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The calcium-sensing receptor in the specification of normal and malignant hematopoietic cell localization in the bone marrow microenvironment
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The calcium-sensing receptor in the specification of normal and malignant hematopoietic cell localization in the bone marrow microenvironment
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Content
THE CALCIUM-SENSING RECEPTOR IN THE SPECIFICATION OF NORMAL
AND MALIGNANT HEMATOPOIETIC CELL LOCALIZATION IN THE
BONE MARROW MICROENVIRONMENT
by
Ben S. Lam
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
(GENETICS, MOLECULAR AND CELLULAR BIOLOGY)
August 2012
Copyright 2012 Ben S. Lam
ii
Acknowledgements
It has been quite some time since I embarked on this journey to pursue my graduate
degree. Along the way, I have met many people who have made the journey memorable
and irreplaceable. First and foremost, I would like to thank my mentor, Dr. Gregor
Adams, PhD, for being the best mentor a student could ever ask for. Under Gregor’s
guidance and support, I have become a more independent and sophisticated thinker,
which will definitely be beneficial to me in my future career in the science field. As his
graduate student, I have also been very lucky to be able to have a lot of freedom in
research, which sharpened my creativity. Gregor never ceases to amaze me with his
memory. Whenever I had trouble finding a certain journal article and could not remember
where to find it, he would provide me with the author’s name, year, and the name of the
journal. His scientific knowledge is equally impressive, and it is my goal to reach his
level and beyond.
Of course, it is more about the people who you work with that makes the whole journey
more memorable. I would like to thank everyone in the laboratory for always being there
during all the good and bad times. Dr. Xiaoying Zhou, MD, PhD, has been a great
colleague during my entire journey, and whenever I had problems with my experiments,
she was always a valuable person for discussion. My fellow graduate student, Sapna
Shah, has been a good friend who I have shared a lot of laughs with, and I will definitely
not forget about her drawer where she hides all the cookies and brownies from tea time. I
also want to thank Tassja Spindler for being such a good sport when I needed her help
iii
with my experiments, though she calls me a “slave-driver” now for making her work too
much at times. I also would like to thank the former members of the Adams Lab. First, I
want to thank our former graduate student, Dr. Narges Rashidi, PhD, for being a great
friend in this entire journey. I am so proud of her, and at the same time very jealous, that
she is in London for her post-doctoral fellowship. I also want to thank our former
laboratory technicians, Sali Liu, Alan Tseng, and Tigue Tozer for all their help and fun
memories along this journey. Last, I also want to thank our former lab manager, Dilani
Rosa, for both her technical assistance with histology and also for planning all the fun
events.
During the past five years, I have met people who have influenced my life in various
ways. First, I would like to acknowledge a very important person in my life, Chris
Terwilleger. Chris has seen every personal hardship and struggle that has come along my
way, especially during this past year, and has always been supportive and caring even
though I might not be the best person to be around with. He has changed my life and
instilled confidence in me, and although I am not the most expressive person, I do
appreciate his presence in my life deeply in my heart. My journey would not have been
the same without my good friends who started the journey with me at the same time:
Yvonne Leung, Christine Cao, Lauren Geary, and Sarah Utley. Yvonne has been a very
good friend of mine during this whole journey and has always been a fun person to
interact with, and I sincerely wish the best for her in the upcoming years and beyond.
Christine is definitely the one to beat in terms of working long, productive hours in the
iv
laboratory. She is also a very good friend of mine who has seen me through my personal
struggles, and has also been a great help whenever I needed some reagents for my
experiments. I also want to thank Lauren for being such a caring friend during this
journey, and for baking all those delicious cupcakes and cookies that always brightened
my day. Last, but not least, I want to thank Sarah for being the most proactive friend in
the group and planning the get-together events. These memories would not have been the
same without any one of you!
Last, but not least, I am also grateful to the present and former staff at the FACS core: Dr.
Cynthia Cunningham, PhD, and Lora Barsky. In the beginning of this journey, I knew
nothing about flow cytometry, but they have both taught me so much during the past
years that flow cytometry is now one of my strongest areas of expertise. I also would like
to thank the members of my PhD Dissertation Committee, Dr. Preet Chaudhary, MD,
PhD, and Krzysztof Kobielak, MD, PhD, for their insightful comments and suggestions. I
also would like to acknowledge the administrative staff in the PIBBS and GMCB
programs: Dawn Burke, Raquel Gallardo, and Marisela Zuniga, for all their help and
support in the past five years.
v
Table of Contents
Acknowledgements ........................................................................................................... ii
List of Tables ................................................................................................................... vii
List of Figures ................................................................................................................. viii
Abbreviations .....................................................................................................................x
Abstract ........................................................................................................................... xvi
Chapter 1: Introduction ....................................................................................................1
1.1 The Development of Hematopoiesis ..............................................................................1
1.2 Adult Hematopoiesis: Localization of Hematopoietic Stem Cells in the
Bone Marrow Microenvironment ........................................................................................9
1.3 The Calcium-Sensing Receptor ...................................................................................23
1.4 The Human Hematopoietic System .............................................................................40
1.5 Multiple Myeloma .......................................................................................................46
Chapter 2: Pharmacologic Modulation of the Calcium-Sensing
Receptor Enhances Hematopoietic Stem Cell Lodgment in the Adult
Bone Marrow ....................................................................................................................53
2.1 Abstract ........................................................................................................................53
2.2 Introduction ..................................................................................................................54
2.3 Materials and Methods .................................................................................................57
2.4 Results ..........................................................................................................................76
2.5 Discussion ....................................................................................................................93
Chapter 3: Calcium-Sensing Receptor-Mediated Regulation of Cell
Proliferation and Interactions with the Bone Marrow
Microenvironment in Multiple Myeloma ......................................................................98
3.1 Abstract ........................................................................................................................98
3.2 Introduction ................................................................................................................100
3.3 Materials and Methods ...............................................................................................103
3.4 Results ........................................................................................................................124
3.5 Discussion ..................................................................................................................143
vi
Chapter 4: The Calcium-Sensing Receptor Specifies a Distinct Cell
Population in the Human Hematopoietic System .......................................................150
4.1 Abstract ......................................................................................................................150
4.2 Introduction ................................................................................................................152
4.3 Materials and Methods ...............................................................................................155
4.5 Discussion ..................................................................................................................167
Chapter 5: Concluding Remarks and Future Perspectives .......................................170
5.1 Concluding Remarks ..................................................................................................170
5.2 Future Perspectives ....................................................................................................174
Bibliography ...................................................................................................................183
vii
List of Tables
Table 2.1 Antibody list 61
Table 3.1 Information in regards to the sources from which the
human MM cell lines were derived 103
Table 3.2 Culture conditions for the three human MM cell lines 104
Table 3.3 Primers for subcloning 110
Table 3.4 Primers for PCR screens 112
Table 3.5 Primers for DNA sequencing 113
viii
List of Figures
Figure 1.1 CaR signaling transduction 30
Figure 1.2 Hierarchy of the hematopoietic system 40
Figure 2.1 CFU-C assay 60
Figure 2.2 Gating schemes for FACS 62
Figure 2.3 CAFC assay 64
Figure 2.4 Cell cycle analysis 67
Figure 2.5 Effects of Cinacalcet treatment on CaR signaling and
expression 77
Figure 2.6 Effects of CaR stimulation on hematopoietic progenitor and
primitive cell activity in vitro 79
Figure 2.7 CaR stimulation does not alter cell cycle status, or cell
survival, but significantly increases HSC adhesion to
collagen I 81
Figure 2.8 Collagen I is expressed ubiquitously along the bone 83
Figure 2.9 Cinacalcet treatment enhances in vivo homing, lodgment,
and engraftment 85
Figure 2.10 CaR stimulation enhances CXCR4 signaling and cell
migration towards SDF-1α, but does not alter the
expression of adhesion molecules 89
Figure 2.11 A proposed model for the role of CaR in HSC lodgment in
vivo 93
Figure 3.1 Diagram of pGIPZ lentiviral vector carrying CaR
shRNAmir 109
Figure 3.2 Diagram of the FUIGW lentiviral vector carrying the CaR-
FLAG insert 113
ix
Figure 3.3 Expression of CaR mRNA in U-266, RPMI-8226, NCI-
H929, and a BM sample from a patient with MM 124
Figure 3.4 Immunocytochemistry of U-266, RPMI-8226, and NCI-
H929 cells for cell surface CaR 125
Figure 3.5 Effects of CaR reduction on MM cell growth in vitro 126
Figure 3.6 Effects of CaR overexpression on MM cell proliferation in
vitro 127
Figure 3.7 Effects of CaR overexpression on apoptosis and cell cycle
status 129
Figure 3.8 Effects of CaR knockdown on cell migration and cell
adhesion 131
Figure 3.9 Effects of CaR overexpression on cell migration and cell
adhesion 132
Figure 3.10 Cell surface expression of CXCR4 133
Figure 3.11 Chemosensitivity of MM cells in the presence of
bortezomib 134
Figure 3.12 Introducing MM cells into the SCID-hu mouse model 137
Figure 3.13 Adapting the SCID-hu model for the study of homing and
localization of MM cells 140
