified and
designated as amniotic fluid stem cells (AFSC) (Figure 2.5). AFSCs represent the most
characterized clonal population of pluripotent stem cells isolated from amniotic fluid.
AFSCs can be isolated by immunoselection with magnetic microsphere or FACS for the
receptor for stem cell factor (c-kit or CD117). After isolation AFSCs will grow slowly
for about one week (this phenomenon differs in AFSCs isolated at different gestational
stages), and will then start to proliferate faster following this initial ‘lag-phase’ [46].
AFSCs grow in absence of feeder layer when plated on Petri dishes and have a doubling
time of about 36 hours [31]. The isolated population can then be cultured quite readily
on plastic or glass. If maintained at a sub-confluent state, AFSCs do not differentiate.
Clones should be cultured in medium containing α-minimal essential medium
supplemented with 20% Chang-B and 2% Chang-C solutions, 20% fetal bovine serum
(FBS), 1% L-glutamine, and 1% antibiotics. Clones should be periodically monitored for
the presence of a correct karyotype, and for the expression of specific markers such as
Oct4, SSEA4, CD29, CD44, and the absence of markers such as CD45, CD34, and
32
CD133 (see De Coppi et al., 2007 for a complete list of specific markers). AFSCs are
pluripotent and can be differentiate in vitro into several lineages [31,46]. Numerous
groups have reported the high renewal capacity of these cells without differentiation or
loss of telomere length [39].
Figure 2.5 Amniotic Fluid Stem Cells. Human amniotic fluid stem cells (left) and
mouse amniotic fluid stem cells (right) that were isolated via selection for the surface
marker CD117. Both cells have similar phenotypes.
2.8 Why use AFSC in Regenerative Medicine?
When selecting a stem cell population for use in a regenerative or therapeutic capacity,
there are a myriad of factors that need to be considered. The pluripotentiality, the ability
of the cells to differentiate into different germ layers and tissue types, is of fundamental
importance if one is isolating cells to treat diseases or developmental deficiencies in
which progenitor cells within the patient are compromised or overwhelmed.
Additionally, the plasticity of the cells and their ability to differentiate to repopulate
33
different populations within an organ, and repopulate them correctly is crucial.
Furthermore, the behavior of the cells after injection must be carefully studied and
characterized. Tumorogenicity, immunogenicity and the propensity to form teratomas
and further exacerbate a disease state can rule out various cellular therapies simply due to
risk. To date, amniotic fluid stem cells have demonstrated the ability to meet all of these
criteria and behave remarkably well in a regenerative and therapeutic capacity.
Amazingly pluripotent, less immunogenic, and not prone to teratoma formation, AFSCs
have quickly risen near the top of the list of stem cell therapies to continue developing.
Furthermore, recently induced pluripotent stem (iPS) cells have been prepared from cells
derived from amniotic fluid (AF-iPS), and have shown high efficiency of transformation
and colony formation after just six days [57]. Although not fully understood, this is
probably due to the presence of an epigenetic status closer to the embryonic state [58].
Reprogramming of somatic cells using the four specific factors, Oct4, Sox2, Klf4, and c-
Myc has the potential to provide pluripotent stem cells specific for patients, thus AF-iPS
seem to be more easily reprogrammed to pluripotency compared to adult cells or cells
from neonates.
2.9 Differentiation of Amniotic Fluid Stem Cells for regenerative medicine
C-kit positive amniotic fluid stem cells are pluripotent and have been successfully
differentiated into all three germ layer cell types: endoderm, ectoderm and mesoderm.
From these pluripotent cells, various phenotypes have been derived in vitro. Osteogenic,
endotheilial, hepatic, neurogenic, adipogenic and myogenic progenitor cell lines are a few
34
of the lineages derived to date. Derivation of these lines has been verified by
morphogenesis, phenotypic analysis and a litany of biochemical assays for characteristic
of each cell type. Culture and manipulation of these cells into various progenitors has
become so streamlined, that various standard protocols have been established [59].
Although a significant milestone, differentiation of AFSC into various lineages in vitro is
quite distinct from the in vivo potential, use and efficacy of these cells. Transplantation
of these cells into a living system, or the use of these cells to create a functional organ
hinge on the ability of these cells not simply to survive in vivo, however; success is
dependent on the physiological functionality of these cells to perform within the
anatomy. The future of regenerative medicine and cellular therapy hinges on this
principle, and not surprisingly, AFSC have also shown remarkable capabilities in vivo in
numerous organs.
2.9.1 Hematopoietic System
AFSC expressing CD117
+
and Lin
-
, derived from both human and mouse, have been
shown to have hematopoietic potential [60]. These cells were capable of differentiating
into erythroid, myeloid, and lymphoid lineages in vitro as well as in vivo, in the
peripheral blood of irradiated mice. Furthermore, single cells analysis was able to assess
the expression of several genes important during different stages of hematopoietic
differentiation.
35
2.9.2 Brain
A fully mature neural differentiation remains to be tested for cells derived from amniotic
fluid. Neural differentiation was fist reported during the initial identification of AFSCs
[31]. Subsequently, a study for the differentiation of AFSCs into dopamine neurons [61],
showed that AFSCs express specific markers of neural progenitors and immature
dopamine neurons, but were unable to fully differentiate in vitro or in vivo. Analyzing
other cell lines isolated from amniotic fluid [62] it was shown that phenotypic
characteristics of dopaminergic neurons are present, while markers for other neurons, like
cholinergic, GABAergic, and adrenergic were absent or had a weak expression.
2.9.3 Bone
AFSC cultured with an osteogenic medium, can secrete alkaline phosphatase and produce
mineralized calcium, characteristic of functional osteoblasts. Furthermore, when
implanted into an immunodeficient mouse, AFSC where able to produce mineralized
tissue in vivo [31]. A comparison between AFDSCs and bone morrow-derived stem cells
(MSCs), has shown that while MSCs undergo a faster differentiation, AFDSCs can
maintain and increase the mineralization for a longer period [63].
2.9.4 Kidney
AFSC therapy in the kidney is progressing quickly and is arguably at the forefront of
AFSC research. Research groups using AFSC in kidney have not only been able to
demonstrate the ability of AFSC to populate the kidney and form renal structures, but
36
also to protect the kidney during injury and aid in the regeneration of renal tissue. The
groundbreaking studies, which follow, paved the way for much of the other organ
specific experimentation, in particular, that of the lung.
In the embryonic kidney, AFSC have been shown to differentiate into tubular and
glomerular structures and express characteristic kidney cell markers and genes [32]. In
this study, metanephric kidneys were isolated from embryonic mice, microinjected with
approximately 1000 CM-dil labeled c-kit positive AFSC and placed on a membrane for
cultivation in an incubator. What is remarkable is that even though the embryonic kidney
was not fully formed at the beginning of the experiment, labeled AFSC were seen to
integrate into developing C and S-shaped structures at day 5, and at day 6, integrated into
tubular and glomerular structures. Reverse transcriptase-PCR for human kidney specific
genes, not previously expressed by the AFSC, identified expression of ZO-1, claudin and
glial-derived neurotrophic factor. This experiment showed the ability of AFSC to survive
within developing tissue, engraft into that tissue, differentiate into the appropriate cell
type and aid in the population of an organ.
Furthermore, it has recently been discovered that AFSC injected into the acutely injured
kidney stimulate the release of anti-inflammatory cytokines and attenuate pro-
inflammatory signaling greatly reducing apoptosis and allowing for proliferation and
repopulation of injured epithelia [64]. In this study, nude mice, deprived of water for a
period of 22 hours, were given an intramuscular injection of a 50% hypertonic glycerol
37
solution in water. This type of injury induces rhabdomyolysis-related acute tubular
necrosis (ATN) ultimately resulting in renal dysfunction. Following intrarenal injection
of 1.2x10
6
cells, AFSC were observed, via luciferase, to persist at the site of injury most
notably at 48 and 72 hours, with persistence in the kidneys for up to 6 days.
Additionally, analysis of the cytokine milieu showed the markedly different expression
patterns of cytokines at 14 days post transplant. Mice with ATN only, and no AFSC
transplant, showed a general trend of increased pro-inflammatory cytokines and
decreased anti-inflammatory cytokine expression. On the other hand, mice with ATN
and intra-renal injection of AFSC demonstrated that the anti inflammatory cytokines
increased over the 14 day study period, while pro-inflammatory cytokines decreased.
In another study after glycerol-induced acute kidney injury [65] a comparison between
mesenchymal stem cells (MSCs) and AFSCs has shown that while MSCs where mainly
inducing proliferation, AFSCs had an antiapoptotic effect. Thus, these data suggests that
AFSCs responds in a paracrine manner in response to injury and/or stress, and
modulation of immune signaling is what contributes to the alleviation of symptoms
associated with the injury.
Most recently in a Col4α5
-/-
deficient Alport mouse model, treatment with AFSC
increased the mean lifespan of treated cohorts by 20% while significantly inhibiting the
progression of interstitial renal fibrosis [66]. Furthermore, following AFSC treatment,
significant reduction in pro-fibrotic pro-inflammatory cytokines was observed providing
38
strong evidence for the therapeutic potential of AFSC in chronic injury models of disease
as well.
2.9.5 Lung
In utero, the developing lungs of the fetus are filled with fetal lung liquid, which is
actively secreted into the amniotic fluid. In the late gestational period, surfactant
produced by the fetal lungs contributes to the composition of amniotic fluid and can be
measured to determine the developmental stage of the surfactant system within the fetal
lungs. Thus, it makes sense that when looking for regenerative therapies for lung tissue,
AFSC are a logical source.
In our preliminary transplantation studies, it was found that c-kit positive AFSC can
incorporate into mouse embryonic lung and express human lung epithelial cell markers
[30]. In the same study, following naphthalene injury in nude mice, and intravenous
transplantation of 1x10
6
AFSC, cells were observed to preferentially remain at the site of
injury when compared to uninjured controls when visualized via luciferase assay.
Additionally, following oxygen injury in the lung it was observed that AFSC appear to
exhibit alveolar epithelial type II phenotypes, widely surmised to be a lung epithelial
stem/progenitor cell, suggesting that once in the lung these cells are stimulated to
differentiate in response to injury. Furthermore, in vivo, the efficiency of AFSC
diapedesis, integration and expression in upper and lower airway epithelia is increased
following injury. After oxygen injury, AFSC were observed to be taken into the SP-C
39
positive alveolar epithelial lineage, whereas after naphthalene injury AFSC are taken up
into the CC10-positive Clara cell lineage. AFSC presence persisted in the lung after
injury, but decreased over time. Although integration into the adult lung following injury
is a relatively rare event, additional therapeutic mechanisms displayed by these cells,
such as the modulation of the inflammatory milieu and their differentiation into type II
lineages demonstrate great potential in the stimulation of lung repair mechanisms.
Lung researchers have also begun investigating the potential of seeding AFSC on a
scaffold to regenerate tissue for transplantation. Due to the overwhelming shortage of
donor lungs, and the inability of modern medicine to effectively treat or halt many
progressive lung diseases such as idiopathic pulmonary fibrosis, research focus has
shifted to the bioengineering of functional lung tissue. Decellularization of lungs, where
all cells are removed from the extracellular matrix of an organ, has become an
investigational target. In 2010 a whole lung decellularization method and tissue
engineering study using neonatal lung epithelia was reported [33]. What is remarkable
about this study is that while it has long been known that epithelial cells seeded on a
decellularized lung matrix were capable of forming alveolar epithelia, this study
demonstrated the functionality of the regenerated tissue. The decellularized, repopulated
and regenerated lungs were transplanted into a rat and were able to support short-term
perfusion and gas exchange. In another study, researchers were able to seed not only
epithelium, but also endothelium as well on a decellularized rat lung. Following
transplantation, blood gas analysis of the engineered lung demonstrated that the lung was
40
capable of gas exchange [34]. Thus decellularized lung matrix seems currently to be the
most promising scaffold for whole lung regeneration and the possibility of using an
autologous source of stem cells such as AFSC to repopulate the scaffold could have great
potential in the future.
2.9.6 Heart
The use of AFSC as a regenerative therapy for cardiac disease and congenital disorders
has shown the efficacy of transplanted cells providing both cardio protective potential, as
well as the engineering of various cardiac components such as valves and tissue [67-69].
The engineering of heart valves, obtained from normal human amniotic fluid samples,
sorted via positive selection for the CD133 molecule, was elegantly demonstrated in 2007
[68]. Both CD133 positive and negative cell populations were cultured in media that
caused differentiation towards endothelial phenotypes. CD133
+
cell populations showed
the ability to produce functional endothelial cells indicated by the expression of eNOS
and CD141, while CD133
-
cells displayed a more mesenchymal phenotype. CD133
-
cells
were then seeded on biodegradable PGA leaflets that were positioned within a mold to
form a valve structure. After 14 days, CD133
+
cells were seeded onto the scaffold as
well. While regeneration of both extracellular matrix and endothelial layers were
generated, functional testing revealed that the heart valves were sufficiently functional
only under low-pressure conditions. This failure to perform at physiological levels was
not due to the scaffold material, which displayed linear properties prior to being seeded
with cells, but instead was a result of the incomplete formation of collagen suggesting
41
that the method of seeding and culture upon the biodegradable scaffolding needs to be
optimized further to be able to transplant these engineered valves into patients.