Figure 4.1 Expression levels of cell surface CaR in human BM, UCB,
MPB, and FL 161
Figure 4.2 Hematopoietic progenitor activities in human BM, UCB,
MPB, and FL as measured by CFU-C frequency 162
Figure 4.3 Primitive hematopoietic cell activities in human BM, UCB,
MPB, and FL 163
Figure 4.4 Cell cycle analysis based on the expression of cell surface
CaR in the primitive CD34
+
CD38
-
cell population 164
Figure 4.5 The engraftment potential of human FL CD34
+
CaR
-
and
CD34
+
CaR
+
cell populations in NSG mouse recipients 165
x
Abbreviations
2βMe β-mercaptoethanol
7-AAD 7-amino-actinomycin
ADP adenosine diphosphate
AGM aorta-gonad-mesonephros
bFGF basic fibroblast growth factor
BFU-E burst-forming unit erythroid
BM bone marrow
BM MNC bone marrow mononuclear cell
BMPR1A bone morphogenetic receptor type 1A
BSA bovine serum albumin
C3 Clostridium botulinum C3
CAFC cobblestone area forming cell
CAM cell adhesion molecule
cAMP cyclic adenosine monophosphate
CaR calcium-sensing receptor
CD cluster of differentiation
cDNA complementary DNA
CFSE carboxyfluorescein diacetate succinimidyl ester
CFU-E colony forming unit-erythroid
CFU-GM colony forming unit-granulocyte macrophage
CFU-S colony forming unit-spleen
xi
c-Kit cellular Kit
Col1a1 type 1 collagen α1
CXCL12 chemokine (C-X-C motif) ligand 12
CXCR4 chemokine (C-X-C motif) receptor 4
DiI 1,1'-Dioctadecyl-3,3,3',3'-Tetramethylindocarbocyanine
DiO 3,3'-Dioctadecyloxacarbocyanine
DKK-1 dickkopf-1
DMEM Dulbecco’s modified eagle medium
ECM extracellular matrix
ERK extracellular signal-regulated kinase
FACS fluorescence activated cell sorting
FAK focal adhesion kinase
FBS fetal bovine serum
FISH fluorescent in situ hybridization
FL fetal liver
G-CSF granulocyte colony-stimulating factor
GFP green fluorescent protein
GM-CSF granulocyte macrophage colony-stimulating factor
GPCR G protein-coupled receptor
Gy gray
G
αi
α subunit of the i-type heterotrimeric G proteins
G
αq
α subunit of the q-type heterotrimeric G proteins
xii
HA hyaluronic acid
HBSS Hank’s balanced salt solution
HEK human embryonic kidney
HGF hepatocyte growth factor
HLA human leukocyte antigen
HoxB4 homeobox B4
HPC hematopoietic progenitor cell
HSC hematopoietic stem cell
IACUC Institutional Animal Care and Use Committee
IGF insulin growth factor
IL interleukin
IP
3
phosphatidylinositol
JNK c-Jun N-terminal kinase
Ki67 nuclear protein associated with cellular proliferation
LFA-1 lymphocyte function associated antigen-1
Lin lineage
LTC-IC long-term culture-initiating cell
LT-HSC long-term hematopoietic stem cell
Ly5.1 CD45.1
Ly5.2 CD45.2
MAPK mitogen-activated protein kinase
mGluR metabotropic glutamate receptor
xiii
MGUS monoclonal gammopathy of undetermined significance
MIP macrophage inflammatory protein
MM multiple myeloma
MMP matrix metalloproteinase
MPB mobilized peripheral blood
MPP multipotent progenitor
NF-κB nuclear factor kappa-light-chain-enhancer of activated B
cells
NOD/SCID non-obese diabetic SCID
NPS R-467 type II calcimimetic
NSG NOD/SCID/IL-2Rg
c
-null
OPN osteopontin
P/S penicillin-streptomycin
p67 laminin receptor
PBS phosphate buffered saline
PI3K phosphatidylinositol-3 kinase
PKA protein kinase A
PKC protein kinase C
PLA
2
phospholipase A
2
PLC phospholipase C
PLD phospholipase D
pSP para-aortic splanchnopleura
PTH parathyroid hormone
xiv
PTHR1 parathyroid hormone receptor 1
PTH-rP parathyroid hormone-related protein
RANK receptor activator of nuclear factor-κB
RANK-L receptor activator of nuclear factor-κB ligand
rh recombinant human
Rh123 rhodamine123
rm recombinant mouse
RT-PCR reverse transcription-polymerase chain reaction
SCF stem cell factor
SCID severe combined immunodeficiency
SDF-1 stromal cell-derived factor 1
SHG second harmonic generation
SLAM signaling lymphocyte activation molecule
SNARF-1 seminaphtharhodafluor
SNO spindle-shaped N-cadherin
+
CD45
-
osteoblastic
SRE serum response element
ST-HSC short-term hematopoietic stem cell
tm-SCF transmembrane isoform of SCF
TNF tumor necrosis factor
trOPN thrombin-cleaved osteopontin
UCB umbilical cord blood
VCAM vascular cell adhesion molecule
xv
VEGF vascular endothelial growth factor
VLA very late antigen
WT wild-type
YS yolk sac
α-MEM minimum essential medium alpha
xvi
Abstract
Hematopoiesis occurs through complex interplay between hematopoietic stem and
progenitor cells and the supportive bone marrow (BM) microenvironment. Defined by the
unique ability to self-renew and differentiate into all of the necessary blood cells to
support lifelong hematopoiesis, hematopoietic stem cells (HSCs) are spatially located in
BM stem cell niches. It was first discovered that primitive HSCs are localized at the
endosteal region of the BM. In this region, bone remodeling mediated by bone-forming
osteoblasts and bone-resorbing osteoclasts dynamically alters the endosteal surface and
releases Ca
2+
ions from the bone. HSCs are able to sense extracellular Ca
2+
concentrations through the cell surface expression of the calcium-sensing receptor (CaR).
Previous studies demonstrated the crucial role of the CaR in the localization of HSCs in
the endosteal BM niche, where CaR
-/-
HSCs displayed stem cell autonomous defects
including impaired lodgment and engraftment. We further explored the role of the CaR
on HSC functions by using a pharmacologic approach to stimulate the receptor ex vivo
with the clinically approved allosteric agonist, Cinacalcet. With CaR stimulaton, HSC
homing, lodgment, and engraftment in the endosteal BM niche were promoted via
mechanisms involving increased CXCR4 signaling and cell adhesion to major ECM
molecules, suggesting that HSC-niche interactions were enhanced to support
hematopoietic recovery in the recipient. Although the regulatory functions of the CaR in
HSC lodgment and localization have been investigated in normal hematopoiesis, the
functional role of the CaR in pathologic hematopoiesis is still poorly understood. In this
regard, we used a genetic approach to study the functional role of the CaR in regulating
xvii
the pathophysiology of multiple myeloma (MM), as this specific hematological
malignancy involves devastating bone destruction caused by an excessively active bone-
resorbing osteoclastic population, leading to heightened levels of Ca
2+
ions in the BM
endosteal region. We demonstrated that the CaR stimulates myeloma cell proliferation,
with mechanisms involving adhesive interactions with the BM microenvironment and
cell cycle entry. We also provided the first evidence that the CaR plays a
chemoprotective role in MM, as genetic manipulations of CaR expression altered the
chemosensitivity of myeloma cells to the chemotherapeutic agent, bortezomib. Finally,
we explored the clinical relevance of the CaR in the specification of a functionally
distinct HSC population in human hematopoietic tissues based on the cell surface
expression of the receptor. Interestingly, we discovered that human HSCs displaying cell
surface CaR have impaired hematopoietic stem and progenitor activity in vitro, but
similar in vivo engraftment potential in xenogeneic recipients compared to human HSCs
lacking cell surface CaR. Although the precise regulatory mechanisms remain to be
elucidated, our findings nonetheless suggest that intrinsic functional differences exist in
these human HSC populations. Collectively, these data have implications in developing
novel therapeutic strategies to enhance HSC engraftment in the transplantation setting by
stimulating the CaR, and also in targeting the CaR to abrogate the pathophysiology of
bone metastatic cancers by disrupting the complex interplay between the cancer cells and
the BM microenvironment.
1
Chapter 1: Introduction
1.1 The Development of Hematopoiesis
Hematopoiesis is defined as the process of blood cell formation from HSCs during
embryogenesis and throughout the lifetime of an animal. This process is intricately
regulated and stage-specific in order to meet the demands of rapid growth in the
developing embryo, and also to support and maintain a functional hematopoietic system
in the adult organism. Unlike adult hematopoiesis, which happens predominantly in the
BM, the pool of HSCs formed during embryonic hematopoiesis involves multiple
anatomical sites, beginning at the extraembryonic yolk sac (YS), the intraembryonic
para-aortic splanchnopleura (pSP), the aorta-gonad-mesonephros (AGM) region, the
placenta, and finally at the fetal liver (FL). Although the specific site of de novo HSC
genesis is unclear, it is generally agreed that the establishment of the hematopoietic
system is a complex process that occurs in two waves in utero: primitive and definitive
hematopoiesis.
1.1.1 Primitive Hematopoiesis
Primitive hematopoiesis occurs in the extraembryonic YS after gastrulation, when
mesodermal precursors migrate from the posterior region of the primitive streak to the
YS (E7.0-E7.5 in mouse; day 15-17 post-fertilization in human) (Huang et al., 2007a;
Tavian and Péault, 2005). These mesodermal precursors, known as hemangioblasts, are
the common precursor for early hematopoietic and endothelial cells (Robb et al., 1998;
2
Robertson et al., 1999). During primitive hematopoiesis, the earliest blood elements in
the hematopoietic system arise. These primitive elements consist mainly of large,
nucleated erythrocytes that express embryonic hemoglobin (ε and βH1) (Huang et al.,
1994; Medvinsky et al., 1993; Godin and Cumano, 2002). They form blood islands that
consist of clusters of hemogenic cells surrounded by a flattened endothelium (Ferkowicz
et al., 2005). However, because these early blood elements are mainly erythroid in nature,
it is unclear whether hematopoiesis originates in extraembryonic or intraembryonic sites.
In addition, because the onset of circulation (E8.5 in mouse; day 21 post-fertilization in
human) (Huang et al., 2007a; Tavian and Péault, 2005) occurs at the same time as
hematopoiesis, and of the difficulties inherent in tracking development during
mammalian embryogenesis, the origin of HSCs capable of contributing to the
establishment of definitive lifelong blood formation remains highly disputed.
1.1.2 Definitive Hematopoiesis
Definitive hematopoiesis describes the process of the generation of HSCs with the
capacity of self-renewal and long-term repopulation in transplant recipients. This second
wave of hematopoiesis occurs later in embryonic development, following the initiation of
somitogenesis, and contributes to the definitive HSC pool. Definitive hematopoiesis may
arise independently in both extraembryonic and intraembryonic sites, since multipotent
hematopoietic cells can be detected in both the extraembryonic YS and the
intraembryonic pSP region prior to FL colonization (Godin et al., 1995; Cumano et al.,
1993). However, multiple studies have documented that the intraembryonic AGM region,
3
developed from the pSP, is a source of definitive HSCs rather than the extraembryonic
YS (Cumano et al., 1996; Medvinsky and Dzierzak, 1996; Muller et al., 1994),
specifically the dorsal aorta area (de Bruijin et al., 2002). Using a transgenic zebrafish
model coupled with imaging and lineage tracing studies, a recent study elegantly
demonstrated that the first HSCs generated arise directly from the hemogenic
endothelielium lining the ventral wall of the dorsal aorta (Bertrand et al., 2010).
Furthermore, a study using isolated pSP and YS tissues before the onset of circulation
also demonstrated that the intraembryonic tissues contained long-term reconstituting
hematopoietic precursors that were capable of generating both lymphoid and myeloid
progenies (Cumano et al., 2001). However, because this study was done using explant
cultures obtained from the pSP and the YS, the intrinsic developmental pathways might
have been altered. Therefore, this study did not conclusively rule out the possibility of the
contribution of the YS to definitive hematopoiesis during embryonic development. The
placenta has recently been identified to harbor a large pool of multipotent HSCs capable
of generating clonogenic hematopoietic progenitors and giving rise to multilineage
engraftment in transplant recipients (Alvarez-Silva et al., 2003; Gekos et al., 2005;
Ottersbach and Dzierzak, 2005).
1.1.2.1 Establishment of Hematopoiesis in the FL
During embryonic development, the number of HSCs and other hematopoietic cell types
must increase substantially in order to keep pace with the growth of the developing fetus
and to constitute the hematopoietic system of the adult. However, before they colonize
4
the FL, HSCs are found in very low numbers in anatomical sites such as the YS and the
AGM. Therefore, in order to ultimately seed the BM and spleen for lifelong
hematopoiesis, HSCs generated at earlier developmental stages must expand in great
numbers. During fetal development, the FL is known to be a major site of HSC
expansion, maintenance, and differentiation (Paige et al., 1984; Ema and Nakauchi,
2000). It is believed that de novo HSC genesis does not occur in the FL, but rather the FL
is seeded by circulating HSCs after the onset of circulation (Johnson and Moore, 1975;
Houssaint, 1981). However, the relative contribution of YS, AGM, and placenta to the FL
and adult HSC pool is still unclear.
The first wave of hematopoietic seeding occurs when myeloerythroid progenitors that
generate definitive erythroid cells migrate from the YS through vitelline vessels to
colonize the FL (E9.5-E10.5 in mouse; week 4.5-5 in human) (Khurana and
Mukhopadhyay, 2008; Tavian and Péault, 2005; Mikkola and Orkin, 2006). The second
wave of hematopoietic seeding is defined by the the appearance of HSCs in the FL
(E11.5 in mouse; day 30 in human) (Mikkola and Orkin, 2006; Tavian and Péault, 2005),
the majority of which are likely to be derived from the AGM and the placenta and
migrate to the FL via the umbilical vessels. A previous study has also shown the
contribution of the YS to FL hematopoiesis by supplying the FL with a large number of
definitive HSCs (Kumaravelu et al., 2002). The FL supports the expansion of HSCs
starting from E12.5, where the number of HSCs increases exponentially until E15-E16
(Morrison et al., 1995; Ema and Nakauchi, 2000). In addition to supporting the expansion
5
of HSCs, the FL also plays an important role in producing differentiated hematopoietic
cells. In fact, the early FL is initially rich in CFU-Es (colony forming unit-erythroid) and
proerythroblasts, then myeloid and lymphoid progenitors accumulate with developmental
age (Mikkola and Orkin, 2006). Early studies have also shown that early B lymphocytes
can be detected in the FL during fetal development (Nossal and Pike, 1973; Denis et al.,
1984). The FL remains the predominant site of hematopoiesis until E16 when HSC
number declines rapidly (Morrison et al., 1995; Ema and Nakauchi, 2000), suggesting the
transitioning of the major site of hematopoiesis from the FL to the spleen and BM in late
fetal development.
1.1.2.2 Establishment of Hematopoiesis in the BM
As the last blood-forming tissue that develops in ontogeny (Tavian and Péault, 2005), the
BM is the primary site of hematopoiesis that occurs throughout the lifetime of an adult
animal. The vertebrate skeleton, composed of cartilage and bone, is developed from
mesenchymal condensations that give rise to either osteoblasts or chondrocytes for the
production of intramembranous bones or the generation of a cartilaginous framework for
mineral deposition, respectively (Olsen et al., 2000). Controlled by a myriad of
transcription factors, growth factors, cytokines, and extracellular matrix molecules, the
cartilaginous framework is eventually replaced by bone and BM by the process of
endochondral ossification (Olsen et al., 2000). During this process, surrounding
mesenchymal cells and blood vessels invade the cartilaginous bone rudiments, leading to
the formation of ossifying trabeculae separated by vascular spaces (Charbord et al.,
6
1996). The seeding of hematopoietic stem and progenitor cells is then facilitated by the
establishment of the vascular system in the developing bone (Mikkola and Orkin, 2006).
Hematopoietic progenitor activity is detectable in the fetal BM as early as E15.5 as
demonstrated by the presence of B cell progenitors in a competitive repopulation assay
(Christensen et al., 2004). Interestingly, this precedes the colonization of long-term
repopulating HSCs in the fetal BM, which occurs at E17.5 (Christensen et al., 2004). A
previous study showed that stromal cell-derived factor 1 (SDF-1), or chemokine (C-X-C
motif) ligand 12 (CXCL12), a ligand for chemokine (C-X-C motif) receptor 4 (CXCR4),
is a major chemokine required for attracting HSCs to the fetal BM, where SDF-1
-/-
embryos displayed severe impairment in HSC colonization in the fetal BM, but without
any significant effects on HSC numbers in the FL (Ara et al., 2003). In addition, because
SDF-1 is produced by the BM stromal cell components, it was suggested that the early
fetal BM microenvironment lacks the ability to attract HSCs, possibly through the
CXCR4/SDF-1 axis, and support their engraftment and self-renewal. Together, these
studies provide evidence for the importance of SDF-1 in attracting HSCs to the fetal BM
during development. On the other hand, the regulation of HSC homing to the adult BM
by the CXCR4/SDF-1 may be different from that to the fetal BM. Studies on HSC
homing to the adult BM showed that CXCR4
-/-
FL cells were able to migrate to the BM
and reconstitute hematopoiesis in lethally irradiated recipients, though with decreased
homing efficiency in more differentiated cells compared to progenitor cells (Ma et al.,
7
1999; Kawabata et al., 1999; Foudi et al., 2006). This suggests that CXCR4 may be
dispensable for the homing of hematopoietic progenitor cells to the adult BM.
1.1.2.3 Differences in FL HSCs and BM HSCs
Unlike the quiescent HSCs from the adult BM, HSCs from the FL are actively engaged in
cell division (Fleming et al., 1993) and can give rise to long-term reconstitution at higher
levels when transplanted into lethally irradiated recipients (Micklem et al., 1972;
Morrison et al., 1995; Rebel et al., 1996). Specifically, it was shown that 18-22% of the
BM HSCs are in the S/G
2
/M phase, while at least 40% of FL HSCs are actively cycling
(Fleming et al., 1993). This suggests that intrinsic differences exist in the mechanisms
that regulate FL HSC and BM HSC self-renewal. In addition, extrinsic differences
residing in the FL and BM microenvironments may also provide distinct signaling cues
that regulate HSC divisions. Unraveling the underlying cellular and molecular
mechanisms that regulate rapid HSC self-renewal in the FL may translate into clinical
applications, including as the expansion of functional HSCs in culture for transplantation.
In fact, efforts have been made to expand functional HSCs in culture using various FL
stromal cell lines and have identified one cell line with the capability of maintaining input
HSCs and expanding primitive progenitors (Moore et al., 1997). By comparing the
expression profiles of the FL-derived stromal cell lines with varying HSC supportive
abilities, multiple differentially expressed genes were identified that may represent the
molecular profile of a candidate stem cell niche (Hackney et al., 2002). The functions of
these genes in the regulation of HSC self-renewal remain to be investigated. Another
8
study demonstrated that FL CD3
+
Ter119
-
cells are capable of supporting and expanding
HSCs in culture, and that microarray expression analysis revealed the expression of
insulin-like growth factor (IGF)-2 and angiopoietin-like proteins in these supportive cells
(Zhang and Lodish 2004; Zhang et al., 2006). Understanding the mechanisms that
regulate HSC self-renewal will be instrumental not only for creating an ex vivo HSC
expansion culture system to advance HSC-based therapies for the treatment of
hematological disorders, but also for elucidating important regulatory pathways that
affect stem cell fate decisions.
9
1.2 Adult Hematopoiesis: Localization of Hematopoietic Stem Cells in
the Bone Marrow Microenvironment
1.2.1 Early Observations on the Spatial Distribution of Hematopoietic Progenitors
Approximately five decades ago, spleen colony forming units (CFU-S) were first
identified as progenitor cells with the ability to initiate colony formation in spleens of
mice recovering from radiation-induced depletion of hematopoietic tissues (Till and
McCulloch, 1961). CFU-Ss were demonstrated to be highly proliferative progenitor cells
that are capable of giving rise to myeloerythroid cells (Becker et al., 1963; Wu et al.,
1967). These cells were later demonstrated to give rise to lymphocytes (Wu et al., 1968).
At that time, CFU-S was equated with HSC because of its capacity to undergo unlimited
proliferation and differentiate into any blood cell lineages. Since the introduction of the
spleen colony assay for the study of HSCs, a series of studies were conducted to study the
localization of CFU-S in the BM. The former assumption that hematopoiesis is a random
process in the adult BM is argued by considerable evidence. Fractionation studies of the
mouse femoral marrow indicated that hematopoiesis is in fact a spatially and functionally
ordered process, with a nonrandom distribution of hematopoietic progenitor cells (HPCs)
in the adult BM (Lord and Hendry, 1972; Lord et al., 1975; Shoeters and Vanderboroght,
1980; Mason et al., 1989; Lord, 1990). Specifically, these studies suggested that there are
higher concentrations of CFU-S in the regions near bone surfaces than in the center of the
BM cavity (Lord and Hendry, 1972; Lord et al., 1975; Shoeters and Vanderboroght,
1980; Mason et al., 1989; Lord, 1990), with the highest frequency of undifferentiated
colony cells in the endosteal region of the bone (Lambertsen and Weiss, 1984). These
10
findings prompted the establishment of the stem cell niche hypothesis by Schofield,
which suggests that the true stem cell is maintained in close association with an
environmental niche that imposes on it the attribute of indefinite self-renewal capacity
and prevents it from differentiation and maturation (Schofield, 1978; Schofield, 1983).
When this association is perturbed, the stem cell becomes committed to a specific line of
blood cell development and loses its stem cell property (Schofield, 1978; Schofield,
1983). The stem cell niche hypothesis has remained largely undisputed and provided the
basis for studies focusing on the interactions of the HSC and niche components.
1.2.2 Hematopoietic Stem Cell Niches
1.2.2.1 The Endosteal BM HSC Niche
Since the introduction of the stem cell niche hypothesis, numerous studies have been
performed to improve our understanding of the extrinsic regulators of stem cell numbers
and functions, including self-renewal, differentiation, apoptosis, and mobilization.
Various cellular and extracellular matrix (ECM) components that comprise the stem cell
niches have been identified. Based on the early observations that CFU-Ss are present in
higher numbers near the endosteal bone surface, Taichman et al. hypothesized that bone-
forming osteoblasts may be the critical cellular component that supports HSC in vivo and
demonstrated that human osteoblasts purified from trabecular bones could support the
expansion of long-term culture-initiating cells (LTC-IC) and progenitor cells in vitro
(Taichman et al., 1996) by secreting hepatocyte growth factor (HGF) and granulocyte
colony-stimulating factor (G-CSF) (Taichman et al., 2001). To further support the
11
localization of primitive HSCs in the endosteum, Nilsson et al. demonstrated that a
primitive population of less metabolically active HSCs preferentially localized to the
endosteal region after transplantation (Nilsson et al., 2001). Taken together, these two
studies suggest that the endosteal region and osteoblastic lineage cells provide a favorable
microenvironment for HSC in vivo.