In an acute myocardial infarction model, ischemia, produced via ligation of the left
anterior descending coronary artery, was followed with intravenous transplantation of
AFSC and reperfusion of the heart for 2 hours. Animals treated with 5x10
6
cells
intravenously, showed a significant decrease in infarct size and number of apoptotic
cardiomyocityes when compared to control animals administered saline alone [67].
Staining to determine the localization and viability of transplanted AFSC showed that
two hours post transplant, cells localized to the lung, spleen and heart. AFSC within the
heart co-stained for epithelia vWf and α-SMA, suggestive of the potential of these cells
to commit to endothelium and smooth muscle following transplant. Long term retention
and engraftment in the injured myocardial tissue did not occur however. The secretion of
thymosin beta 4 in vitro, a cardio protective factor, suggests that the transplantation of
AFSC in this model exert a paracrine effect.
2.10 Conclusion
The studies outlined in this chapter demonstrating the capability that AFSC have shown
in vitro and in vivo show that AFSC are viable targets for regenerative medicine and for
future therapeutic treatment strategies. Although the relatively early stage of AFSC
research limits a full understanding of the behaviors, properties and characteristics of
42
these fascinating cells, research to date demonstrates two important mechanisms of action
that need to be investigated further.
First, AFSC have the potential to serve as an in vivo treatment to stimulate endogenous
cell populations, repopulate injured tissue or ameliorate inflammatory or disease states.
These properties are advantageous when dealing with disease or injury states in which
there is enough functional tissue remaining in an organ to drive repopulation. The only
caveat to endogenous cellular stimulation is that the remaining tissue (that is being
stimulated) must be functional, meaning that it is free of genetic disorders or mutations.
If remaining tissue within an organ meets these standards, exogenous AFSC
transplantation can be used to stimulate endogenous progenitors to repopulate, protect
progenitor or other cell types from further injury, or AFSC may be driven to differentiate
to repopulate this tissue, as was indicated in the aforementioned embryonic studies.
Second, AFSC have the potential to engineer whole organs in vitro to be transplanted into
a recipient. This strategy is advantageous in situations where, for whatever reason,
enough functional tissue does not remain to repopulate with a cell transplant. Whole
organ re-engineering, perhaps the holy grail of regenerative medicine, involves a
symphony of factors, events and coordinated expression patterns to form intricate niche
structures including endothelium, epithelium, extracellular matrix and so on. The
engineering of a whole organ will in fact require a much deeper understanding of these
cells as signaling cascades and response elements need to be coordinated to engineer
43
every cell type within a specific organ. However the recent findings of Kajtsura et al
(2011) support the our notion that the genome within a single stem cell type may prove to
be sufficiently plastic to simultaneously derive all of the cell lineages required for
complex organ repair or engineering.
AFSCs therapy in the clinic may still be a few years in the future, however it is not too
early to begin investigating the potential of these cells within various conditions. A “one
size fits all” strategy for these cells in clinical applications is not likely to be effective.
44
CHAPTER THREE: MODULATING THE ALVEOLAR MILIEU TO ENHANCE
RESOLUTION OF FIBROTIC LUNG INJURY
3.1 Introduction
Pulmonary diseases are one of the leading causes of death worldwide (18.7% of 50.5
million deaths registered in 1990, according to WHO statistics). Terminal lung failure can
be a consequence of infectious disease, cancer, and genetic disorders, or interstitial lung
disease that starts with inflammation and ends up as pulmonary fibrosis [70]. Allogeneic
lung transplantation remains the only option for patients with terminal pulmonary failure,
despite a scarcity of donated organs. However, lung transplants have relatively low
survival rates, in comparison to transplantation of other parenchymal organs [71].
Therefore there is an urgent need for novel therapies that will limit progression of
epithelial damage to end stage lung disease.
At the root of many lung fibrotic diseases is the disruption of alveolar homeostasis and
the loss of alveolar epithelial type I (AECI) or alveolar epithelial type II cells (AECII)
coupled with the inability of type II progenitor cells to resist and repair epithelial injury.
The alveolar epithelium has a relatively finite repertoire of responses to injury, which are
dictated by the alveolar milieu, a repository of cytokines and growth factors that affect
recruitment of other cells to the site of injury, or the proliferation of resident cells at the
site of injury. Alveolar lung injury resultant in the loss of homeostasis is often
accompanied by inflammation, abnormal repair, endoplasmic reticulum stress, inability
45
of progenitor cells to repopulate injured epithelia, epithelial to mesenchymal transition
(EMT), activation of resident lung fibroblasts or recruitment of circulating fibrocytes
[72]. Alveolar injury can ideally either resolve to heal the gas diffusion surface, or, as is
the case in disease, chronic wounding of the alveolus can lead to fibrotic scarring and
alveolar destruction, which eventually contributes to end stage lung disease.
The identification and characterization of the cytokines, growth factors, and other
biomarkers that dictate the response to disease is key to understanding, diagnosing,
treating and determining the trajectory of various lung disorders. Identification of the
soluble factors that promote epithelial versus fibrotic healing offers the translational
potential of correcting the alveolar milieu to minimize or ameliorate fibrosis, with the
goal of prevention of progression to end-stage disease in currently intractable diseases
such as Idiopathic Pulmonary Fibrosis (IPF), Bronchopulmonary Dysplasia (BPD) and
Chronic Obstructive Pulmonary Disease (COPD). In addition, biomarkers of normal
versus aberrant epithelial healing could be used to monitor and/or predict disease
prognosis.
3.2 Response of the alveolar epithelium to injury/disease
The alveolar epithelium has a relatively limited repertoire of responses to injury that
either lead to resolution or to chronic wounding of the alveolus, which eventually
contributes to end stage lung disease (Figure 3.1). With the hallmarks of epithelial injury
being apoptosis and necrosis, other responses such as epithelial to mesencyhmal
46
transition and chronic inflammation comprise the most typical responses of alveolar
epithelium to injury or disease [73]. In addition to AECI and AECII participating in
maintenance of alveolar homeostasis, fibroblasts and macrophages are garnering much
attention as their responses in wound healing and repair have more recently been
elucidated. All four of these cell types participate in distinct stages within wound
healing. Injury to AECI and AECII triggers the release of inflammatory mediators,
which trigger the activation and migration of inflammatory cells such as macrophages;
resident fibroblasts or circulating fibrocytes are then activated to proliferate and/or
migrate to wound sites, followed by tissue remodeling which can result in wound healing
or fibrotic scarring [74]. Corrective therapy of the alveolar milieu may therefore prove to
be beneficial in many presently serious and incurable lung diseases including BPD,
COPD and IPF that likely begin and progress with injury to the alveolar epithelium. Our
recently published [75] data show that the alveolar milieu can indeed both maintain
alveolar epithelial homeostasis and mediate alveolar repair. Importantly, the alveolar
milieu also highly chemoattractive to exogenous progenitor cells, such as AFSC, and can
induce infiltrating stem cells to augment endogenous healing, most probably through
specific combinatorial cytokine mediator release, and can also induce expression of
AECII lineage markers in previously uncommitted stem cells.
47
Figure 3.1 Response of the alveolar epithelium to injury/disease.
3.3 The distal alveolar epithelial milieu is the key to alveolar epithelial homeostasis
The alveolar milieu contains autocrine and exocrine factors that maintain the epithelium
in health, and dictate the nature of repair in disease. The distal alveolar epithelial milieu
is critical for maintaining a functional population of alveolar epithelial cells containing a
balance of autocrine and exocrine factors that maintain homeostasis. Some of the
cytokines and signaling molecules involved in this response are listed in Table 3.1. A
functional population of AECII, the putative alveolar epithelial progenitors, is necessary
to maintain the distal epithelium for efficient gas exchange, as well as providing lung
compliance, elastance and innate immunity. Biomarkers of AECII injury are seen in the
plasma of patients with acute exacerbations of IPF [76], suggesting a systemic response
to AECII damage. This balance of soluble factors released by damaged AECII, along
with local and infiltrating cells and matrix will determine whether the wound will be
!
!"#$%&'()'
*+,-)+*%'
-.!(/-+!$0'
*.).()1!12'
"-3%)1!12'-0('
!"4+*00*(!)"'
!"4+*00*(!)"' %-.*!%' %-5-"-%*(!)"'
48
sealed by epithelial or mesenchymal cells. Well-studied soluble factors in the AECII
damage milieu that have been shown to reduce fibrosis in animal studies include KGF
(by supplementation), or TGF-b (by inhibition) [77].
Table 3.1 Key signaling molecules within the pro-fibrotic alveolar milieu.
Signaling Molecule Origin Target
TNFα
Leukocytes, endothelial cells,
epithelial cells, lymphocytes,
macrophages
Fibroblasts, myofibroblasts
TGF-β
Macrophages, leukocytes,
lymphocytes, epithelial cells,
endothelial cells, mesenchymal
cells
Fibroblasts, myofibroblasts,
epithelial cells, T cells,
monocytes
Reactive oxygen species
(ROS)
Macrophages, neutrophils Fibroblasts, myofibroblasts
PDGF
Epithelial cells, endothelial
cells, macrophages, fibroblasts
Fibroblasts
MIP-1α
Neutrophils, smooth muscle
cells
Neutrophils, activated
macrophages, basophils,
eosinophils, T cells, B cells
IL-1
Epithelial cells, leukocytes,
macrophages
Fibroblasts, myofibroblasts
IL-4 T cells Myofibroblasts
IL-8 Macrophages, epithelial cells
Neutrophils, endothelial cells,
macrophages
IL-10 Monocytes, lymphocytes Fibrocytes, fibroblasts
IL-13 Leukocytes, T cells
Fibroblasts, myofibroblasts, T
cells
IL-17A CD4
+
T cells Neutrophils
IL-21 T cells T cells
IL-25 Epithelial cells, T cells T cells
IL-33 Epithelial cells T cells
CXCL5, CXCL8,
CXCL12
Macrophages, epithelial cells,
fibroblasts
fibrocytes
CCL22 Macrophages, monocytes Monocytes, lymphocytes
CCL2/MCP-1, CCL3,
CCL5, CCL12, CCL19,
CCL21b
Fibroblasts, endothelial cells,
macrophages, lymphocytes,
smooth muscle cells, epithelial
cells
Monocytes, lymphocytes,
basophils, fibrocytes, fibroblasts
CCL17
Epithelial cells, endothelial
cells, macrophages,
Monocytes, lymphocytes
49
3.4 Proteomic analysis of bronchoalveolar lavage (BAL) can be used to characterize
the alveolar milieu
A dynamic proteomic database for human BAL is available on the web [78]. The
database was established to hunt for new disease markers in BAL, to examine post-
translational modifications of BAL proteins, and to compare BAL in health and disease,
thus the database expands with advances in technology. Increased resolution and
sensitivity now enable the detection of low abundance proteins, and low MW peptides
[78]. A comparison of BAL from IPF, sarcoidosis and healthy controls [79], versus from
BAL associated with inflammatory myopathies [80] and ARDS [81] showed protein
patterns in the AECII milieu specific to each disease. The ability to identify proteins in a
high-throughput manner and build “proteomic profiles” for specific diseases is a major
leap in the early and most importantly minimally invasive detection of pulmonary disease
via biomarkers.
3.5 Cell based therapies have the potential to limit progression of fibrotic lung
disease
Cell-based therapies encompass the idea that endogenous (cells residing within an organ)
or exogenous cells (isolated from outside the organ) can be manipulated to ameliorate
disease progression or repair, regenerate or replace a diseased organ. Cell-based
therapies using circulating blood cells, including those from bone marrow, have
augmented repair and prevented subsequent emphysema or fibrosis in several animal
lung damage models, including LPS, elastase, irradiation, naphthalene, and bleomycin-
50
mediated acute lung injury [23,82-85]. Stem cells have also been used to target
therapeutic agents to the site of damage, e.g. KGF delivered by BMSC and expressed via
an inducible lentivirus [86]. However, as these therapies cannot reverse scar formation or
restore fibrotic alveolar microstructure to normal, thus our emphasis is placed on devising
methods for attenuation of fibrotic disease progression. The relatively low engraftment
of the donor cells in repairing damage models suggests that their protective effect is
paracrine in nature, augmenting the alveolar epithelial membrane’s endogenous healing
milieu. Identification of factors contributing to a pro-healing, pro-epithelial milieu would
allow pre-conditioning of cells pre transplant, or direct manipulation of the alveolar
milieu, leading to novel therapeutic approaches to prevent progression to end stage lung
disease. If harnessed, the potential of cellular therapies extends beyond simply
ameliorating symptoms of lung disease, as many pharmaceuticals do, but instead halting
and preventing further injury or degradation.
3.6 Amniotic fluid stem cells augment endogenous alveolar epithelial healing
In our hands, amniotic fluid stem cells (AFSC) are particularly efficient for cell-based
therapy (CBT) [87]. The AFSC, an intermediate type of stem or progenitor cell between
embryonic stem cells (ESC) and adult stem cells resident in differentiated organs, has
been well characterized [31]. AFSC satisfy the requirements of a stem cell potentially
suitable for CBT, since they are derived from a non controversial source, are readily
available, scalable and are pluripotent and non-teratogenic. We have chosen the amniotic
fluid stem cell AFSC to test the concept of stem cell-augmented alveolar epithelial repair,
51
rather than mesenchymal stem cells (MSC), based on our preliminary unpublished
studies, which show that the AFSC are significantly less immunomodulatory than MSCs.