Direct in vivo evidence for the role of osteoblasts in HSC regulation and maintenance
came from two studies in which osteoblast numbers were manipulated experimentally. In
the first study conducted by Calvi et al., osteoblast-specific expression of a constitutively
active form of parathyroid hormone 1 receptor (PTHR1), the common receptor for
parathyroid hormone (PTH) and PTH-related protein (PTH-rP), was achieved using the
type 1 collagen α1 (Col1a1) promoter, which is specific to maturing and mature
osteoblasts (Calvi et al., 2003). These mutant mice displayed simultaneous increase in the
number of both osteoblasts and HSCs in the BM (Calvi et al., 2003). In the other study
conducted by Zhang et al., mutant mice with conditional inactivation of bone
morphogenetic receptor type 1A (BMPR1A), which is expressed by osteoblasts and most
hematopoietic lineages, displayed a simultaneous increase in the number of both spindle-
shaped N-cadherin
+
CD45
-
osteoblastic (SNO) cells and HSCs in the BM (Zhang et al.,
2003). This suggests that SNO cells lining the bone surface is a key cellular component
of the niche to support HSCs. Taken together, these two studies indicated that osteoblasts
are an essential component of the endosteal niche.
12
1.2.2.2 The Vascular BM HSC Niche
Evidence for the presence of a second specialized HSC microenvironment termed the
vascular BM HSC niche came forth when a large proportion of long-term repopulating
CD150
+
HSCs were observed to be in close contact with the BM sinusoidal endothelium
(Kiel et al., 2005). This study proposed the possibility of isolating highly purified HSCs
and other progenitors based on the combinations of signaling lymphocyte activation
molecule (SLAM) family members, a group of 10-11 cell surface receptors that are
tandemly arrayed at a single locus on chromosome 1 to regulate the proliferation and
activation of lymphocytes (Howie et al., 2002; Wang et al., 2004). The idea that
endothelial cells possess the capacity for HSC maintenance or expansion is no surprise
given that hematopoietic and endothelial cells both arise from a common embryonic
precursor called the hemangioblast (Huber et al., 2004). In fact, previous studies using
endothelial cells derived from the YS or the AGM have revealed the ability of these cells
to support and expand adult HSCs in vitro (Ohneda et al., 1998; Li et al., 2003). It is
important to note that BM sinusoidal endothelial cells are functionally distinct in their
ability to maintain repopulating HSCs in vitro because endothelial cells isolated from
various non-hematopoietic organs have little or no ability to maintain HSCs in vitro (Li et
al., 2004).
The existence of two HSC niches in the BM raises the question of how functionally
different HSCs from the vascular niche are compared to HSCs from the endosteal niche.
HSCs with label retaining activity are almost exclusively located in the endosteal niche
13
(Zhang et al., 2003), which indicates that this niche contains the most dormant HSCs.
Therefore, the endosteal niche might serve as storage for quiescent HSCs. In contrast,
HSCs from the vascular niche possess a proliferative capacity that allows them to rapidly
and robustly respond to hematological stress. In fact, the vascular niche was previously
predicted to form during HSC mobilization after myeloablation, when quiescent HSCs
detach from the endosteal niche and migrate to the vascular niche at the central marrow
region to re-establish hematopoiesis through proliferation and differentiation (Kopp et al.,
2005; Avecilla et al., 2004; Heissig et al., 2002). However, recent imaging studies
showed that the HSCs substantially proliferate in the endosteal region in irradiated
recipients to reconstitute hematopoiesis, suggesting that the endosteal niche can convert
to a stimulatory niche under stress (Lo Celso et al., 2009; Xie et al., 2009). In terms of
the spatial distinction between the two niches, it was discovered that all transplanted
hematopoietic progenitor and stem cells localized in close proximity to the vasculature,
but only at variable distances from the endosteum (Lo Celso et al., 2009). In addition, it
was demonstrated that there was a non-random co-association of osteoblastic cells and
the vasculature in the trabecular bone (Xie et al., 2009), and that the proposed dichotomy
between the two niches is actually not anatomically feasible in the calvarium because
osteoblasts were found to be perivascular (Lo Celso et al., 2009), further questioning the
real distinction between the endosteal and vascular niche. As a result, it remains to be
investigated the true functional differences between the endosteal and vascular niches in
the maintenance of HSCs.
14
1.2.3 Regulation of HSC Functions and Localization in the Adult BM
1.2.3.1 Early Observations
The signals that influence stem cell functions are transmitted from the niche to the stem
cell via direct cell contact, adhesion molecules, components of the ECM, growth factors,
and chemokines. Because of the fact that stem cells are extremely rare and in vivo
tracking technologies were underdeveloped, identifying and characterizing stem cell
niches were particularly challenging. Some of the early studies on the
microenvironmental effects on HSC localization were conducted using the anemic Sl/Sl
d
and W/W
v
mutant mice. The Sl and W alleles encode stem cell factor (SCF) and its
receptor c-Kit, respectively. The Sl defect results in a reduction of the ability in the
affected microenvironmental cells, such as fibroblasts and endothelial cells, to produce
SCF and to support hematopoiesis, while the W defect is intrinsic to hematopoietic cells
(Dexter and Moore, 1977). In these studies, lodgment was simply defined as the trapping
of normal stem cells from the circulating blood. Here, it was found that lodgment of
injected CFU is normal in the Sl/Sl
d
spleen, but erythrocytic commitment (loss of
multipotency) is affected (Wolf, 1974). In another study, the hematopoietic defects in
W/W
v
mice can be cured by transplanting BM cells obtained from Sl/Sl
d
mice (Dexter and
Moore, 1977), indicating a role of the microenvironment of the stem cells in supporting
self-renewal and differentiation. An in vivo transplantation study using the monoclonal
antibody ACK2 to selectively block the c-Kit receptor has shown a reduction in burst-
forming unit erythroid (BFU-E) and colony forming unit-granulocyte macrophage (CFU-
GM) retrieved from the BM and spleen, suggesting that interaction of SCF with the c-Kit
15
receptor mediates the lodgment of HPCs (Broudy et al., 1996). Altogether, these early
studies point to the importance of stem cell-niche interaction in mediating cell lodgment
as well as in sustaining normal hematopoiesis following transplantation into irradiated
recipients.
1.2.3.2 Adhesion Molecules and the ECM
In the adult mammal, hematopoiesis is restricted to the extravascular compartment where
HSCs are in contact or close proximity with a heterogeneous population of stromal cells
in the niche. Cellular interactions between HSCs and stromal cells involve tightly
coordinated participation of various cell adhesion molecules (CAM), including integrins,
selectins, sialomucins, and the immunoglobulin gene superfamily, which are
subsequently translated into signaling mechanisms in regulating the localization of cells
within the niche. Studies on the identification of molecules that define the anatomical
localization of HSCs are complicated by the extensive array of CAMs that are expressed
on HSCs (Simmons et al., 1997). In addition, this is further complicated by the broad
range of potential ligands correspondingly presented by BM stromal cells (Simmons et
al., 1997). One in vitro study explored the capacity of human HPCs to adhere to the ECM
secreted by human marrow fibroblasts and discovered that adherence to the matrix varied
both with the cell lineage and maturation stage of the progenitor, and that fibronectin may
be one ECM component involved in HPC adhesion (Coulombel et al., 1988). A study on
the spatial location of ECM proteins including fibronectin, collagen types I, III, and IV,
and laminin in murine femoral bone marrow by immunofluorescence labeling has
16
revealed distinct locations of each protein, supporting the notion that they have an
important role in the homing and lodgment of transplanted cells (Nilsson et al., 1998).
Whether there is a preference for a specific subset of primitive hematopoietic cells to
adhere to certain ECM proteins within the adult BM remains to be explored.
Many reports have documented the importance of β
1
integrins, particularly α
4
β
1
, also
known as very late antigen 4 (VLA-4), and α
5
β
1
or VLA-5, in modulating adhesive
interactions between HPCs and the cellular and ECM components that comprise the stem
cell niche (Simmons et al., 1992; Teixidó et al., 1992; Williams et al., 1991; Miyake et
al., 1991a; Miyake et al., 1991b). It has been shown that VLA-5 is expressed on mouse
and human long-term repopulating hematopoietic cells and binds to fibronectin in the
ECM, and that disruption of this binding can lead to decreased engraftment in the BM
(van der Loo et al., 1998). A recent study that focuses on the functional implications of α
4
integrin deficiency induced by Mx.cre deletion has shown that α
4
integrin-deficient HPCs
accumulate in the peripheral blood (Scott et al., 2003). Furthermore, in transplantation
studies, α
4
-/-
cells displayed impaired homing to the BM, and short-term engraftment was
severely delayed (Scott et al., 2003). Despite the number of studies documenting
impaired homing in α
4
-deficient cells, the ability of these cells to localize to different
anatomic regions within the BM was not examined until recently. Using fluorescent-
labeled Lin
-
c-Kit
+
cells harvested from the murine BM, it was discovered that although
α
4
-deficient cells were capable of interstitial transmigration within the BM, the retention
of these cells in the endosteal region following transplantation into irradiated recipients
17
was compromised compared to normal control cells (Jiang et al., 2009). Altogether, these
studies suggest the importance of β
1
integrins in the homing and lodgment of HSCs
within specialized regions of the BM microenvironment.
To identify cell surface molecules with potential roles in influencing the site of HSC
lodgment following homing to the BM, a group conducted an extensive analysis of CAM
expression on Lin
+
and Lin
-
Rh123
dull
populations and found that the Lin
-
Rh123
dull
population displayed a reduced cell surface expression of the single-chain carbohydrate
hyaluronic acid (HA) (Nilsson et al., 2003). HA is also a component of the ECM within
the BM microenvironment and cell surface expression affects the adhesion, motility, and
growth of many cell types (Fraser et al., 1997). To investigate the role of HA in the
lodgment of transplanted HSCs, HSCs were treated with hyaluronidase prior to
transplantation in order to facilitate the enzymatic removal of HA. The results showed a
significant alteration in spatial distribution of the transplanted HSCs, suggesting a key
role for HA in lodgment (Nilsson et al., 2003). Another study on the cell-matrix
interaction was focused on laminins, which are a group of ECM proteins expressed in
BM that HPCs can bind to through their cognate receptors (Gu et al., 1999; Gu et al.,
2003; Siler et al., 2000). Specifically, expression of the p67 laminin receptor was found
on erythroid HPCs, and that blocking p67 binding of donor cells with anti-functional
antibody could lead to reduced BM homing of BFU-Es (Bonig et al., 2006). However,
whether nonerythroid HPCs can interact with laminins through other receptors is still
unanswered.