We have shown that AFSC track in vivo to damaged alveolar epithelium, where they can
differentiate into epithelial cells expressing markers of AECII [30]. Furthermore, AFSC
are capable of responding to various types of in vivo alveolar injury, including
naphthalene, hyperoxia and bleomycin induced fibrosis. This tracking to the site of
damaged AECII, and damage-induced differentiation can be recapitulated in vitro, where
AFSC are attracted to damaged AECII milieu per se, and expedite wound healing by
contributing healing cytokines, such as the inhibition of MIF and PAI-1, to the milieu
[75]. Our unpublished observations indicate that AFSC can also ameliorate bleomycin-
induced fibrosis in a rodent model, demonstrating their potential functionality in
preventing fibrotic alveolar epithelial wound responses. Identification of the proteins in
the damaged AECII milieu that attract and promote differentiation of AFSC into cells
expressing markers of AECII (SPC, TTF-1 and ABCA3), and the proteins secreted by
AFSC into the damage milieu that promote wound healing, would enable the design of
therapeutic strategies to promote non fibrotic re-epithelialization and the attenuation of
disease progression.
3.7 Conclusion
The alveolar milieu contains key signaling molecules during both homeostasis and
injurious states. These cytokines and regulatory molecules play a key role in either
52
pathogenesis or normal wound healing. The elucidation of the factors within the alveolar
milieu, which dictate repair processes, is an important target for therapies aimed at
diseases, which affect the alveoli. The milieu of damaged alveolar epithelium provides a
niche for inflammatory cells and innate bone marrow stem cells, as well as for
systemically delivered stem cells. A critical, but as yet undefined balance of soluble
factors will dictate not only which cells are attracted to the site of damage but also the in
situ response of the infiltrating cells. Whether these cells engraft, contribute
cytokines/growth factors, and/or differentiate to expedite appropriate epithelial repair, or
further compromise the epithelium through inflammation and/or induction of fibrosis will
be dependent on the specific contents of the damage milieu. Thus, a detailed knowledge
of the soluble protein constituents within the AECII milieu that maintain homeostasis in
health and restore it after damage would facilitate the design of therapeutic interventions
to avert inappropriate repair mechanisms after epithelial damage.
53
CHAPTER FOUR: AMNIOTIC FLUID STEM CELLS INHIBIT THE
PROGRESSION OF BLEOMYCIN INDUCED PULMONARY FIBROSIS VIA
CCL2 MODULATION IN BRONCHOALVEOLAR LAVAGE.
4.1 Introduction
IPF is a chronic, progressive and fatal lung disease, surmised to result from a myriad of
factors. The improvement of diagnostic technology and criteria, coupled with an increase
in aged populations worldwide, virtually ensures that morbidity and mortality attributed
to IPF will increase [88]. The histopathology of IPF demonstrates a characteristic
heterogeneity: areas of normal parenchyma interspersed with areas of paraseptal and
subpleural fibrosis [89]. At the cellular level IPF is characterized by alveolar epithelial
injury, the initiation of inflammatory cascades, exaggerated pro-fibrotic cytokine
expression, increased extracellular matrix deposition, and the development of fibrotic
lesions termed ‘foci’[89-92]. The only effective and definitive treatment for IPF is lung
transplantation; however this option is limited by the quality and availability of donor
lungs.
Recently, treatment strategies for IPF have focused on immunomodulation of cytokine
targets [93]. In particular, expression of the pro-fibrotic cytokine CCL2 plays a
significant role in IPF as it is secreted by type II alveolar epithelia (AECII) and its
secretion is significantly increased during inflammatory and fibrotic remodeling events;
thus CCL2 has become a biomarker for the progression of IPF [94-98]. Furthermore, in
54
experimental models of lung fibrosis, increased expression of CCL2 attracts fibroblasts
and stimulates their proliferation [99,100]. Inhibition of CCL2 production, deletion of
CCR2 (the high affinity receptor for CCL2), or CCR2 antagonism inhibits the deposition
of collagen and attenuates the experimental development of fibrosis [101-105]. Thus
emerges the importance of CCL2/CCR2 signaling in the pathogenesis of pulmonary
fibrosis.
We have previously investigated the therapeutic potential of AFSC in both acute and
chronic injury models. Our studies have demonstrated the immunomodulatory functions
of AFSC in the kidney, which correlate with improvements in organ function, inhibition
of the development of interstitial fibrosis and increased life span in experimental animals
[64,66]. Amniotic fluid stem cells (AFSC) are a distinct population of multipotent cells
isolated from amniotic fluid based on the expression of c-kit [106]. Recent studies have
demonstrated their potential for reprogramming into a pluripotent state through simple
manipulation of culture media [107]. AFSC express additional stem cell markers found in
Embryonic Stem Cells (ESCs) such as Oct4 and SSEA-4 and can be induced to
differentiate into cell lineages of all three embryonic germ layers without forming
terratomas in vivo [31]. AFSC are negative for hematopoetic markers: CD34, CD45 and
CD133, but express CD29, CD44, CD73 and CD105, markers also found on
mesenchymal and neuronal stem cells [30]. Unlike mesenchymal stem cells (MSC),
AFSC have not been linked to development or exacerbation of fibrosis [108,109].
55
In the present study, we used the murine bleomycin injury model to induce the
parenchymal remodeling and elevated CCL2 production seen in human IPF. We then
treated cohorts intravenously with AFSC to test whether AFSC can inhibit the
progression of experimentally induced pulmonary fibrosis [110,111]. We determined that
AFSC treatment, administered during either acute or chronic fibrotic remodeling events,
inhibits changes in histology and pulmonary function associated with the development of
pulmonary fibrosis. We also observed that AFSC express CCR2, the high affinity
receptor for CCL2, home to fibrotic foci in vivo and inhibit increased CCL2 levels in
bronchoalveolar lavage (BAL) following bleomycin-induced lung injury. Through in
vitro migration assays, we discovered that AFSC migrate toward increased CCL2
concentrations found in bleomycin-injured BAL. Finally, we provide data in support of a
potential mechanism for the reduction of CCL2 by AFSC: the proteolytic cleavage of
CCL2 by secreted MMP2, inducing formation of a previously described CCR2 receptor
antagonist cleavage product [112-114].
Although the use of various cell populations to attenuate the progression of pulmonary
fibrosis, with varying degrees of success has been previously described, we are the first to
demonstrate that AFSC directly respond to increased CCL2 gradients found in injured
lung BAL. The retention of AFSC within fibrotic lesions, and their homing ability
toward CCL2 gradients suggests the potential for AFSC to deliver therapeutic effects
specifically to sites of injury, which may prove to be superior to single agent non-specific
therapies. Furthermore, we are the first to propose a potential mechanism for CCL2
56
reduction in BAL following AFSC treatment and to provide data in support of this
hypothesized mechanism [115-118]. This novel cell based therapy and proposed
mechanism suggest the translational potential for AFSC to arrest the progression of
pulmonary fibrosis at the stage at which AFSC are administered.
4.2 Methods
4.2.1 Ethics Statement: Samples of human amniotic fluid from male fetuses (12–18
week of gestation) were provided to our laboratory by Genzyme Genetics Corporation
(Monrovia, CA, USA) after karyotyping analysis. No written or verbal consent was
required since samples were not identified and information obtained about the samples
was limited to karyotype and fetal health status. All animal studies were performed in
adherence to the National Institutes of Health Guide for the Care and Use of Laboratory
Animals and approved by and performed according to the protocols and guidelines of the
Institutional Animal Care and Use Committee at Children’s Hospital Los Angeles
(Animal Welfare Assurance Number: A3276-01). All surgery was performed under
isoflurane anesthesia, and every effort was made to minimize suffering.
4.2.2 Isolation and Culture of AFSC: The isolation, culture and characterization of the
pluripotency of human and mouse AFSC is a well established protocol in our laboratory,
and clones used in these experiments are the same as those used in our previous
publications[30,32,64,66,75]. Samples of murine amniotic fluid, used in in vivo
experiments, were obtained from E13.5 embryos and samples of human amniotic fluid,
57
used in in vitro experiments, were obtained from amniocentesis samples form 12-18
weeks of gestation. Briefly, the stem cell population was isolated from the general
amniotic cellular milieu using standard Magnetic Sorting (MACS) techniques (Miltenyi
Biotech, Auburn, CA) against the cell surface marker, c-kit. Clones derived from a single
sample of amniotic fluid were cultured in petri dishes in medium containing α-MEM
Medium, 20% Fetal Bovine Serum, 1% L-Glutamine and 1% antibiotics (pen-strep)
(Gibco/ BRL, Rockville, MD) supplemented with 20% Chang Medium B and 2% Chang
Medium C (Irvine Scientific, Santa Ana, CA). All in vivo AFSC treatment experiments
were conducted with murine AFSC obtained from embryos of the same background as
the experimental animals. Prior to injection, a clonal murine AFSC population was
labeled with a cell surface marker, chloromethylbenzamine-1,1’-diactaolecyl-3,3,3’,3’-
tetramethylindocarbocyanine perchlorate (CM-Dil) (Invitrogen, Carlsbad, CA), in order
to track the cells after injection according to the manufacturers specifications.
4.2.3 Bleomycin Induced Lung Injury and AFSC Treatment: Female C57Bl/6J mice 8-12
wks. of age (Jackson Laboratories, Bar Harbor, Maine) were randomly selected, for
bleomycin-injury or saline controls; a minimum of 6 mice were used for each
experimental condition and time point. For bleomycin treatment, 1.5U/kg bleomycin
(Sigma, St. Louis, MO) was dissolved in 50µl of saline and injected into the trachea
using a sterile 28
G
needle under isoflurane anesthesia. Mice were housed in plastic cages
on a 12-hour light/12-hour dark cycle with access to food and water ad libitum until
harvest. AFSC treated mice received 1x10
6
murine AFSC intravenously (IV) in a 50µl
58
volume of sterile PBS at either 2 hours (Bleo+AFSC day 0) or 14 days (Bleo+AFSC day
14) post-bleomycin-injury. Mice receiving AFSC on day 0 were sacrificed at either 3
days (acute time point) or 28 days (chronic time point) post-bleomycin. Mice receiving
AFSC on day 14 were sacrificed at 28 days post-bleomycin.
4.2.4 Histology: Lung tissues were fixed in 4% paraformaldehyde at 20-25 cm H
2
O
inflation pressure, embedded in paraffin and cut into 5-7µm thick sections. Mouse lung
sections were stained with Sirius Red/Fast Green FCF (Sigma, St. Louis, MO) for
collagen visualization. Morphological changes in 225 randomly chosen microscopic
fields, spanning all experimental conditions at the chronic time point, photographed with
20-fold magnification, were quantified according to the numerical scale proposed by
Ashcroft et al [119]. Visualization and quantification of CM-Dil labeled cells was
achieved through counterstaining with 4’,6-diamidino-2-phenylindole (DAPI) (Vector
Laboratories, Burlingame, CA). In vitro adherent cells were fixed in 4%
paraformaldehyde for staining. Overnight incubation with primary antibodies used for
immunofluorescence included CCR2 [1µg/ml] (Abcam, Cambridge, MA) and α-smooth
muscle actin (α-SMA) [4µg/ml] (Sigma, St. Louis, MO).
4.2.5 Measurements of Lung Mechanics and Collagen Quantification: Mice were
anesthetized with 70–90 mg/kg pentobarbital sodium solution, tracheotomized, placed in
a plethysmograph and connected to the Scireq small animal ventilator (Scireq, Montreal,
Canada). Mice were mechanically ventilated at a rate of 150 breaths/min, tidal volume of
59
10 ml/kg, and positive end-expiratory pressure of 2–3 cmH2O. All maneuvers were
computer controlled via Flexivent v5.2 software (Scireq, Montreal, Canada). Pressure-
volume loops were generated by a sequential delivery of seven increments of air into the
lungs from resting pressure to total lung capacity followed by seven expiratory steps
during which air was incrementally released. Pressures at each of the incremental
volumes delivered were recorded and graphed to give pressure-volume loops. The
Salazar-Knowles equation was applied to measurements resulting from the pressure
volume manipulations to calculate quasi-static compliance and hysteresis (the area
enclosed by the pressure volume loop), which provides an estimate of the amount of
airspace closure that existed before the P-V loop maneuver [120]. Negative pressure
forced expirations were preformed via rapidly switching the airway opening to negative
pressure, resulting in the ability to measure Forced Vital Capacity (FVC). All
measurements and maneuvers were preformed in triplicate. Total collagen content of
whole lung samples was assessed by hydroxyproline assay kit (BioVision, Milpitas, CA)
according to the manufacturers instructions and assessed by spectrophotometry at 570nm.
4.2.6 Collection of Bronchoalveolar Lavage (BAL) and Lung Tissue: BAL and lung
tissue were collected according to our previously published protocol [13]. Protein
concentrations in BAL and tissue supernatants were determined using a Bradford assay
kit (Bio-Rad Laboratories, Hercules, CA). BAL supernatants were frozen at -20°C; lung
homogenates were frozen at -80°C.
60
4.2.7 Proteomic Cytokine Analysis: Cytokine levels were assessed using multiplex
cytokine assay membranes (R&D Systems, Minneapolis, MN). Membranes were
developed with Super Signal West Pico Chemiluminescent Substrate (Thermo Fisher
Scientific Inc., Rockford, IL), scanned and analyzed using Image J Software.
4.2.8 CCL2 ELISA: BAL was analyzed for CCL2 concentration using the mouse MCP-1
ELISA KIT (Invitrogen, Camarillo, CA) according to the manufacturer’s suggested
protocol with spectrophotometry at 450nm.
4.2.9 In Vitro Collagen Assay: To study the ability of the damaged cytokine milieu
secreted into BAL, specifically CCL2, to induce collagen synthesis in fibroblasts we
developed an in vitro assay system to measure newly synthesized soluble collagen.
Murine 3T3 fibroblasts (ATCC, Manassas, VA) were plated at 4x10
4
cells per well in 24
well plates and cultured in DMEM + 10% FBS + 1% penicillin-streptomycin. Once
confluent, duplicates of 100µl of BAL (n=6 per time point), conditioned media (n=10) or
recombinant murine CCL2 at 0, 50, 100 or 200 pg/ml (R&D Systems, Minneapolis, MN),
was added to each culture for an additional 48 hours. At the end of the 48-hour growth
period, conditioned media supernatants were removed, adherent cells were washed in
PBS, and incubated with 0.5M acetic acid for 2 hours. Acetic acid fractions containing
newly synthesized solubilized pro-collagen was then collected and assayed according to
the manufacturers instructions via the Sircol Assay system (Biocolor Life Sciences,
County Antrim, UK) and measured with spectrophotometry at 570nm.
61
4.2.10 Western blotting and Zymography: BAL samples were concentrated using Amicon
Ultra-4 Centrifugal Filter Units (3kDa) (Milipore, Bilerica, MA). 20µg total protein from
BAL or cell lysate was loaded onto 4-12% Bis-Tris Gels with MOPS buffer (Novex,
Grand Island, NY). Anti-CCL2 [1:200] (R&D Systems, Minneapolis, MN) was
incubated with membranes overnight. BAL zymography was performed using either 10
or 20 µg protein (indicated in figure legend) on 10% gelatin gels (Novex, Grand Island,
NY) with MMP-9 control (Amersham Life Science, Pittsburg, PA) and MMP-2 control
collected from conditioned media from CCL-201 human lung fibroblasts (ATCC,
Manassas, VA). Western blots and zymograms were each repeated a minimum of 3
times per sample.
4.2.11 Migration Assay: Murine AFSC were assayed for migration toward recombinant
mouse CCL2 (R&D Systems, Minneapolis, MN) at concentrations of 0, 25, 50, 75 and
100 pg/ml. Human AFSC were assayed for migration toward recombinant CCL2 at
concentrations of 0, 12.5, 25 and 50 ng/ml. Murine AFSC migration toward duplicates of
100µl BAL samples (n=6 per time point, CCL2 concentrations having been previously
determined via ELISA) from mice with acute and chronic bleomycin injury were also
assayed. Furthermore, to determine the extent of acutely produced CCL2 on murine
AFSC migration, a mouse CCL2 neutralization antibody (R&D Systems, Minneapolis,
MN) was used on acute BAL samples in additional wells. Migration assays were
preformed according to the Boyden chamber method as previously described [75]. Media
62
containing recombinant CCL2 or BAL was added to the well underneath the insert. As
positive and negative controls DMEM and 0.1% BSA (random migration) or DMEM
with 2.5% FBS (stimulated migration) were used in additional wells. AFSC were
allowed to migrate for 24 hours. Membranes were then stained with crystal violet, rinsed
in distilled water, eluted from the membrane using 0.1M HCl and read at 600nm in a
spectrophotometer.
4.2.12 In Vitro AFSC Rescue: To determine the extent of the hypothesized direct
interaction between AFSC and AECII during acute inflammation we devised an assay
system, which employed in vivo bleomycin lung injury, followed by in vitro murine
AFSC rescue. We isolated AECII from control or bleomycin-injured mice, three days
post-intratracheal instillation of 1.5U/kg bleomycin according to our previously published
protocol [13]. Briefly, AECII from lavaged lungs were isolated by dispase digestion
followed by differential adherence on IgG plates. Isolated AECII were plated in 6 well
tissue culture plates (BD Falcon, Franklin Lakes, NJ) coated with fibronectin (Sigma-
Aldrich, St. Louis, MO) at a density of 5x10
5
cells/well. Cells were allowed to attach
overnight. Once attached, 5x10
4
murine AFSC were added to experimental wells and
allowed to remain in culture with AECII for and additional 24 hours. At the end of the
24 hour period CCL2 levels in conditioned media was determined via ELISA (Invitrogen,
Camarillo, CA) then assayed for its ability to induce collagen synthesis according to our
previously described in vitro soluble collagen assay.
63
4.2.13 In Vitro MMP2 inhibition: AECII from non-injured lavaged lungs were isolated by
dispase digestion followed by differential adherence on IgG plates as previously
described [13]. Isolated AECII were plated in 6 well tissue culture plates (BD Falcon,
Franklin Lakes, NJ) coated with fibronectin (Sigma-Aldrich, St. Louis, MO) at a density
of 5x10
5
cells/well. Cells were allowed to attach overnight, before being injured with
100-mU/ml bleomycin. Two hours post-bleomycin injury 5x10
4
murine AFSC with and
without an MMP-2 specific inhibitor (MMPi), cis-9-Octadecenoyl-N-hydroxylamide,
Oleoyl-N-hydroxylamide, [10mM] in ethanol (EMB Biosciences, San Diego, CA) were
added to experimental wells and allowed to remain in culture with AECII for and
additional 24 hours. At the end of the 24-hour period conditioned media was collected
and CCL2 levels in conditioned media was determined via ELISA (Invitrogen,
Camarillo, CA).
4.2.14 Data Presentation and Statistical Analysis: Data are expressed as mean ± SEM
unless otherwise stated. Comparisons between two groups were determined using a two-
tailed Student’s t-test. For multiple comparisons, one-factor analysis of variance was
used, followed by the appropriate ad-hoc test as dictated by the normality and distribution
of the data. All statistical analyses were performed using SigmaPlot 12 (Systat Software
Inc., San Jose, CA). P values less than or equal to 0.05 were considered significant and
expressed as *p< 0.05; **p<0.001.
64
4.3 Results
4.3.1 AFSC treatment inhibits fibrotic parenchymal destruction 28 days post-bleomycin
injury in vivo. Histological specimens receiving AFSC treatment IV during the acute
(day 0) or chronic (day 14) phases of parenchymal destruction associated with bleomycin
injury were stained with Sirius Red/FCF Green. Analyses showed no fibrotic lesions or
alveolar destruction in control animals, while animals injured with bleomycin exhibited a
large amount of alveolar destruction and collagen deposition (red fibers) (Figure 4.1, A-
B). Animals treated with AFSC at day 0 showed minimal fibrotic changes, limited to
minor alveolar septal thickening and marginal alveolar destruction. Mice treated with
AFSC at day 14, the period that coincides with the initiation of fibrotic remodeling,
showed some collagen deposition, alveolar destruction and cellular infiltrate, which
occurred mostly in distal subpleural regions of the lung. The Ashcroft score for
histological sections from bleomycin-injured lung measured a median of 4 [119]. In
contrast, development of fibrosis in mice that received AFSC either at day 0 or day 14
was significantly diminished, generating median Ashcroft scores of 1 and 2 respectively
(p<0.05) (Figure 4.1, C). Furthermore, bleomycin-injured mice demonstrated a
significant increase in measurable hydroxyproline content when compared to controls
(p<0.001), but mice treated with AFSC showed a significant reduction in hydroxyproline
content when compared to bleomycin-injured cohorts, whether AFSC were administered
at day 0 (p<0.05) or at day 14 (p<0.05) (Figure 4.1, D). Sham injured control animals
injected with AFSC at either day 0 or day 14 did not develop fibrotic lesions or display
changes in hydroxyproline content (data not shown). These data demonstrate that AFSC
65
transplantation during either the initiating inflammatory events (2 hours post bleomycin
injury) or the inception of fibrotic remodeling (14 days post bleomycin injury)
significantly inhibits the progression of bleomycin-induced fibrosis past the extent at the
time at which the transplant occurs, preventing the progression of further fibrotic
remodeling.
66
Figure 4.1 IV administration of murine AFSC inhibits fibrotic alveolar and
parenchymal remodeling when injected during either acute or chronic periods
following bleomycin induced lung injury. (A) IV AFSC injection was administered
during either the acute period, 2 hours post-bleomycin injury, or during the chronic
fibrotic remodeling period, 14 days post-bleomycin injury. Lungs were studied at day 28
post-bleomycin injury to visualize the full extent of fibrotic remodeling. Control
animals: n=6, bleomycin injured: n=8, animals treated with murine AFSC two hours post
bleomycin injury: n=6, animals treated with AFSC 14 days post bleomycin injury: n=6.
(B) Histological analysis of adult mouse lung tissue embedded in paraffin, stained with
67
Sirius Red/FCF Green, for collagen visualization examined at 10X and 20X. All collagen
types-red; non-collagenous tissue-green/blue. (C) Ashcroft scoring of histological
sections from bleomycin-injured mice. Distributions are presented as box plots with lines
at the lower quartile, median and upper quartile, whiskers are representative of the
minimum and maximums excluding outliers, dots are representative of outliers. (D) The
measurement of total collagen content, as quantified by the hydroxyproline assay, was
used to determine the amount of collagen present within the total lungs of the
experimental cohorts. Distributions are presented as dot plots with lines indicating
median values.
4.3.2 AFSC treatment inhibits loss of pulmonary function associated with the
development of pulmonary fibrosis 28 days post-bleomycin injury in vivo. Pressure-
volume (PV) loops describe the mechanical behavior of the lungs and chest wall during
inflation and deflation. A shift of the PV-loop downwards along the volume axis occurs
due to the development of fibrotic disease, indicating that more pressure is required to
inflate the lungs to a given volume [120]. Following bleomycin injury, we indeed
recorded a significant downward shift of the PV-loop along the volume axis as compared
to control animals. Animals given AFSC at day 0 post-bleomycin injury displayed a P-V
loop at day 28 nearly identical to control animals. Mice treated with AFSC at day 14
post-bleomycin injury showed an upward shift of the PV-loop along the volume axis as
compared to control and bleomycin-injured animals. This upward shift indicates that less
pressure was required to inflate the lungs to a given volume and could be attributable to
the enlarged air-space size observed in day 14 treated mice in Figure 4.1, B. (Figure 4.2,
A). We applied the Salazar-Knowles equation to the PV-loop data to quantify hysteresis
(the area contained within the pressure-volume loop) (Figure 4.2, B) [120]. When
compared to control animals, bleomycin-injured mice showed a decrease in hysteresis
(p<0.05). Animals that received AFSC treatment at either day 0 or day 14 showed an
68
increase in hysteresis (p<0.05) when compared to bleomycin-injured mice. Following
bleomycin injury, forced vital capacity routinely decreased (p<0.05), but was improved in
cohorts treated with AFSC at day 0 (p<0.05) and day 14 (p<0.001) (Figure 4.2, C).
Quasi-static compliance, which measures the elastic recoil pressure of the lungs at a
given volume, decreased following bleomycin injury (p<0.05), but improved in both day
0 and day 14 (p<0.05) AFSC treated cohorts (Figure 4.2, D). Taken together, these
results demonstrate that following AFSC transplant, at either of the two key events in
bleomycin induced lung fibrosis, further loss of pulmonary function is impeded
supporting our hypothesis that AFSC inhibit the progression of parenchymal remodeling
associated with the development of fibrosis.
69
Figure 4.2 IV administration of AFSC attenuates loss of pulmonary function when
injected during either acute or chronic periods following bleomycin induced lung
injury. (A) Pressure-volume loops, describing the mechanical behavior of the lungs and
chest wall during inflation and deflation (B) Area of hysteresis as calculated via the
Salazar-Knowles equation. (C) Forced vital capacity. (D) Quasi-static compliance.
Control animals: n=5, bleomycin injured: n=9, animals treated with murine AFSC two
hours post bleomycin injury: n=6, animals treated with murine AFSC 14 days post
bleomycin injury: n=6.
70
4.3.3 AFSC modulate acute inflammatory cytokine expression in BAL and lung tissue in
vivo. To test our hypothesis that AFSC exert immunomodulatory effects in response to
bleomycin injury, we used proteomic arrays to examine BAL and lung tissue cytokine
profiles 3 days post-bleomycin injury. Control, bleomycin-injured and bleomycin-
injured mice that received AFSC treatment at day 0 were compared. BAL cytokine
profiles demonstrated significant changes in C5α (p<0.05), CCL2 (p<0.001) and TIMP-1
(p<0.05) levels following bleomycin injury and AFSC treatment (Figure 4.3, A).
Analysis of whole lung tissue homogenates (Figure 4.3, B) showed increases in CCL1,
CXCL1 and CCL5 (p<0.05) and a decrease in CXCL9 (p<0.05) following both
bleomycin injury and bleomycin injury with day 0 AFSC treatment.