18
A recent study on the role of β
1
integrins have demonstrated their interactions with a
thrombin cleavage fragment of osteopontin (OPN), which is a multifunctional acidic
glycoprotein expressed by osteoblasts within the endosteal region of the BM and plays a
role in suppressing HSC proliferation and regulating their lodgment within the BM after
transplantation (Grassinger et al., 2009). In this study, thrombin-cleaved OPN (trOPN)
was shown to be a chemoattractant for HSCs through interaction with α
9
β
1
and α
4
β
1
integrins (Grassinger et al., 2009). Furthermore, using the transgenic Opn
-/-
mouse model,
it was depicted that the lack of OPN in the hematopoietic microenvironment results in a
significant impairment in the retention of HSCs in the endosteal region where OPN is
expressed at high levels (Grassinger et al., 2009). However, despite the absence of OPN
in the hematopoietic microenvironment in Opn
-/-
mice, homing of HSCs isolated from the
endosteal region of the BM was not altered (Grassinger et al., 2009). The mechanism
behind this remains unclear and further investigation is anticipated.
1.2.3.3 Growth Factors and Chemokines
During ontogeny and adult life, HSC migration to various organ sites and retention in a
specific anatomic location is mediated by soluble factors and chemokines. One
interesting study that looks at CXCR4 expression and function in human CD34
+
cells
obtained from distinct tissue sources have demonstrated that despite lower levels of
CXCR4, responsiveness of the cells to SDF-1 was proportionally the highest in cells
derived from the BM (Glodek et al., 2007). This suggested that preserved chemokine
receptor signaling was highly associated with BM rather than blood localization, further
19
confirming the role of SDF-1 and CXCR4 in HSC retention. A recent study that
attempted to investigate the mechanisms of the CXCL12-CXCR4 pathway has indicated
the involvement of focal adhesion kinase (FAK) in mediating the chemotactic effect of
CXCL12 in HPCs, as Cre-mediated FAK deletion resulted in impaired CXCL12-induced
migration (Glodek et al., 2007). In summary, this study raises the possibility that FAK
may function as an intermediary in signaling pathways controlling hematopoietic cell
lodgment and lineage development.
As mentioned previously, the interaction of SCF and its receptor c-Kit has been
implicated in regulating HSC lodgment following transplantation. In fact, SCF can exist
as a soluble form or a transmembrane isoform through alternative splicing to mediate the
adhesion of cells that express its receptor (Flanagan et al., 1991; Long et al., 1992). It is
important to note that expression of Sl
d
mutant cDNA in transfected cells produced
functionally active soluble SCF, but without the expression of the transmembrane
isoform of SCF (tm-SCF) (Flanagan et al., 1991). To analyze role of tm-SCF in cell
lodgment, fluorescent-labeled HSCs were transplanted into Sl/Sl
d
mouse recipients and
the spatial distribution of the transplanted HSCs were monitored (Driessen et al., 2003).
Compared to WT recipients, there was a significant reduction in the number of
transplanted HSCs within the endosteal region of the BM in Sl/Sl
d
recipients (Driessen et
al., 2003). Interestingly, HSC homing to the BM was not impaired, as equivalent number
of donor cells were detected per central femoral section in both types of recipients
20
(Driessen et al., 2003). Altogether, these studies suggest a specific key role for tm-SCF in
the lodgment of HSC within the endosteal region of the BM.
1.2.4 Technologies for Studying the Localization of Transplanted HSCs
Previously, in-depth analysis of BM transplantation was restricted by the technologies
available to detect donor cells in the recipient. To allow cell tracking, techniques
including immunophenotypic analysis of cell surface markers expressed on donor cells,
congenic models such as the Ly5.1/5.2 mouse, southern blot analysis to detect Y
chromosome-specific DNA sequences in male donor cells within female recipients, and
fluorescent in situ hybridization (FISH) were utilized (Hendrikx et al., 1996; Spangrude
et al., 1995; Lamar and Palmer, 1984; Slezak and Muirhead, 1991; Nilsson et al., 1996;
Nilsson et al., 1997). Some of these techniques have the problems of low sensitivity and
specificity, while others introduce too much background for the positive signal to be
readily detected. Although the use of fluorescent dyes such as PKH-26 has shown to
provide information on the cell cycle status of transplanted HSCs in the BM and spleen
(Lanzkron et al., 1999), this technique is generally limited by the number of cell divisions
the transplanted cells make, with each cell division resulting in a halving of cellular
fluorescence (Parish, 1999). As a result, the use of fluorescent dyes to track transplanted
HSCs may be limited to short-term analysis. Depending on the study objective, this may
be sufficient to track specifically the transplanted HSCs that are homed and lodged in the
BM microenvironment but not their progeny.
21
One elegant study by Nilsson et al. incorporated the use of the fluorescent dye
carboxyfluorescein diacetate succinimidyl ester (CFSE) to label different subpopulations
of BM cells for transplantation in nonablated recipients and allow for post-transplant
visualization and lodgment analysis (Nilsson et al., 2001). This study clearly
demonstrated that there is a distinction in the spatial distribution of HSCs and HPCs,
according to their respective level of lineage commitment. Specifically, although the
majority of both Lin
-
and Lin
+
cells localized in the central marrow region immediately
after transplantation, Lin
-
cells selectively redistributed away from the central marrow
region and predominantly localized in the endosteal region, whereas Lin
+
cells selectively
redistributed away from the endosteal region and predominantly localized in the central
marrow region. This redistribution process was shown to be nonrandom, and the results
supported the presence of HSC niches within the endosteal region of the BM (Nilsson et
al., 2001).
Recently, real-time live imaging has been utilized to track the migration of transplanted
HSCs to the BM in irradiated and non-irradiated recipients (Xie et al., 2009; Lo Celso et
al., 2009). In one study, green fluorescent-protein-expressing (GFP
+
) HSCs (Flk2
-
Lin
-
Sca-1
+
c-Kit
+
) were transplanted into the recipient and the migration and proliferation of
the transplanted HSCs were monitored by ex vivo real-time imaging technology (Xie et
al., 2009). Imaging analyses from the study showed that HSCs home to the BM through
the vascular system, and that they are randomly distributed throughout the BM in non-
irradiated recipients but tend to home to the endosteal region of the BM in irradiated
22
recipients (Xie et al., 2009). In addition, expansion of homed HSCs in irradiated
recipients was also observed using an immunoassay with the Ki67 marker (Xie et al.,
2009). Another study utilized a combination of high-resolution confocal microscopy and
two-photon video imaging to track individual HSCs in the calvarium BM following
transplantation into irradiated and c-Kit receptor-deficient recipients (Lo Celso et al.,
2009). This study also demonstrated that transplanted HSCs tend to home and lodge in
the endosteal region of the BM in irradiated recipients, with the long-term HSC localizing
closest to the endosteum and osteoblasts and more mature subsets localizing
progressively farther away (Lo Celso et al., 2009). HSCs that were transplanted into c-Kit
receptor-deficient recipients also tend to home and lodge in the endosteal region of the
BM, suggesting the possibility of overtaking endogenous HSC production in the
recipients (Lo Celso et al., 2009). In summary, these studies provide an alternative way to
track transplanted HSCs in the BM of the recipients, further confirming the tendency for
HSCs to localize in the endosteal region of the BM.
23
1.3 The Calcium-Sensing Receptor
1.3.1 Discovering the Cell Surface Receptor with an Extracellular Ca
2+
-Sensing
Mechanism
The CaR is a G protein-coupled receptor (GPCR) that plays a key role in calcium
homeostasis as well as in many other biological processes. Its primary physiological role
is the maintenance of constant blood Ca
2+
levels (1.1-1.3mM) through continuous
adjustments of PTH release from the parathyroid chief cells that are highly sensitive to
the small changes in extracellular Ca
2+
levels (Saidak et al., 2009a). The CaR is able to
mediate specific cellular functions in response to various extracellular messages,
including but not limited to its physiological ligand, Ca
2+
. Long before the discovery of
the CaR, the importance of Ca
2+
was revealed when it was discovered be a crucial
element for the contraction of isolated hearts (Ringer, 1883). Since then, the role of Ca
2+
in the regulation of various physiological functions in biological systems has been
recognized, and more importantly the requirement for organisms to maintain calcium
homeostasis (Carafoli, 2003). Systemic calcium homeostasis is maintained primarily
through the tight regulation PTH secretion in response to extracellular calcium
concentrations, followed by the action of the secreted PTH on the targeted tissue to
absorb or release calcium (Magno et al., 2011). However, evidence documenting the
existence of a surface receptor sensitive to extracellular calcium did not arise until the
1980s, when accumulating studies reported the receptor-mediated mobilization of
intracellular Ca
2+
with increases in extracellular Ca
2+
(Brown et al., 1987; Kifor and
Brown, 1988; Shoback et al., 1988), even in the absence of Ca
2+
transmembrane influx
24
(Nemeth and Scarpa, 1987). Finally, in 1993, the CaR was cloned from the bovine
parathyroid and characterized to possess a cell surface Ca
2+
-sensing mechanism that
regulates PTH secretion to maintain system calcium homeostasis (Brown et al., 1993).
1.3.2 Structural and Functional Properties of the CaR
The first CaR identified was 5276bp long and had an open reading frame of 3255bp that
encoded a protein of 1085 amino acids (Brown et al., 1993). In 1995, the human CaR
equivalent, consisting of seven exons, was cloned from an adenomatous parathyroid
gland (Garrett et al., 1995). The first exon is a 5’-untranslated region possibly for the
alternative splicing of the parathyroid CaR mRNA, while the other six exons encode a
protein of 1078 amino acids with 93% sequence similarity to the bovine CaR (Garrett et
al., 1995). Similar to the bovine CaR, the protein sequence of the human CaR also
showed potential sites for N-linked glycosylation and phosphorylation by protein kinase
C as well as a seven membrane-spanning region characteristic of GPCRs (Garrett et al.,
1995). Furthermore, as with the bovine CaR when expressed in Xenopus laevis oocytes,
the isolated human CaR clone also displayed a pharmacologic profile resembling to that
observed in parathyroid cells (Garrett et al., 1995). Specifically, the expressed CaR
responded to a range of CaR agonists besides Ca
2+
in a similar fashion and potency as the
native receptor (Garrett et al., 1995).
Like other GPCRs, the CaR three structural domains: a large, predominantly hydrophilic
amino-terminal domain of 612 amino containing a hydrophobic segment characteristic of
25
eukaryotic signal sequences and 9 potential N-linked glycosylation sites; a central core of
250 amino acids containing seven potential membrane-spanning helices characteristic of
the GPCR superfamily; and a hydrophilic carboxy-terminal domain of 222 amino acids
(Brown et al., 1993).