Other cytokine modulations in BAL and lung tissue were detected, but were found not to
be statistically significant (Appendix 1). Furthermore, the cellular component of BAL
fluid, which was comprised mainly of macrophages, lymphocytes and neutrophils was
analyzed to determine the effect of cytokine modulation on inflammatory cell populations
(Appendix 2).
CCL2 concentrations in BAL from animals 3 days post-bleomycin injury were further
quantified via ELISA. BAL collected from control mice exhibited CCL2 levels of
42.67±4.01 pg/ml, while CCL2 levels in BAL from bleomycin-injured animals increased
2-fold to 84.97±13.87 pg/ml (p<0.05). Animals that received AFSC at day 0
71
demonstrated a decrease in CCL2 levels to 42.78±5.10 pg/ml (p<0.05) (Figure 4.3, C).
BAL collected from bleomycin-injured animals 28 days post-injury demonstrated an
increase in CCL2 when compared to control animals (80.89±13.07 pg/ml versus
53.15±4.45 pg/ml), in contrast with animals that received AFSC at either day 0 or day 14,
which showed significantly decreased levels of CCL2 at the 28 day post-injury time point
(20.18±6.23 (p<0.001) and 32.48±5.49 pg/ml (p<0.05), respectively) (Figure 4.3, D).
To determine the impact increased CCL2 concentrations found in BAL post-bleomycin
injury could have on collagen synthesis by fibroblasts, 3T3 fibroblasts were exposed to
increasing concentrations of recombinant CCL2 in culture. This resulted in a 2.5-fold
increase in collagen synthesis when cells were exposed to 100 pg/ml CCL2, similar to
CCL2 levels detected in bleomycin-injured murine BAL (Figure 4.3, E). BAL samples
analyzed via ELISA in Figure 4.3, C-D were then used in this same assay. BAL from
mice 3 days post-bleomycin injury elicited a moderate but noticeable increase in collagen
synthesis by 3T3 cells as compared to control BAL. Decreased levels of collagen were
synthesized by 3T3 cells exposed to BAL from AFSC treated mice (Figure 4.3, F). BAL
from animals 28 days post-bleomycin injury induced a significant increase in collagen
synthesis when compared to control animals (p<0.05). Treatment of bleomycin-injured
mice with AFSC at days 0 or 14 post-injury resulted in production of BAL that induced
significantly less collagen synthesis when compared to BAL from bleomycin-injured
mice, with 3.34-fold (p<0.001) and 1.77-fold (p<0.05) reductions in 3T3 collagen
synthesis respectively (Figure 4.3, G).
72
These data demonstrate that AFSC are immunomodulatory in the bleomycin lung injury
model, and that AFSC have the capacity to modulate key pro-inflammatory cytokines,
most notably CCL2 following treatment. Furthermore, these data show the direct impact
of recombinant CCL2 alone or BAL containing varying levels of CCL2 (associated with
AFSC treatment), on collagen synthesis by fibroblasts in vitro. The link to CCL2
mediated processes following bleomycin injury and AFSC rescue is further supported by
the recruitment of CCL2 responsive immune cell populations into the BAL with and
without AFSC treatment.
73
Figure 4.3 IV AFSC treatment modulates the acute inflammatory cytokine milieu in
both BAL and tissue following bleomycin induced lung injury. Samples from BAL
extracts (A) and whole lung homogenates (B) were analyzed via protein array to
determine their acute inflammatory profiles. Cohorts included control (n=6), bleomycin
injured (n=6) and bleomycin injured receiving AFSC transplant 2 hours post injury (n=6)
(C) CCL2 concentration in BAL quantified by ELISA during acute inflammation, 3 days
post-bleomycin injury. (D) CCL2 concentration in BAL quantified by ELISA during the
chronic injury period, 28 days post-bleomycin injury. (E) In vitro assay to determine the
direct effect of varying concentrations of recombinant CCL2 on 3T3 fibroblast collagen
synthesis. (F) Collagen synthesis induced in 3T3 fibroblasts following exposure to in
vivo acute BAL samples. (G) Collagen synthesis induced in 3T3 fibroblasts following
exposure to in vivo chronic BAL samples.
74
4.3.4 AFSC modulate CCL2 through MMP2 mediated proteolytic cleavage. To
investigate a potential mechanism of CCL2 regulation in BAL following AFSC
treatment, we preformed Western blot analysis of CCL2 in AECII cellular fractions 3
days post-bleomycin injury (immunomodulatory time point of interest), using a gradient
gel with high resolving capacity. Since we previously demonstrated that AFSC treatment
most significantly attenuates CCL2 secreted into the BAL following bleomycin injury
(Figure 4.3, A-D), Western blots of cell fractions were not used to measure changes in
levels of CCL2 secretion, but instead as an indicator of the presence and type of CCL2.
In controls, as well as following bleomycin injury, mouse-specific CCL2 was present as a
band of the expected molecular weight, 25 kDa (Figure 4.4, A). However, following
AFSC treatment, a subtle shift downward in the CCL2 band was visualized, indicating
the cleavage of a 0.4 kDa peptide. This shortened, cleaved form of CCL2 has been
previously reported to function as a CCR2 receptor antagonist, thereby rendering the
cleaved form of CCL2 found here a putative CCR2 receptor antagonist in BAL [112-
114]. In parallel studies, gelatin zymography of BAL harvested 3 days post-bleomycin
injury plus AFSC day 0 treatment showed a significant increase in MMP2 activity
(Figure 4.4, B). This effect was transient, as elevated levels of MMP2 did not persist to
28 days and this was true whether AFSC were given at 0 or 14 days post-bleomycin
injury. (Figure 4.4, C). These data are significant, as CCL2 is a known target of MMP2
proteolytic cleavage, with the CCL2 cleavage product forming the aforementioned
putative receptor antagonist for CCR2 [112]. These data suggest a potential mechanism
75
for CCL2 modulation: the proteolytic cleavage of CCL2, forming a receptor antagonist,
which maintains its binding affinity for CCR2, yet does not induce a response upon
binding. We hypothesize that the transient increased MMP2 secretion into BAL
following AFSC transplantation could be the protease cleaving CCL2 and regulating its
activity.
Figure 4.4 AFSC modulate AECII secreted CCL2 in BAL through proteolytic
cleavage by transient MMP2 expression. (A) Representative Bis-Tris SDS-PAGE
analysis of AECII fractions from control (n=6), bleomycin-injured (n=6) and bleomycin-
injured with AFSC treatment at 2 hours post-bleomycin (n=6), harvested at day 3,
demonstrated a subtle 0.4 KDa shift of CCL2 to a putative inhibitory form. (B)
Representative gelatin zymography of 20µg BAL fluid from control, bleomycin-injured
and bleomycin-injured with AFSC treatment at 2 hours post-bleomycin, harvested at day
3 (C) Representative gelatin zymography of 10µg control, bleomycin-injured and
bleomycin-injured with AFSC treatment harvested at day three as compared to BAL
fractions from animals harvested at 28 days post-bleomycin injury (receiving AFSC at
either day 0 or day 14 post-bleomycin injury), demonstrates transient nature of MMP2
increase.
76
4.3.5 AFSC chemotactically respond to increased CCL2 gradients. Control mice injected
with CM-Dil labeled AFSC did not exhibit fibrotic changes in lung tissue when analyzed
using Sirius Red/Fast Green FCF and did not demonstrate retention of AFSC (data not
shown). Mice injured with bleomycin and treated with AFSC at either day 0 or day 14
exhibited preferential AFSC retention within fibrotic regions of the lung when examined
at day 28 (Figure 4.5, A). To rule out any contribution by AFSC to the development of
fibrotic lesions, we analyzed α-SMA expression in AFSC in situ in bleomycin-injured
and treated lung and never observed co-localization of α-SMA expression with AFSC
(data not shown).
It has been previously characterized that in both human IPF and murine bleomycin-
induced lung injury increased CCL2 expression is noted within activated epithelium in
fibrotic areas [94]. To determine if the presence of AFSC in fibrotic lesions in this study
was the result of chemotaxis toward areas of increased CCL2 expression, we analyzed
AFSC prior to injection for CCR2 receptor expression. Immunofluorescent staining for
CCR2 demonstrated that indeed murine AFSC express this receptor prior to injection
(Figure 4.5, B).
Furthermore, we assayed both human and mouse AFSC were for chemotaxis in vitro
towards increasing concentrations of recombinant mouse CCL2, which is a
chemoattractant for both human and mouse cells that express CCR2 [121]. Murine
AFSC migrated, in a dose dependent manner, toward increasing concentrations of CCL2
77
in culture (Figure 4.5, C). The greatest AFSC migration observed, toward a
concentration of 100 pg/ml (similar to what is found in murine BAL following bleomycin
injury), demonstrated a 2.24-fold increase when compared to AFSC not exposed to
CCL2. Migration of human AFSC, assayed to determine the translational potential of
this chemotactic response, demonstrated a moderate peak at 50 ng/ml, a CCL2
concentration similar to that reported in BAL of IPF patients (Figure 4.5, D) [97]. BAL
samples previously analyzed via ELISA (Figure 4.3, C-D) were also tested for the ability
to chemoattract murine AFSC. Murine AFSC were significantly more attracted to
bleomycin-injured BAL than to BAL from control and day 0 AFSC-treated mice
(p<0.05) (Figure 4.5, E) demonstrating the specificity of CCL2 as a potent
chemoattractant for AFSC. Furthermore, upon CCL2 neutralization in BAL samples 3
days post-bleomycin injury using a CCL2 neutralizing antibody (Nab), migration toward
control, bleomycin-injured, and bleomycin-injured plus AFSC day 0 treated samples all
decreased significantly (p<0.001) (Figure 4.5, E). Finally, murine AFSC migration
toward BAL samples harvested at day 28 post-bleomycin injury demonstrated a
significant increase (p<0.05) in chemotaxis when compared to controls. Chemotaxis
decreased in both day 0 and day 14 AFSC treated mouse BAL (Figure 4.5, F).
These experiments demonstrated that AFSC express CCR2, are preferentially retained
within fibrotic lesions, and are not contributing to the deposition of collagen.
Additionally AFSC can actively respond to the chemotactic gradient induced following
bleomycin injury, specifically CCL2 within that injurious gradient, which once
78
neutralized inhibits the chemotaxis of AFSC. Finally, we demonstrate the translational
potential of human AFSC to respond to CCL2 gradients at concentrations found in
human IPF patients.
79
Figure 4.5 AFSC are retained within fibrotic lesions and migrate toward increased
CCL2 concentrations. (A) Sections from lungs injured with bleomycin and injected
with CM-Dil labeled AFSC 14 days post bleomycin injury, stained with Sirius Red/FCF
80
Green and DAPI and visualized at 10x show increased retention of AFSC within fibrotic
lesions (arrows). (B) Cultured murine AFSC express CCR2, the cognate receptor for
CCL2, visualized by immunofluorescence, prior to injection. (C) Murine AFSC migrate
toward a recombinant CCL2 gradient. (D) Migration elicited by CCL2 in human AFSC
toward recombinant CCL2. (E) AFSC migration toward BAL harvested at day 3 from
control, bleomycin-injured versus bleomycin-injured with AFSC treatment at day 0,
assayed for the ability to chemoattract AFSC (gray boxes). Migration toward BAL with
CCL2 neutralized using a neutralizing antibody (Nab) elicited a diminished migratory
response in AFSC (hashed boxes). Distributions are presented as box plots with lines at
the lower quartile, median and upper quartile, whiskers are representative of the
minimum and maximums excluding outliers, dots are representative of outliers. (F)
AFSC migration toward BAL samples harvested at the 28-day time point having either
received no treatment, or treatment at days 0 or 14. Distributions are presented as box
plots with lines at the lower quartile, median and upper quartile, whiskers are
representative of the minimum and maximums excluding outliers, dots are representative
of outliers.
4.3.6 AFSC co-cultured with bleomycin injured AECII inhibit increased CCL2
expression in vitro. To examine the direct interaction between bleomycin-injured AECII,
which secrete CCL2 [94], and AFSC during the acute inflammatory period, we injured
mice with bleomycin or saline, harvested AECII 3 days post-injection, and then co-
cultured AECII with murine AFSC. AECII harvested from saline injected animals grew
in circular colonies on fibronectin-coated plates, while AECII from bleomycin injected
animals grew sporadically and did not appear to attach well (Figure 4.6, A). Addition of
AFSC to bleomycin-injured AECII in culture resulted in the AFSC surrounding
bleomycin-injured AECII, which then formed colonies (arrows). Further visualization
with CM-Dil labeled AFSC and unlabeled AECII demonstrated a similar phenomenon, in
which AFSC surround bleomycin injured AECII colonies (Figure 4.6, B).
81
In experiments on cultured AECII that paralleled our previous observations using BAL,
elevated levels of secreted CCL2 in conditioned media as measured by ELISA were
observed in bleomycin-injured AECII wells at 70.47±6.83 pg/ml compared to 35.85±1.68
pg/ml measured in control AECII wells (p<0.05). Conditioned media from wells
containing AECII co-cultured with AFSC demonstrated a decrease in secreted CCL2
levels to 44.91±7.98 pg/ml (p<0.05) (Figure 4.6, C). Conditioned media from cultured
cells isolated from bleomycin-injured lung induced a significant increase in collagen
synthesis in 3T3 fibroblasts (p<0.05) as compared to control AECII conditioned media.
This ability to stimulate collagen synthesis was reduced when injured AECII were co-
cultured with AFSC (Figure 4.6, D). These experiments demonstrate that direct
interaction between AFSC and injured AECII results in CCL2 modulation identical to
that seen in our in vivo modeling.
82
Figure 4.6 In vitro AFSC co-culture with in vivo injured AECII recapitulates in vivo
CCL2 regulation. (A) Bright field microscopy of in vivo sham and bleomycin-injured
AECII at 3 days post-injury cultured with and without AFSC. Scale bar =200µm. (B)
Phase contrast microscopy of in vivo injured AECII co-cultured with CM-Dil stained
(red) AFSC visualized via fluorescence microscopy. Scale bar =100µm. (C) In vitro
levels of CCL2 in conditioned media as measured by ELISA. (D) Effect of conditioned
media from in vitro AFSC co-culture experiments on induction of 3T3 collagen
synthesis.
4.3.7 Inhibition of MMP2 in vitro attenuates the ability of AFSC to reduce CCL2
expression. Finally, to test our hypothesis that MMP2 plays a role in CCL2 regulation,
we sought to determine if inhibition of MMP2 using an MMP2 specific inhibitor (Oleoyl-
83
N-hydroxylamide) would restore increased secreted CCL2 levels observed in cultured
AECII following bleomycin injury, we employed an assay in which AECII were injured
with bleomycin in vitro and co-cultured with AFSC two hours post-injury. Control levels
of CCL2 in AFSC and non-injured AECII conditioned media were measured at
27.18±1.78 pg/ml and 38.18±1.75 pg/ml, respectively (Figure 4.7, A). After AECII
bleomycin injury in vitro, CCL2 levels in the media doubled to 72.13±2.68 pg/ml
(p<0.05). Following co-culture with murine AFSC, CCL2 levels significantly decreased
to 47.86±1.15 pg/ml. Finally, addition of the MMP2 inhibitor to bleomycin-injured
AECII co-cultured with AFSC resulted in a significant increase in CCL2 to 62.22±1.42
pg/ml (p<0.05). Furthermore, this increase in CCL2 was not significantly different from
levels in wells that had experienced bleomycin injury alone. These data demonstrate that
CCL2 reduction following AFSC co-culture, is linked to increased MMP2 expression by
AFSC, and that inhibition of MMP2 results in increased CCL2 levels that are statistically
indistinguishable from levels expressed by AECII injured with bleomycin.
Based on the data in its entirety, we propose the following mechanism (Figure 4.7, B).
Following AECII injury, CCL2 is secreted [94]. CCL2 triggers fibroblast migration,
proliferation and survival. AFSC express CCR2, the receptor for CCL2 and respond
chemotactically to increased CCL2 gradients. MMP2 is secreted during migration, a
known mechanism utilized by highly motile cells [122]. Secreted MMP2 cleaves CCL2
forming a receptor antagonist [114]. Binding of the CCR2 antagonist downregulates the
further secretion of CCL2, thus inhibiting the further development of CCL2 induced
84
fibrosis. We suggest that the proposed mechanism of CCL2 reduction by AFSC is a
temporal response. We surmise that AFSC home to fibrotic lesions during a period that
CCL2 is increased and that subsequent secretion of MMP2 by AFSC reduces CCL2
levels following localization within these lesions.
Figure 4.7 Inhibition of MMP2 in vitro inhibits the ability of AFSC to reduce CCL2
levels following AECII bleomycin injury. (A) CCL2 ELISA of conditioned media
from in vitro injury of AECII with 100 mU/ml of bleomycin with AFSC co-culture with
and without the addition of an MMP2 inhibitor. (B) Proposed mechanisms of AFSC
mediated CCL2 modulation within the alveolus.
4.4 Discussion
IPF is a disease that lacks both a cause and definitive treatment and is typically not
diagnosed until the chronic stage in which fibrotic lesions have been established and
patients present with diminished lung function [123]. The need for treatment strategies
for IPF during clinically relevant diagnostic periods; targeted toward clinically relevant
pro-fibrotic mediators is essential. The data from our in vivo and in vitro models of lung
fibrosis is clinically and physiologically relevant to CCL2 dependent events characteristic
of human IPF.
85
We have demonstrated that AFSC treatment inhibits changes in lung function associated
with the development of bleomycin-induced fibrosis when administered during both
acute and chronic fibrotic remodeling events. While the acute intervention (day 0)
allowed us to investigate a novel mechanism of action of AFSC, the chronic intervention
(day 14) provided data that are clinically relevant. Although lung function in day 14
treated cohorts could not fully be restored to normal levels, due to alveolar destruction
caused by the development of fibrosis prior to treatment, we demonstrated that following
AFSC treatment lung function and destruction of the alveolar architecture did not
progress to the fibrotic extent seen in untreated cohorts. In all cohorts, the inhibition of
the development of fibrosis was demonstrated by a decrease in measured hydroxyproline
content, measured Ashcroft score and the preservation of lung mechanics and pulmonary
function.
Based upon clinical and experimental characterizations of cellular and molecular
responses in human IPF, specifically exaggerated CCL2 expression [96,97,124,125], we
have presented data, which indicate that AFSC attenuate increased CCL2 in BAL in an
experimental model of lung fibrosis. We demonstrated that transient and local MMP2 up
regulation, by AFSC in BAL was associated with the proteolytic cleavage of CCL2, thus
creating a localized CCL2/CCR2 antagonist within the injured alveolar milieu. We
hypothesize that this antagonist downregulates further pro-fibrotic CCL2/CCR2
signaling. Our data indicate that CCL2 regulation by MMP2-mediated proteolytic
86
cleavage occurs acutely following AFSC treatment, but continues chronically in vivo to
downregulate CCL2 in BAL. We surmise that this transient MMP2 expression is
sufficient to cleave excess CCL2 produced during the active disease state, yet transitory
enough to avoid the parenchymal degradation typically associated with chronic up
regulation of MMPs [126]. We supported this proposed mechanism through analyses of
two, independent, in vitro AECII injury models in which significant, secreted CCL2
expression was attenuated following AFSC co-culture. Finally, it is important to note that
in all experiments that utilized AFSC, CCL2 secretion was not completely abrogated,
which may be critical for the protection of the homeostatic arm of the CCL2/CCR2
signaling pathway [121]. Another potential impact of these data lies in the ability of
AFSC to not only target their salutary therapeutic properties during clinically relevant
intervention periods, but to home to the diseased region of the lung, foregoing non-
diseased regions, as seen in our in vitro migration assays and in vivo histology that shows
AFSC chemotaxis toward CCL2 and retention within fibrotic regions.
AFSC therapy, unlike previously published MSC based therapy, has yet to show
deleterious secondary effects such as tumorogenesis or expression of fibrotic phenotypes
in experimental models of chronic fibrotic injury [30,64,66,127]. Furthermore, unlike
specific CCL2 inhibitors, these studies coupled with our previously published findings
demonstrate the plasticity of the mechanisms of action of AFSC, which are dependent on
the type and location of injury. We suggest that this makes AFSC perhaps more
translationally applicable in the context of disease than many single agent systemic drug
87
therapies. Our findings not only demonstrate the efficacy of AFSC within the bleomycin
injury model, we provide data which suggest a novel mechanistic role for AFSC
regulation of CCL2 resulting in the inhibition of parenchymal remodeling and the
development of pulmonary fibrosis. These data provide insight into the potential
tractability of targeting the CCL2/CCR2 pathway in fibrotic lung diseases via a novel
AFSC cell-based therapy, and provides a treatment strategy that we think deserves further
evaluation.
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CHAPTER FIVE: REPOPULATION OF DECELLULARIZED LUNG MATRIX
USING AMNIOTIC FLUID STEM CELLS
5.1 Introduction
The American Lung Association estimates that there are currently more than 35 million
Americans living with chronic lung disease; 400,000 of which will die within the year.
In cases where lung disease or injury has progressed to the point where transplantation is
the only treatment option, clinicians and researchers are investigating the possibility of
whole organ replacement via tissue-engineered lung. Tissue engineered lung can be
derived from either endogenous or exogenous stem or progenitor cell populations seeded
on allogeneic or xenogeneic lung matrix. Seeded matrices would then, in theory, be
cultured to repopulate the organ with functional organ specific tissue. Such a strategy
reflects a novel approach to regenerative medicine which could result in a decrease in
transplant waiting time limited only by organ regeneration time, improved patient
survival rates and a reduction in transplant rejection as exact donor matches would be
unnecessary. As the American Lung Association estimates that lung disease is the third
leading cause of death in the United States, responsible for one in six deaths, the potential
impact, scope and scalability of this translational treatment strategy is enormous.
The successful creation of tissue-engineered lung involves a two-fold approach: selection
of an appropriate candidate stem cell population and the use of appropriate scaffolding
for repopulation. Thus, we first sought to investigate the potential of Amniotic Fluid
89
Stem Cells (AFSC), to generate lung tissue via expression of lung specific lineage
markers. The use of these cells is a novel yet logical hypothesis, as in utero, the lungs of
the fetus are filled with amniotic fluid, which contributes to their development,
maturation, growth and maintenance. Our published studies have found that AFSC can
incorporate into developing mouse embryonic lung and express human lung epithelial
cell markers not previously expressed, demonstrating their regenerative and
differentiation potential [30]. Additionally, following injury in the lung it was observed
that AFSC appear to be retained at sites of injury, and exhibit alveolar epithelial type II
(AECII) phenotypes following transplantation suggesting that once in the lung, these
cells are stimulated to differentiate into alveolar epithelial phenotypes in response to
injury.
The second facet of creating a tissue engineered lung is to seed AFSC on an
architecturally, physiologically and functionally relevant scaffold. Thus, we investigated
the use of decellularized lung matrix as the biological scaffolding for cells to “rebuild”
tissue. Decellularized matrix provides an advantage over synthetically engineered matrix
as research in the field has recently demonstrated the importance of the mesenchyme in
directing cell morphology, differentiation, repair and development [128,129].
Additionally, decellularized matrix can be obtained from either human or animal donors,
such as the pig. The ability to obtain these scaffolds from a variety of donor species
makes this strategy limited only by organ regeneration time.
90
Our characterization of human AFSC elucidated the expression of type I and type II lung
epithelial cell markers prior to culture on decellularized matrix, which had not previously
been described. Furthermore, protocol devised herein, for organ decellularization has
allowed us to successfully and efficiently decellularize the lung while maintaining the
extracellular architecture intact. Finally, we have established seeding and culture
protocols that have allowed us to culture AFSC on decellularized lung matrices for up to
35 days. Furthermore, throughout culture, we characterized AFSC secretions of growth
and angiogenic factors critical to the formation of functional organoid units. These
preliminary findings strongly support our previously published claims and working
hypotheses that these unique and novel amniotic fluid derived cells have the potential to
be utilized for in vitro tissue engineering.
5.2 Methods
5.2.1 Isolation and characterization of human AFSC
Five samples of human amniotic (Table 5.1) fluid from 12-18 weeks of gestation were
isolated and characterized as previously described[30,32,64,66,75]. Briefly, the stem cell
population was isolated from the general amniotic cellular milieu using standard
Magnetic Sorting (MACS) techniques (Miltenyi Biotech, Auburn, CA) against the cell
surface marker, c-kit. Pluripotential characteristics of the clonal and subclonal groups
were tested according to previously published protocols [30,32,39,64,66]. Clones derived
from a single sample of amniotic fluid were cultured in petri dishes in medium containing
α-MEM Medium, 20% Fetal Bovine Serum, 1% L-Glutamine and 1% antibiotics (pen-
91
strep) (Gibco/ BRL, Rockville, MD) supplemented with 20% Chang Medium B and 2%
Chang Medium C (Irvine Scientific, Santa Ana, CA).
Line Passage Weeks of Gestation
14 7 Unknown
1 8 16.6
21 9 16.6
13 19 17.4
6 7 16.4
Table 5.1 Identification of the 5 human AFSC lines tested.
5.2.2 Characterization of human AFSC via qPCR for subpopulations with lung specific
markers
Aliquots of clones derived from a single samples of amniotic fluid were tested for various
markers characteristic of lung epithelial development, type I alveolar epithelia, type II
alveolar epithelia, stem and Clara cell lineages via qPCR (Table 5.2). RNA was extracted
using the Qiagen RNeasy kit (Qiagen, Valencia, CA) according to the manufacturer’s
instructions. Quantitative PCR was performed using a Roche Light Cycler 480. Each
qPCR analysis was performed in triplicate.