1.3.2.1 The Extracellular Domain of the CaR
The extracellular domain of the CaR is important in the proper functioning of the
receptor, as naturally occurring mutations in this domain have been linked to calcium
homeostatic disorders (Hu and Spiegel, 2007). In addition, the extracellular domain has
been proposed to contain sites of ligand binding (Reyes-Cruz et al., 2003). Specifically,
mutational studies have revealed that amino acids Ser147, Ser170, Asp190, Tyr218, and
Glu297 are critical to the proper binding of the physiological ligand, Ca
2+
, to the receptor
(Huang et al., 2007b; Silve et al., 2005). Recently, the CaR was identified to contain five
Ca
2+
-binding sites contributing to a cooperative Ca
2+
response (Huang et al., 2009).
1.3.2.2 The Transmembrane Domain of the CaR
The transmembrane domain spans residues 613-862 and contains seven hydrophobic
regions that form helices linked by alternating intracellular and extracellular loops
(Magno et al., 2011). It was proposed that when a ligand binds to the extracellular
domain of the GPCR, conformational changes in the transmembrane domain result, and
this subsequently leads to the promotion of signal transduction (Wess, 1997). A previous
study reported that a missense mutation in the transmembrane domain of the CaR results
26
in constitutive activation of the receptor, without depending on the binding of Ca
2+
(Zhao
et al., 1999). In addition, the transmembrane domain is involved in receptor dimerization
through non-covalent, possibly hydrophobic, interactions (Zhang et al., 2001), and that
dimerization of the receptor has functional implications (Pace et al., 1999; Ray et al.,
1999). CaR allosteric modulators such as calcimimetics and calcilytics were identified to
interact with the transmembrane domain to cause conformational changes of the receptor
(Petrel et al., 2004; Hu et al., 2006)
1.3.2.3 The Intracellular Domain of the CaR
The carboxyl terminal intracellular domain of the CaR is actually the least conserved
domain among different species and the most variable domain due to alternative splicing
(Pin et al., 2003). This domain was predicted to contain three protein kinase C (PKC)
phosphorylation sites at Thr888, Ser895, and Ser915 (Bai et al., 1998a), and mutations at
Thr888 led to both a decrease in responsiveness of the CaR to agonist stimulation and a
reduction in sensitivity to PKC activity (Jiang et al., 2002). PKC is actually involved in
inhibiting CaR activity by negative feedback. Specifically, when activated by the CaR,
PKC in turn phosphorylates the receptor to inhibit CaR activity, ultimately preventing G
protein subtypes from interacting with the region of the receptor critical for releasing
Ca
2+
from internal stores (Jiang et al., 2002; Davies et al., 2007). In addition to the PKC
phosphorylation sites, there are two putative protein kinase A (PKA) phosphorylation
sites, Ser899 and Ser900 in the intracellular domain of the CaR. However, it was
previously reported through pharmacological inhibition studies that blocking PKA alone
27
does not increase CaR activity, and that both PKA and PKC synergize in inhibiting the
CaR (Bösel et al., 2003).
1.3.2.4 Dimerization of the CaR Has Functional Implications
A previous study performed on transfected human embryonic kidney (HEK293) cells has
shown that the CaR mostly exists in the form of a dimer (Bai et al., 1998b). In fact,
dimerization of the CaR is functionally important for intracellular signaling and can be
achieved via disulfide linkages between cysteine resides in the extracellular domain
(covalent interactions) and hydrophobic interactions in the transmembrane domain (non-
covalent interactions) (Pace et al., 1999; Ray et al., 1999; Zhang et al., 2001; Bai 2004).
Specifically, it was documented that Cys-129 and Cys-131 are important in mediating
disulfide-linked dimerization of the CaR, where Cys-129 and Cys-131 mutants displayed
reduced dimerization of the CaR on the cell surface (Zhang et al., 2001). In contrast, Cys-
101 and Cys-236 substantially reduced overall expression of the CaR, but without
substantial conversion of dimers to monomers, suggesting the participation of non-
covalent interactions in the dimerization of the CaR (Zhang et al., 2001).
1.3.3 Sequence Homology of the CaR in Different Species
The CaR is well conserved across different species such as human, rat, and rabbit,
sharing more than 90% homology to the bovine CaR. (Aida et al., 1995; Riccardi et al.,
1995; Butters et al., 1997). In terms of homology between the CaR and other GPCRs, the
bovine CaR and the metabotropic glutamate receptors (mGluR) share limited but overall
28
significant homology in amino acid sequences (19-25% identity, 25-30% similarity for
mGluR1-6) (Brown et al., 1993). Both receptors belong to a subgroup of GPCRs known
as family 3 (or C) with a characteristically large extracellular domain called a Venus
flytrap motif (Pin et al., 2003) near the N-terminus.
1.3.4 Modulators of CaR Signaling
The CaR is able to detect changes in the extracellular environment through binding of its
large extracellular domain to the agonists, then transmit this signal to modulate
intracellular signaling events. Previous studies on the affinity of various putative agonists
of the CaR were performed using Xenopus laevis oocytes and HEK293 cells transfected
with recombinant CaR cDNA (Brown et al., 1993; Bai et al., 1995). Because there were
no assays for the binding of extracellular Ca
2+
to the CaR, the responses of agonist-
mediated CaR activation were measured in various ways: as altered Cl
-
inward currents in
oocytes, or as intracellular Ca
2+
mobilization, phosphatidylinositol (IP
3
) formation,
extracellular signal-regulated kinase (ERK) activation, or phospholipase A
2
(PLA
2
)
activation in the transfected HEK293 cells (Riccardi, 2002).
Although extracellular Ca
2+
is the primary physiological agonist, the CaR is also
responsive to a wide range of endogenous and exogenous agonists, including divalent and
trivalent cations, L-amino acids, polyamines, polypeptides, aminoglycoside antibiotics,
and pharmacological agents (Figure 1.1) (Riccardi, 2002). These agonists are divided into
two types, depending on their mechanism of inducing CaR activity: type I and type II
29
agonists. Organic and inorganic polycations are type I agonists of the CaR. These
polycations bind to the extracellular domain of the CaR to directly induce downstream
intracellular signaling (Magno et al., 2011). The potency of known polycation agonists
were reported as follows: La
3+
> Gd
3+
> Be
2+
> Ca
2+
= Ba
2+
> Sr
2+
> Mg
2+
(Riccardi,
2002). In contrast, type II agonists are allosteric modulators that sensitize the CaR to type
I agonists by interacting with the transmembrane domain of the receptor (Nemeth et al.,
2004). Therefore, type II agonists are ineffective in the absence of type I agonists,
particularly extracellular Ca
2+
. Interactions between type II agonists and the
transmembrane domain of the CaR induce conformational changes in the receptor, which
presumably reduce the threshold for CaR activation by type I agonists, particularly Ca
2+
(Nemeth et al., 2004). Subsequently, PTH secretion is reduced in the absence of a change
in the level of extracellular Ca
2+
. The IC
50
for extracellular Ca
2+
-mediated activation of
CaR and inhibition of PTH secretion in the parathyroid gland is ~1.2mM (Brown and
MacLeod, 2001; Brown, 1981).
30
Figure 1.1 CaR signaling transduction. The CaR can be activated by a variety of type I and type
II agonists to activate downstream signaling cascades, one of which involves the release of Ca
2+
ions from the internal calcium store into the cytosol. CaR, calcium-sensing receptor; AC,
adenylate cyclase; cAMP, cyclic AMP; DAG, diacylglycerol; PLC, phospholipase C; PKC,
protein kinase C; G
αi
and G
αq
, α subunits of the i- and q-type heterotrimeric G proteins,
respectively; IP
3
, inositol-1,4,5-triphosphate; PI(4,5)P
2
, phosphatidylinositol-4,5-bisphosphate.
1.3.5 Intracellular CaR Signaling Pathways
The CaR is capable of modulating an extensive and complex array of intracellular
signaling pathways through at least two G proteins, G
αi
and G
αq
, by controlling the
production of various secondary messengers (Handlogten et al., 2001) and consequently
suppressing PTH secretion. Some of the major, better characterized CaR-mediated
31
signaling pathways include phospholipase signaling, Mitogen-Activated Protein Kinase
(MAPK) signaling, Akt signaling, Rho signaling, and inhibition of cyclic adenosine
monophosphate (cAMP) (Magno et al., 2011).
1.3.5.1 Phospholipase Signaling
Early studies on the bovine CaR reported the activation of phospholipase C (PLC)
pathway and the subsequent elevation in IP
3
levels upon stimulation of the CaR with
extracellular Ca
2+
(Brown et al., 1993), followed by the mobilization of intracellular Ca
2+
from the internal calcium stores (Figure 1.1) (Nemeth and Scarpa, 1986; Membreño et
al., 1989). In addition to PLC, phospholipase D (PLD) and PLA
2
were also documented
to be activated by the CaR in bovine parathyroid cells and in transfected HEK293 cells
(Kifor et al., 1997). It was demonstrated that the CaR activates PLD via a pathway that
involves Gα
12/13
, and that the CaR is capable of coupling to Gα
12/13
(Huang et al., 2003).
Furthermore, PLC and PLA
2
could be activated by the CaR via a pathway that involves
G
αq
, and that CaR-dependent stimulation of PLA
2
activity could be inhibited by chelation
of extracellular Ca
2+
, confirming the role of the CaR in regulating PLA
2
activity
(Handlogten et al., 2001).
1.3.5.2 MAPK Signaling
Previous studies depicted the role of the CaR in modulating MAPK signaling cascades
through the tyrosine kinase, Src, and the PLC pathway. Specifically, these studies
demonstrated that agonists of the CaR can stimulate Src kinase activity and induce the
32
signaling cascade leading to ERK activation (McNeil et al., 1998; Hobson et al., 2000;
Hobson et al., 2003), and that the CaR can also increase ERK activity by initiating the
PLC pathway and activate tyrosine kinases (Kifor et al., 2001). The CaR has also been
implicated in the activation of other subtypes of MAPL signaling cascades, including c-
Jun N-terminal kinase (JNK), p42/44 MAPK, and p38 MAPK, where agonists of the CaR
were reported to enhance cyclooxygenase-2 expression via JNK signaling (Ogata et al.,
2006) and increase phosphorylation of the p42/44 MAPK and p38 MAPK to stimulate
mitogenic responses (Yamaguchi et al., 2000).
1.3.5.3 Akt Signaling
Another major signaling pathway that was demonstrated to be activated by CaR signaling
is the Akt signaling pathway. Experiments using fetal rat calvarial cells and renal
proximal tubule-derived opossum kidney cells exposed to elevated extracellular Ca
2+
or
other agonists found an increase in the phosphorylation and activation of the anti-
apoptotic protein kinase, Akt (Dvorak et al., 2004; Ward et al., 2005), while inhibition
experiments confirmed that CaR-mediated phosphorylation of Akt was via
phosphatidylinositol-3 kinase (PI3K) signaling (Ward et al., 2005). Results from these
studies suggest that the CaR is important for cell survival and proliferation via Akt
signaling.