92
Gene Lineage Primer Sequence
18S control
F: AAATCAGTTATGGTTCCTTTGGTC
R:GCTCTAGAATTACCACAGTTATCCAA
AQP5
Type I Alveolar Epithelial
Cell
F: CCTGCGGTGGTCATGAAT
R: TTGGGGAAGAGCAGGTAGAA
TTF-1/
Nkx2.1
Early embryonic lung
epithelium; Type II
Alveolar Epithelial Cell
F: TCATTTGTTGGCGACTGG
R: TGCTTTGGACTCATCGACAT
FoxP1
Lung epithelial
development
F: CATGCTGGAAAACAGCCTAAA
R: AAAAGCTGCTGCTGAAAAGC
FoxA2
Lung epithelial
development
F: TGCGTGCTTTATTTATGGCTTA
R: GGGATAAAACCGGGTATAACACTT
ABCA3
Type II Alveolar
Epithelial Cell
F:AGCACCTTCTTCAGCAAAGC
R: CCACGAAGAAGTAGGGGATGT
SP-A
Type II Alveolar
Epithelial Cell
F: CATCAAATCCTGCAGACAAGG
R: TGGCATCAAAAGTGATGGAC
SP-B
Type II Alveolar
Epithelial Cell
F: ATCTCTCCGAGCAGCAATTC
R: ACAGCTAGCGCACCCTTG
SP-C
Type II Alveolar
Epithelial Cell
F: CGTGGTGTATGACTACCAGCA
R: GGATGCTCTCTGGAGCTATCTT
SP-D
Type II Alveolar
Epithelial Cell
F: GACGGGATGGGAGAGAGG
R: CTCCCTTTGGTCCAGGTTCT
CC10 Clara Cell
F: CTGCAATGGAACTTTTCAGC
R: CACAGTGAGCTTTGGGCTATT
PDPN/
T1a
Type I Alveolar Epithelial
Cell
F: CTGCTCTTCGTTTTGGGAAG
R: CGCCTTCCAAACCTGTAGTCT
Sca-1/Ly6E Stem Cell Antigen
F: GCCATCCTCTCCAGAATGAA
R: GCAGGAGAAGCACATCAGC
Table 5.2 Gene, lineage and primer sequences for lung lineage markers tested.
5.2.3 Western blot analysis of lung lineage markers: Western blot analysis was performed
on freshly cultured and lysed AFSC samples. Protein concentrations were determined
using a Bradford assay kit (Bio-Rad Laboratories, Hercules, CA) to ensure equivalent
sample loading. Samples were probed with for FoxP1 (Cell Signaling, Danvers, MA
1:1000), FoxA2 (Seven Hills, Cincinnati, OH 1:5000), AQP5 (ABCAM, Cambridge, MA
1:1000), ABCA3 (Seven Hills, Cincinnati, OH 1:1000), PDPN (ABCAM, Cambridge,
MA 1:1000) and Actin (Cell Signaling, Danvers, MA 1:1000). Membranes were
developed with SuperSignal West Femto Chemiluminescent Substrate (Thermo Fisher
Scientific Inc., Rockford, IL).
5.2.4 Decellularization of lung matrix: De-identified autopsy specimens obtained from a
previous collaboration with Dr. G. Orlando at Wake Forrest University were cut into
93
300
3
µm sections and placed into DI solution: sterile filtered deionized (DI) water with 5%
pen/strep (Gibco/ BRL, Rockville, MD) and incubated for 1 h at 4°C. Specimens were
then rinsed 5 times in DI solution and incubated in deoxycholate solution: sterile filtered
2% sodium deoxycholate (Sigma-Aldrich, St. Louis, MO) with 1% pen/strep for 24 h at
4°C. Following incubation specimens were removed from deoxycholate solution and
rinsed 5 times in DI solution. Specimens were then incubated in NaCl solution: sterile
filtered 1 M NaCl (Sigma-Aldrich, St. Louis, MO) with 5% pen/strep for 1 h at room
temperature and rinsed with DI solution five times as previously described. Following
the final rinse, specimens were incubated in DNase solution: sterile filtered 30 mg/mL
bovine pancreatic DNase (Sigma-Aldrich, St. Louis, MO) in 1.3 mM MgSO4 (Sigma-
Aldrich, St. Louis, MO) and 2 mM CaCl2 (Sigma-Aldrich, St. Louis, MO) with 5%
pen/strep for 1 h at room temperature followed by 5 rinse steps with DI solution.
Incubation and rise steps with DI solution, deoxycholate solution, NaCl solution, and
DNase solution were then repeated two more times to ensure complete cellular removal.
5.2.5 Seeding and Culture of AFSC on decellularized lung: 1x10
6
AFSC (line 6) were
seeded per matrix in Corning ultra low attachment 25cm
2
culture flasks (Corning, Inc.,
Tewksbury, MA). Cells and matrices were cultured using medium containing α-MEM
Medium, 20% Fetal Bovine Serum, 1% L-Glutamine and 1% antibiotics (pen-strep)
(Gibco/ BRL, Rockville, MD) supplemented with 20% Chang Medium B and 2% Chang
Medium C (Irvine Scientific, Santa Ana, CA) as described previously.
94
5.2.6 Histology: Lung tissues were fixed in 4% paraformaldehyde, embedded in paraffin
and cut into 5-7µm thick sections. Adherent cells were fixed in 4% paraformaldehyde
and washed with phosphate buffered saline prior to staining. Lung tissue sections were
stained with Hematoxylin and Eosin and Sirius Red/Fast Green FCF (Sigma, St. Louis,
MO). Overnight incubation with primary antibodies used for immunofluorescence
included FoxP1 (Cell Signaling, Danvers, MA 1:200), FoxA2 (Seven Hills, Cincinnati,
OH 1:200), AQP5 (Abcam, Cambridge, MA 1:500), ABCA3 (Seven Hills, Cincinnati,
OH 1:250), PDPN 25 kDa (Abcam, Cambridge, MA 1:500) and Sca-1 (Fitzgerald, Acton,
MA: 1:200).
5.2.7 Analysis of secreted cytokines, growth factors and pluripotential characteristics:
Cytokine, growth factor and pluripotentiality of AFSC prior to and throughout culture on
decellularized matrices was assessed using MILLIPLEX MAP Human Stem Cell
Pluripotency Magnetic Bead Panel 2 and Human Cytokine/Chemokine Magnetic Bead
Panel (EMD Millipore, Billerica, MA) according to the manufacturers instructions and
analyzed using the luminex multiplex system (EMD Millipore, Billerica, MA).
5.3 Results
5.3.1 AFSC express Type I and Type II alveolar epithelial lineage markers prior to
culture on decellularized lung matrices. qPCR expression profiles indicate the presence
of pulmonary epithelial and stem cell lineages as demonstrated by expression of FoxP1,
FoxA2, ABCA3, AQP5, PDPN/T1α, Sca-1 (Table 5.3 and Figure 5.1). Lines 14, 6, 1 and
95
13 expressed FoxP1 mRNA at comparable levels, with increased expression in line 21.
Fox A2 was mRNA expression was low in all lines except line 13 which demonstrated
high levels of FoxA2 mRNA expression. ABCA3 mRNA expression increased as the
age of gestation at cell harvest increased. AQP5 was expressed at low levels in all lines
except line 1, which demonstrated markedly increased AQP5 mRNA. PDPN/ T1α
mRNA expression was increased in lines 6 and 21. Finally, Sca-1 mRNA expression was
expressed at comparable levels in all lines examined.
Protein analysis via western blot of all 5 hAFSC lines (Figure 5.2) demonstrated
comparable expression of FoxP1 protein in hAFSC lines 1, 6 and 21, increased
expression in hAFSC lines 13 and 14. FoxA2, AQP5, ABCA3 and PDPN/T1a protein
levels in all 5 hAFSC lines were expressed at comparable levels when normalized against
total protein concentration as demonstrated by actin staining. Finally, immunofluorescent
staining for lung lineage markers within the 5 hAFSC cell lines (Figure 5.3) demonstrated
positive staining for Sca-1, FoxA2, FoxP1, PDPN, ABCA3 and AQP5.
96
Gene
mRNA
Expressed
Lineage
18S Yes control
AQP5 Yes Type I Alveolar Epithelial Cell
TTF-1/Nkx2.1 No
Early embryonic lung epithelium; Type II
Alveolar Epithelial Cell
FoxP1 Yes Lung epithelial development
FoxA2 Yes Lung epithelial development
ABCA3 Yes Type II Alveolar Epithelial Cell
SP-A No Type II Alveolar Epithelial Cell
SP-B No Type II Alveolar Epithelial Cell
SP-C No Type II Alveolar Epithelial Cell
SP-D No Type II Alveolar Epithelial Cell
CC10 No Clara Cell
PDPN/T1a Yes Type I Alveolar Epithelial Cell
Sca-1/Ly6E Yes Stem Cell Antigen
Table 5.3 Gene, expression and lineage of lung markers tested.
97
Figure 5.1 qPCR analysis expressed as relative mRNA expression for lineage
markers in all 5 hAFSC lines. FoxP1 (A), FoxA2 (B), ABCA3 (C), AQP5 (D),
PDPN/T1α (E), Sca-1 (F).
98
Figure 5.2 Western blot analysis of lung lineage markers. (A) Western blot analysis
for FoxP1 demonstrates characteristic staining at 70 kDa and ~90 kDa (Cell Signaling
1:1000), FoxA2 48 kDa (Seven Hills 1:5000), AQP5 28 kDa (ABCAM 1:1000) (B)
ABCA3 191 kDa (Seven Hills 1:1000), PDPN 25 kDa (ABCAM 1:1000) and Actin 42
kDA (Cell Signaling 1:1000).
99
Figure 5.3. Immunofluorescent staining of lung lineage markers within all 5 hAFSC
lines. 10X magnification of cultured AFSC stained for Sca-1 (pink, denoted by white
arrowhead), FoxA2 (green), FoxP1 (green), PDPN (red), negative control, ABCA3
(green), AQP5 (green).
5.3.2 Characterization of secreted cytokines, growth factors and interleukins prior to
culture on lung matrix. Characterization of factors secreted into culture media prior to
culture on decellularized lung matrix, corrected for background, revealed increased levels
of anti-inflammatory IL-6 and pro-angiogenic IL-8 with increased AFSC gestational age
100
(Figure 5.4 A). Furthermore, increased regulator of adhesion CX3CL1 (fractalkine) and
neurogenic GRO (growth related oncogene) was observed in gestationally older AFSC
along with decreasing levels of CCL2 (monocyte chemotactic protein-1) with increased
gestational age (Figure 5.4 B). AFSC line 13 demonstrated increased TGFα
(transforming growth factor alpha), an activator for cell proliferation, differentiation and
development and increased PDGF-BB (platelet derived growth factor-BB), a modulator
of endothelial proliferation and angiogenesis, and GM-CSF (granulocyte macrophage
colony stimulating factor), when compared to the other AFSC lines. All lines
demonstrated expression of PDGF-AA (platelet derived growth factor-AA), PDGF-BB
and VEGF (vascular endothelial growth factor) at variable levels (Figure 5.4 C). Finally,
testing for pluripotency markers in lysed cells demonstrated minimal levels of Tra-1-60
and Tra-1-81 (both human pluripotent stem cell markers), SSEA-1 (stage specific
embryonic antigen-1), KLF4 (Krueppel-like factor 4), E-Cadherin and Lin-28 (regulator
of self renewal). Significant but variable levels of EpCAM were detected in all 5 AFSC
lines (Figure 5.4 D).
101
Figure 5.4 Detection of secreted interleukins, cytokines, growth factors and
intracellular pluripotency markers within AFSC prior to culture on decellularized
lung matrices. Characterization of secreted (interleukins, cytokines and growth factors)
and intracellular factors (pluripotency) in AFSC lines, AFSC line tested denoted by
number and color in legend.
5.3.3 Decellularization removes cellular material from extracellular matrix without
disrupting alveolar structure. Three cycles of detergent enzymatic decellularization
effectively removed all cellular material from lung specimens leaving behind only the
intact extracellular matrix free of cellular material as detected by hematoxylin and eosin
staining (Figure 5.5 A) and DAPI staining (Figure 5.5 B).
102
Figure 5.5 Decellularization of lung specimens produces acellular matrices. (A) 20X
magnification of decellularized lung specimen stained with hematoxylin and eosin
demonstrates complete cellular removal. (B) DAPI staining of decellularized lung
matrices demonstrates absence of nuclei. Scale bar = 25µm.
5.3.4 AFSC seeded on decellularized matrices can be cultured for up to 35 days. AFSC
seeded on matrices were cultured up to 35 days and maintained expression of T1α/PDPN
throughout the culture period (Figure 5.6, up to day 28 shown). Furthermore, AFSC were
observed to localize within large airways and alveoli. Analysis of secreted interleukins
throughout 28 day culture demonstrated a decrease in IL6 and IL8 until days 16-17 in
culture, which then increased gradually until day 28 (Figure 5.7 A). Levels of IL7
remained low, but detectable throughout culture, all other interleukin levels were
considered minimal when corrected for background. Similarly, levels of CX3CL1 were
low, but detectable throughout culture (Figure 5.7 B). GRO demonstrated a decrease
similar to IL6 and IL8 until day 16-17, followed by a gradual increase until day 28.
Finally, levels of the cytokine CCL2 remained low throughout culture until day 21, where
a gradual increase was observed until day 28. Secreted growth factors VEGF and PDGF-
BB decreased until days 16-17 and increased through day 28 (Figure 5.7 C). Other
103
secreted growth factors: EGF (epidermal growth factor), TGFα, G-CSF (granulocyte
colony stimulating factor), GM-CSF and PDGF-AA demonstrated similar decreases
throughout the first 16-17 days, which increased in the final 11 days; however levels
detected were not as notable as those of VEGF and PDGF-BB. Analysis of intracellular
pluripotency factors revealed elevated levels of the adhesion marker EpCAM throughout
culture, up to 35 days (Figure 5.7 D).