33
1.3.5.4 Rho Signaling
The CaR also plays a role in activating PLD signaling via the cytoskeletal associated Rho
protein in Madin-Darby canine kidney cells by coupling to Gα
12/13
, where extracellular
Ca
2+
stimulation of the cells led to an increase in the membrane association of Rho, and
Clostridium botulinum C3 (C3) exoenzyme treatment, which inhibits Rho by adenosine
diphosphate (ADP) ribosylation, led to repression of CaR-stimulated PLD activity
(Huang et al., 2004). Other studies also reported the involvement of Rho in CaR
activation of serum response element (SRE), as evidenced by the abolition of CaR-
mediated SRE activity in HEK293-CaR cells when Rho was inhibited by C3 exoenzyme
treatment (Pi et al., 2002). The role of the CaR in Rho signaling was further examined by
transfecting HEK293-CaR cells with a constitutively active mutant of Rho, where
activated Rho induced intracellular Ca
2+
oscillations (Rey et al., 2005).
1.3.5.5 Inhibition of cAMP
Previous studies depicted the reductions in cAMP levels upon stimulation of HEK293-
CaR cells with the cations, Ca
2+
, Mg
2+
, and Gd
3+
, or the positive allosteric modulators,
NPS R-467 and γ-glutamyl peptides (Chang et al., 1998; Broadhead et al., 2011).
Furthermore, it was shown that the antidiuretic hormone-stimulated accumulation of
cAMP in thick ascending limb from rat kidneys showed the involvement of the CaR in
inhibiting this effect (De Jesus Ferreira and Bailly, 1998). The inhibition of cAMP levels
by extracellular Ca
2+
was shown to be coupled to PLC activation, further demonstrating
that the regulation of cAMP levels is highly sensitive to variations in extracellular Ca
2+
34
that are physiologically relevant for allowing the modulation of cAMP-mediated cellular
processes (de Jesus Ferreira et al., 1998).
1.3.6 Regulation of Cellular Functions by the CaR
The CaR is regarded as the major central mediator of systemic calcium homeostasis that
enables organisms to adapt to wide variations in extracellular Ca
2+
concentrations by
modulating the functions of the kidney, intestine, and bone to maintain a near constancy
of serum Ca
2+
levels (Brown, 2000). For example, in response to low extracellular Ca
2+
concentrations, the CaR increases PTH secretion from the parathyroid gland and
decreases the secretion of calcitonin from the thyroid to promote bone resorption and
calcium retention in the kidney (Brown, 2000). Besides the parathyroid gland and the
thyroid, the CaR is also expressed in a variety of tissues, including the gastrointestinal
tract (Cheng et al., 1999), kidney (Riccardi et al., 1995), heart (Wang et al., 2003), (skin
(Bikle et al., 1996), brain (Chattopadhyay et al., 1997), breast (Cheng et al., 1998), and
bone (Chang et al., 1999).
Despite its primary physiological role in systemic calcium homeostasis, the CaR is also
implicated in a multitude of other cellular processes. The CaR is expressed on osteoclasts
and was shown to be involved in regulating bone resorption, where stimulation of the
receptor resulted in the inhibition of osteoclastic bone resorption (1998). CaR activity
was also demonstrated to stimulate both differentiation and apoptosis in osteoclasts
through a signaling pathway that involves activation of PLC and nuclear factor kappa-
35
light-chain-enhancer of activated B cells (NF-κB) (Mentaverri et al., 2006). Using
knockdown technology, the CaR was also discovered to be important in the cell survival
of human keratinocytes, where decrease in CaR expression correlated with an increase in
apoptosis (Tu et al., 2008). It was also shown that a decrease in CaR expression inhibited
the extracellular Ca
2+
-induced expression of terminal differentiation genes in human
keratinocytes (Tu et al., 2008). The tissue-specific role of the CaR in the regulation of
cell proliferation has also been identified in previous studies. In osteoblasts (Yamaguchi
et al., 1998) and gastric mucous epithelial cells (Rutten et al., 1999), an increase in CaR
activity resulted in increased cell proliferation. However, in intestinal cells (Kállay et al.,
1997) and parathyroid cells (Colloton et al., 2005), an increase in CaR activity resulted in
decreased cell proliferation. Clinical evidence has also depicted that the CaR negatively
regulates the proliferation of parathyroid cells based on the presence of greater
parathyroid hyperplasia in the parathyroid glands associated with decreased CaR function
(Farnebo et al., 1997).
In vivo evidence of the functional relevance of the CaR has been reported using
generalized and tissue specific deletion of the receptor in the parathyroid gland, bone, and
cartilage. Germline-derived CaR
-/-
mice have high mortality, with very few surviving
beyond 7 days of age (Garner et al., 2001). Analysis of the skeleton in CaR
-/-
mice
revealed rickets, a disorder of impaired ECM mineralization of the growth plate and
newly formed trabecular and cortical bone, as the predominant skeletal abnormality
(Garner et al., 2001). As expected from the loss of parathyroid gland CaR, CaR
-/-
mice
36
have an extremely high circulating concentration of PTH, approximately 90-fold higher
compared to WT controls (Garner et al., 2001). Tissue-specific deletion of the CaR in the
parathyroid gland or bone resulted in a profound lack of skeletal development, while CaR
deletion in chondrocytes resulted in death before E13 due to a lack of postnatal growth
(Chang et al., 2008). Interestingly, transgenic mice with a constitutively active mutant
CaR in osteoblasts have reduced bone volume and density accompanied by a diminished
trabecular network (Dvorak et al., 2007). In addition, these mice have an increased
number and activity of osteoclasts due to the constitutively active CaR signaling in
mature osteoblasts, which resulted in increased expression of receptor activator of nuclear
factor-κB ligand (RANK-L), the major stimulator of osteoclast differentiation and
activation (Dvorak et al., 2007). Altogether, these studies support the critical role of the
CaR in early skeletal development, bone formation, and embryogenesis.
1.3.7 Regulation of the Localization of HSCs and Monocytes by the CaR
A distinctive characteristic of the bone is its mineral content. Classical physiological
experiments have depicted that although calcium concentration is maintained rigidly in
serum at 1.1-1.3mM, the concentration reaches 40mM near bone resorbing osteoclasts at
the surface of remodeling bone (Silver et al., 1988). Because of this distinctive
characteristic at the surface of the bone, it is interesting to investigate whether the
preferential localization of HSC in the endosteal region of the BM is partly dictated by
this feature. In fact, a previous study has shown the expression of the CaR on HSCs
(Adams et al., 2006). In this study, CaR was clearly demonstrated to play a key role in
37
HSC lodgment and engraftment in the BM, as HSCs deficient in CaR were found to have
lost their ability to lodge at the endosteal surface of the bone, leading to defective
engraftment (Adams et al., 2006). Interestingly, these HSCs were found to be normal in
migration and homing to the BM (Adams et al., 2006), suggesting that the defect was
solely affecting their ability to lodge in the BM. Moreover, this defect was correlated
with a diminished adhesive interaction with collagen I (Adams et al., 2006), indicating
that the altered localization of HSCs deficient in CaR might be due to their reduction in
the ability to adhere to collagen I in the BM microenvironment.
Besides the regulation of HSC localization in the BM, the CaR has also been implicated
in the regulation of monocyte localization in response to injury or inflammation.
Extracellular fluids at sites at sites of injury or infection are known to contain high
calcium concentrations (Menkin, 1960; Kaslick et al., 1970). Because of this
characteristic, it was hypothesized that the recruitment of immune cells such as mature
monocytes or macrophages to sites of injury or infection might be mediated through the
CaR (Olszak et al., 2000). Through a series of in vitro migration assays, it was
demonstrated that while monocytes derived from WT mice migrated in response to
extracellular calcium in a dose-dependent manner, monocytes derived from CaR
-/-
mice
lacked this chemotactic response (Olszak et al., 2000). Interestingly, when calcium was
subcutaneously injected into mice, there was a significant infiltration of monocytes and
macrophages at the skin (Olszak et al., 2000), providing in vivo evidence for the role of
38
the CaR in the localization of monocytes and macrophages at sites of injury or
inflammation where calcium concentrations are high.
1.3.8 The Differential Roles of the CaR in Bone Metastatic Cancers
Defining the functional characteristics of the CaR can be useful not only for
understanding the biological functions of the CaR in normal physiology, but also for
developing therapeutic strategies for the treatment of pathophysiological conditions
including cancer. In fact, the CaR has been reported to be expressed in a variety of benign
tumors and malignancies at expression levels that differ from those in their healthy
counterparts. These cancers include breast cancer (Cheng et al., 1998; Sanders et al.,
2000), prostate cancer (Sanders et al., 2001), and cancers originating from organs
involved in extracellular Ca
2+
homeostasis, such as colorectal cancer (Sheinin et al.,
2000) and parathyroid adenomas and carcinomas (Gogusev et al., 1997; Kifor et al.,
1996; Farnebo et al., 1997; Haven et al., 2004).
Interestingly, breast and prostate cancers both display preferential bone metastasis,
indicating that the bone provides a favorable microenvironment for its localization.
Containing a large repository of immobilized growth factors that can be released and
activated, the bone is indeed a fertile ground for cancer cell growth (Roodman, 2004).
Elevated levels of CaR mRNA and protein were depicted in highly metastatic breast
cancer cells in a clinical study (Mihai et al., 2006), suggesting an oncogenic property of
the CaR in breast cancer. Similarly, skeletal metastatic prostate cancer cell lines
39
displayed enhanced proliferation in elevated levels of extracellular Ca
2+
, which was
found to be associated with higher CaR expression (Liao et al., 2006). In addition, CaR
knockdown by RNA interference reduced in vitro cell proliferation and in vivo metastatic
progression, indicating a requirement of the CaR for facilitating these effects (Liao et al.,
2006).
In contrast, the CaR was reported to possibly serve as a tumor suppressor gene in
parathyroid and colorectal cancers. Although malignant transformation of parathyroid
tumors is rare, a significant correlation was found between CaR down-regulation and
primary parathyroid carcinomas with a high proliferative index (Haven et al., 2004). In
addition, decreased CaR expression was identified to be associated with parathyroid
hyperplasia in CaR knockout mice and also in patients with inactivating mutations of the
CaR (Ho et al., 1995; Yano et al., 2000; Brown et al., 1999), suggesting an inhibitory
effect of the CaR on parathyroid cellular proliferation. Similarly, CaR expression was
shown to be reduced and associated with a greater progression of malignancy in
colorectal cancer (Sheinin et al., 2000; Chakrabarty et al., 2003; Chakrabarty et al.,
2005). Furthermore, the inhibitory role of CaR in colorectal cancer was depicted in a
previous study using siRNA to down-regulate the CaR, where siRNA-transfected cells
did not exhibit reduction in cell growth upon addition of extracellular Ca
2+
(Bhagavathula
et al., 2007).