Figure 5.6 Acellular matrices provide a culture system for AFSC for up to 28 days.
Paraffin embedded sections of AFSC seeded on lung matrices stained with T1α/PDPN
(red) and counterstained with DAPI (blue). Scale bar on 3 and 14 day specimens 50µm,
28 day specimen scale bar 25µm.
104
Figure 5.7 Detection of secreted interleukins, cytokines, growth factors and
intracellular pluripotency markers within AFSC during culture on decellularized
lung matrices. Characterization of secreted (interleukins, cytokines and growth factors)
and intracellular factors (pluripotency) throughout culture. Measured factor denoted in
legend.
5.4 Conclusion
As is the case with many progressive diseases, eventual loss of organ anatomy and
function is inevitable, resulting in the need for transplant. Although some progressive
diseases can be attenuated prior to complete loss of function, in the case of idiopathic
pulmonary fibrosis (IPF), diagnosis can often be too late [130]. Thus, recent research
strategies have focused on the possibility of engineering organoid units in vitro in an
attempt to restore organ function, improve patient outcomes, and alleviate reliance on
donor organs, which are often scarce.
105
The successful engineering of organoid units will require two significant components: the
selection of appropriate cell populations to create organ specific functional tissue and
architecturally appropriate scaffolds upon which these cells are to be seeded. Our
preliminary data presented indicates that AFSC are a potential candidate cell population
for the creation of lung epithelia. We have demonstrated that ckit
+
AFSC express
markers of type I and type II in addition to stem cell specific markers. Furthermore, we
have demonstrated that through our detergent/enzymatic protocols we can efficiently
remove cellular material from lung specimens leaving behind acellular matrices that
serve as culture systems for AFSC. We have also demonstrated that AFSC can be
cultured on these matrices for up to 35 days while concurrently secreting growth and
angiogenic factors, as well as anti-inflammatory, proliferative and pluripotent factors.
The scope of this project provides several important milestones, both in regenerative
medicine, and scientific discovery. This novel strategy has the potential to revolutionize
treatment for terminal diseases and reduce dependence on pharmaceuticals and prolonged
clinical care. Furthermore, this strategy provides an interesting opportunity to
demonstrate proof of principle, and establish a model for organ reengineering where
organs can be cultured and engineered for eventual therapeutic use. We are confident
that through continued protocol refinement and investigation we can demonstrate
functionality, as well as a better characterization of cellular responses to culture on these
matrices and potentially provide the groundwork for developing the use of acellular
AFSC seeded matrices as a therapeutic strategy.
106
CHAPTER SIX: CONCLUDING REMARKS AND FUTURE DIRECTIONS
The lung is a complex organ that involves a large surface area that interfaces with the
outside environment as well as the circulatory system, making it particularly vulnerable
to injury and disease. Epithelial cell turnover in the lung is quite slow when compared to
other organs, particularly in response to injury and disease. Endogenous cell populations
can easily be overwhelmed when the disease or injury to the lung is great enough.
Furthermore, as most lung diseases are progressive, lung disease and regeneration
becomes a considerable obstacle in aging populations. To address these growing public
health concerns, novel therapies that stimulate and protect endogenous lung stem or
progenitor cells or supply exogenous cells for regenerative or immunomodulatory
purposes have become a therapeutic target. Furthermore, in cases where lung disease or
injury has progressed to the point that cellular therapy is not an option, clinicians and
researchers are investigating the possibility of whole organ replacement via tissue-
engineered lung, derived either from decellularized lung matrix, or bioengineered
scaffolding. These novel and groundbreaking approaches may have the potential to
contribute to repair processes, slow the decline in lung function, or even perhaps to
regenerate functioning lung.
Fibrotic lung injury, in particular, is often attributed to a myriad of factors including
environmental exposure, age, genetic predisposition, epigenetics, co-existing conditions,
acute lung injury and viral infection. No effective therapies, other than lung
107
transplantation, have proven effective against lung fibrosis. Loss of cellular homeostasis
mechanisms in alveolar epithelial type I cells and any inability of type II progenitor cells
to resist and repair epithelial injury are indicators that impaired response to injury and
regeneration is a critical component of this disorder. The alveolar epithelium has a
limited repertoire of responses to injury, which are dictated by the alveolar milieu, a
repository of cytokines and growth factors that affect recruitment of other cells to the site
of injury, or the proliferation of resident cells at the site of injury. The identification and
characterization of the cytokines, growth factors, and other biomarkers that dictate the
response to disease is key to understanding, diagnosing, treating and determining the
trajectory of various lung disorders. Corrective therapy of the alveolar milieu may
therefore prove to be beneficial in many presently serious and incurable lung diseases
that likely begin and progress with injury to the alveolar epithelium.
This dissertation has explored the potential for Amniotic Fluid Stem Cell based therapy
to arrest or delay the progression of fibrotic lung disease through investigation of a novel
proteolytic immunomodulatory mechanism within the alveolar milieu. We have also
demonstrated that AFSC are a viable candidate cell population for in vitro lung tissue
engineering. Critical to the further development of AFSC as a treatment strategy is the
development of bold and innovative techniques and experimental systems that more
effectively mimic the clinical etiology of the disease so that therapeutic strategies can be
rigorously tested.
108
In the context of the use of AFSC as an immunomodulatory treatment strategy to rescue
or delay the progression of pulmonary fibrosis, our data demonstrate that AFSC based
therapies provide have the potential to yield targeted, lesion specific treatments for IPF,
that indeed arrest the progression of experimental pulmonary fibrosis. Most exciting
about this prospect is that AFSC provide similar effects to single agent therapeutic
compounds while targeting to specific lesions, something that is still not possible using
systemically administered pharmaceuticals. In order for this cellular therapy to progress,
a full understanding of mechanisms of AFSC targeting is necessary in order to optimize
delivery and retention of the therapeutic properties of AFSC to specific lesions in order to
improve patient outcomes.
In the context of using AFSC as a cell population for bioengineering of lung tissue,
further investigation, including functional testing, maintenance of or differentiation into
pulmonary lineages, is critical to developing this strategy. The expression of pulmonary
epithelial lineages both prior to and after seeding on acellular matrices is exciting,
however maintenance of these lineages and proof of proper function in vitro and in vivo
is essential to understanding and improving the potential of these cells as an engineering
tool. Furthermore, as the lung is an organ composed of a myriad of cell types, perhaps
seeding of various cell types on scaffolding is key to further development of this strategy.
109
The prospect of using cell based therapies, specifically AFSC, as a treatment strategy for
pulmonary fibrosis is exciting in that this versatile cell population has demonstrated the
ability to serve as a useful therapeutic tool at multiple intervention periods: prior to the
development of fibrotic lesions, during the development of fibrotic lesions, and to
reengineer tissue once fibrotic lesions have progressed. The further development of this
therapeutic cell population, and the investigation of their mechanisms of action, will
surely continue to elucidate the potential for AFSC in this and numerous other injury and
disease states.
110
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APPENDICES
Appendix 1 AFSC modulation of the acute inflammatory cytokine milieu in both
BAL and tissue following bleomycin induced lung injury. (A) Table of all cytokine
modulations detected in BAL. (B) Graph of samples from BAL extracts that were
moderately, but not statistically significantly modulated. (C) Table containing all
cytokine modulations detected in tissue homogenates. (D) Graph of samples from tissue
homogenates that were moderately, but not statistically significantly modulated.
139
Appendix 2 AFSC modulation of the acute inflammatory cellular populations in
BAL following bleomycin induced lung injury. (A) Total cell count modulations
detected in BAL. (B) Differential BAL macrophage analysis. (C) Differential BAL
lymphocyte analysis. (D) Differential BAL neutrophil analysis. Distributions for B-D
are presented as box plots with lines at the lower quartile, median and upper quartile,
whiskers are representative of the minimum and maximums excluding outliers.
Abstract (if available)
Abstract
Idiopathic Pulmonary Fibrosis (IPF) is a chronic, progressive and fatal lung disease, with no effective or definitive treatment other than lung transplantation. Stem cell therapy has the potential to treat IPF through the specific targeting of therapeutic properties or compounds to inhibit or delay disease progression and aid in repair at specific sites of injury. Furthermore, as is the case with most progressive diseases, once the progression of fibrosis has diminished organ function to the point where transplantation is necessary, the use of stem cells to create bioengineered organoid units provides a therapeutic strategy reliant only upon organ reengineering time, and not on donor availability. This dissertation investigates the use of Amniotic Fluid Stem Cells as a therapeutic intervention strategy and a candidate cell population for organ engineering. ❧ The potential for amniotic fluid stem cell (AFSC) treatment to inhibit the progression of fibrotic lung injury has not been described. We have previously demonstrated that AFSC can attenuate both acute and chronic-fibrotic kidney injury through modification of the cytokine environment. Fibrotic lung injury, such as in Idiopathic Pulmonary Fibrosis (IPF), is mediated through pro-fibrotic and pro-inflammatory cytokine activity. Thus, we hypothesized that AFSC treatment might inhibit the progression of bleomycin-induced pulmonary fibrosis through cytokine modulation. In particular, we aimed to modulate the pro-fibrotic cytokine CCL2, which is increased in human IPF patients and is correlated with poor prognoses, advanced disease states and worse fibrotic outcomes. The impact of intravenous AFSC given at acute (day 0) or chronic (day 14) intervention time points after bleomycin injury were analyzed at either day 3 or day 28 post-injury. AFSC treatment at either day 0 or day 14 post-bleomycin injury significantly inhibited collagen deposition and preserved pulmonary function. CCL2 expression increased in bleomycin-injured bronchoalveolar lavage (BAL), but significantly decreased following AFSC treatment at either day 0 or at day 14. AFSC were observed to localize within fibrotic lesions in the lung, showing specific targeting of AFSC to the area of fibrosis. We also observed that MMP2 was transiently increased following AFSC treatment. Increased MMP2 activity was further associated with cleavage of CCL2, rendering it a putative receptor antagonist for CCR2, which we surmise is a potential mechanism for CCL2 reduction in BAL following AFSC treatment. ❧ When fibrotic lung injury has progressed beyond the point that a therapeutic intervention would be effective, organ transplant remains the only viable option. With the scarcity of donor lungs, researchers have begun investigating the potential for whole lung engineering. The complexity of the architecture and cell populations contained within the lung, and specifically the alveolus, present a challenge. Thus, in vitro engineering of bioartificial lungs require the selection of an appropriate candidate cell population and a physiologically relevant scaffold. Our preliminary data indicates that AFSC are a viable candidate cell population that express type I and type II alveolar epithelial lineage markers as well as stem cell markers. Furthermore, our data demonstrates that we have developed detergent/enzymatic protocols that efficiently remove all cellular material from lung extracellular matrices, leaving behind biological scaffolds that AFSC can be cultured on. Finally, we demonstrate that throughout culture on these scaffolds, AFSC secrete growth and angiogenic factors, as well as anti-inflammatory, proliferative and pluripotent factors, all critical for bioengineering functional organoid units. ❧ Based on the entirety of this data, we have concluded that AFSC have the potential to inhibit the development or progression of fibrosis in a bleomycin injury model during both acute and chronic remodeling events and have the potential to repopulate decellularized lung matrix as an in vitro bioarticifical engineering strategy.
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University of Southern California Dissertations and Theses
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Asset Metadata
Creator
Garcia, Orquidea Helen
(author)
Core Title
Immunomodulatory and regenerative potential of amniotic fluid stem cells as a treatment strategy for pulmonary fibrosis
School
Keck School of Medicine
Degree
Doctor of Philosophy
Degree Program
Systems Biology and Disease
Publication Date
04/28/2015
Defense Date
03/18/2013
Publisher
University of Southern California
(original),
University of Southern California. Libraries
(digital)
Tag
amniotic fluid stem cells (AFSC),CCL2,MMP2,OAI-PMH Harvest,pulmonary fibrosis,regenerative medicine
Language
English
Contributor
Electronically uploaded by the author
(provenance)
Advisor
Shi, Wei (
committee chair
), Warburton, David (
committee chair
), Driscoll, Barbara (
committee member
), Keens, Tom (
committee member
), Perin, Laura (
committee member
)
Creator Email
orchid1284@aol.com,orogers@usc.edu
Permanent Link (DOI)
https://doi.org/10.25549/usctheses-c3-247469
Unique identifier
UC11287850
Identifier
etd-GarciaOrqu-1588.pdf (filename),usctheses-c3-247469 (legacy record id)
Legacy Identifier
etd-GarciaOrqu-1588.pdf
Dmrecord
247469
Document Type
Dissertation
Rights
Garcia, Orquidea Helen
Type
texts
Source
University of Southern California
(contributing entity),
University of Southern California Dissertations and Theses
(collection)
Access Conditions
The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the a...
Repository Name
University of Southern California Digital Library
Repository Location
USC Digital Library, University of Southern California, University Park Campus MC 2810, 3434 South Grand Avenue, 2nd Floor, Los Angeles, California 90089-2810, USA
Tags
amniotic fluid stem cells (AFSC)
CCL2
MMP2
pulmonary fibrosis
regenerative medicine