40
1.4 The Human Hematopoietic System
1.4.1 Identification of Human HSCs
The human hematopoietic system comprises a hierarchy of progenitor cells and mature
lineage cells that have differential capacities to proliferate and differentiate. At the top of
the hierarchy is the most primitive hematopoietic cell that is equipped with an intrinsic
capacity to self-renew and repopulate all the blood cell lineages (Figure 1.2).
Figure 1.2 Hierarchy of the hematopoietic system. Cell surface markers for identifying and
isolating mouse and human stem and progenitor cells are shown. HSC, hematopoietic stem cell;
MPP, multi-potent progenitors; CMP, common myeloid progenitor; CLP, common lymphoid
progenitor; MEP, megakaryocyte/erythroid progenitor; GMP, granulocyte/macrophage
progenitor; MkP, megakaryocyte progenitor; ErP, erythroid progenitor; Pro-DC, dendritic cell
progenitor; Pro-NK, natural killer cell progenitor; Pro-T, T cell progenitor; Pro-B, B cell
progenitor.
41
Therefore, the true functional measure that defines a HSC resides in its capacity to
engraft myeloablated recipients, repopulate their hematopoietic system, and sustain long-
term multilineage hematopoiesis in vivo. Over the last three decades, there have been
ongoing efforts to develop animal recipients for human cell transplantation for the
purpose of studying human diseases identifying functional human stem cells. In the study
of human hematopoiesis, xenogeneic model systems are widely utilized. The first
documentation of human stem cell engraftment in a xenogeneic model was performed in
severe combined immunodeficiency (SCID) mice, in which human hematopoietic organs,
such as the FL, fetal thymus, and fetal lymph node, were engrafted under the renal
capsule (McCune et al., 1988). This xenogeneic model, called the SCID-hu mouse
model, was demonstrated to be able to maintain long-term multilineage human
hematopoietic activity (Namikawa et al., 1990). SCID-hu mice were also generated by
subcutaneously implanting human fetal bone fragments (Chen et al., 1994). These
implanted human fetal bone fragments were shown to be able to support the maintenance
of a cell population that has multilineage repopulation potential (Chen et al., 1994). This
suggests the fetal bone implants in SCID mice are able to provide a functional human
hematopoietic microenvironment to support hematopoiesis. The study of human
hematopoiesis was also performed in pre-immune fetal sheep, where human BM or FL
were transferred in utero to the sheep, and long-term multilineage contribution of the
human donor cells were monitored (Zanjani et al., 1991; Zanjani et al., 1994; Srour et al.,
1992). However, because of the costs required in maintaining large animals for long term
studies, the use of immunodeficient mice as recipients for human stem cell graft is the
42
most common and cost-effective strategy available. Since the development of the SCID-
hu mouse model, other improved xenogeneic models, such as the widely used non-obese
diabetic SCID (NOD/SCID) and the most recent NOD/SCID/IL-2Rg
c
-null (NSG) mice,
have been developed and utilized for the study of human hematopoiesis. The NSG mouse
model was demonstrated to be superior in the engraftment of human hematopoietic cells
compared to other immunodeficient mouse models (Ito et al., 2002).
1.4.2 Isolation of Human HSCs
With advances in flow cytometry, isolation of highly enriched HSC populations was
made possible and became an essential process for the study of human hematopoiesis.
The first step towards the isolation of human HSCs was the creation of monoclonal
antibodies against the antigen, My-10, or CD34, a single chain transmembrane molecule
present on the surface of BM cells (Civin et al., 1984; Civin et al., 1987). The CD34
glycoprotein is a hematopoietic cell surface antigen that constitutes about 1-4% of normal
adult BM leukocytes (Strauss et al., 1986). Later, the CD34
+
population was shown to be
enriched for primitive human hematopoietic progenitors in adult and fetal BM using the
in vitro colony forming cell (CFC) assay (Andrews et al., 1986; Andrews et al., 1989;
Strauss et al., 1986), and also demonstrated to be able to provide in vivo long-term
repopulation activity in SCID-hu mice (DiGiusto et al., 1994). In combination with
CD34, other cell surface markers have been used to isolate more enriched HSC
populations from different human hematopoietic tissues. For example, hematopoietic
cells expressing CD34 and Thy-1 (CD90), a phosphatidylinositol-anchored cell surface
43
molecule, were shown to highly enrich for HSCs from fetal and adult BM, FL, umbilical
cord blood (UCB), and mobilized peripheral blood (MPB) (Baum et al., 1992; Craig et
al., 1993; Murray et al., 1995a; Murray et al., 1995b; Mayani and Lansdorp, 1994) with
high clonogenic activity and engraftment potential. Primitive human hematopoietic cells
were also shown to express low levels of the c-Kit antigen as demonstrated by the LTC-
IC assay (Gunji et al., 1993). In addition, HSCs were shown to express high levels of
CD59, but have low or undetectable expression for CD38, CD71, CD45RA, or lineage
markers (Terstappen et al., 1991; Murray et al., 1995b; Hill et al., 1996; Lansdrop and
Dragowska, 1992; Lansdorp et al., 1990; Chen et al., 1997). Rhodamine123 (Rh123) was
also proven to be useful for the isolation of human HSCs, where the CD34
+
Thy-1
+
Lin
-
Rh123
lo
subset from human fetal BM was characterized to be highly enriched for
cobblestone area forming cell (CAFC) activity (Baum et al., 1992). Interestingly, the use
of human leukocyte antigen (HLA)-DR expression to identify human HSCs relies on the
tissue source. It was reported that human HSCs obtained from adult BM are CD34
+
HLA-
DR
-
(Srour et al., 1991), whereas human HSCs obtained from fetal BM and cord blood
are CD34
+
HLA-DR
+
(Huang and Terstappen, 1994; Traycoff et al., 1994).
1.4.3 Sources of Human HSCs
Currently, sources of human HSCs include BM, UCB, and MPB. In the clinical
transplantation setting, decisions regarding HSC source for the treatment of
hematological disorders are based upon guidelines that include individual patient needs,
urgency of the transplant, and size of the patients for determining the adequacy of cell
44
dose (Smith and Wagner, 2009). In addition, many of the functional properties of various
human HSC sources need to be considered as well. With the advent of cord blood
banking, the use of UCB for transplantation is becoming increasingly popular because it
circumvents the issue of donor availability and HLA-restrictions that are associated with
BM or MPB transplants. However, UCB transplantation is not particularly suitable for
treatment in adult patients due to cell dose limitations (Smith and Wagner, 2009). This
problem appears to be linked to delayed hematopoietic recovery after UCB
transplantation (Gluckman et al., 1997; Rubinstein et al., 1998; Wagner et al., 2002;
Locatelli et al., 2003; Laughlin et al., 2004; Rocha et al., 2004). On the other hand, when
compared to BM grafts, MPB stem cells are associated with shortened period required for
hematopoietic recovery (Bensinger et al., 1995; Körbling et al., 1995).
1.4.4 Regulation of Human HSC Functions
The ability to maintain and expand functional human HSCs in culture is a long-standing
challenge. Although ex vivo culture of human hematopoietic cells on stroma is
established, and cytokines can be used to stimulate the proliferation of human HSCs, the
in vivo repopulating stem cell function is lost. This was demonstrated to be associated
with a decline in the number of repopulating cells by studies on human cord blood and
mobilized peripheral blood in immunodeficient mouse models (Gan et al., 1997). The
impairment in the ability of ex vivo cultured human HSCs to repopulate the BM of
immunodeficient mouse recipients was explained by the loss of quiescence as the cells
transition from G
0
to G
1
phase of the cell cycle (Gothot et al., 1998). In terms of homing
45
of human HSCs to the BM, the most well studied regulator is the CXCR4/SDF-1 axis.
Using human cord blood HSCs, it was demonstrated that CD34
+
CD38
-/low
CXCR4
+
cells
efficiently and rapidly home to the BM and spleen of NOD/SCID mice and
NOD/SCID/B2m
null
mice, and that homing of enriched human CD34
+
cells could be
inhibited by pre-treatment with anti-CXCR4 antibodies (Kollet et al., 2001). Human
CD34
+
cells express the chemokine receptor CXCR4, which is activated by SDF-1 to
regulate cellular interactions and engraftment in the BM microenvironment (Peled et al.,
1999). When the human CD34
+
cells are activated by SDF-1, major integrins such as
VLA-4, VLA-5, and lymphocyte function associated antigen-1 (LFA-1) are activated to
support firm adhesion to fibronectin and promote transendothelial migration (Peled et al.,
2000). Using the sheep model of in utero hematopoietic stem cell transplantation, the
importance of VLA-4 in regulating homing of transplanted human CD34
+
cells was
exemplified, where pre-treatment of cells with an anti-VLA-4 monoclonal antibody
decreased BM homing in vivo (Zanjani et al., 1999). Recent studies have shown the role
of tetraspanins, a large family of evolutionarily conserved 4-transmembrane domain
proteins important for facilitating the assembly of specialized molecular aggregates on
plasma and intracellular membranes, in mediating human HSC homing in
immunodeficient mice (Larochelle et al., 2011; Leung et al., 2011). Specifically, using
neutralizing antibodies, these studies demonstrated that CD82 (Larochelle et al., 2011)
and CD9 (Leung et al., 2011) positively regulate the homing of transplanted CD34
+
cells
to the BM of immunodeficient mice.
46
1.5 Multiple Myeloma
MM is a devastating disease that occurs in the bone and represents the second most
commonly diagnosed hematological malignancy, with an annual incidence of
approximately 15,000 cases in the United States alone (Singhal and Mehta, 2006). It is
characterized by osteolytic bone lesions, hypercalcemia, and plasmacytosis, or the
aberrant monoclonal outgrowth of plasma cells or terminally differentiated B cells in the
BM. It is a devastating bone disease that encompasses a multitude of clinical
complications including severe bone pain and fractures, impaired humoral immunity,
susceptibility to infection, BM defects due to the presence of neoplastic marrow cells,
and renal failure (Mundy, 1998; Oyajobi, 2007). Interestingly, MM is more prevalent in
the senior population (median age at diagnosis is 67 years of age) (Singhal and Mehta,
2006).
1.5.1 The BM Microenvironment
The mechanism behind bone metastasis was first introduced by Paget in 1889 as the
“seed-and-soil hypothesis” (Paget, 1889), where the seed is the cancer cell and the soil is
the BM microenvironment that provides a fertile ground for the cancer cell to grow and
develop. It is becoming increasingly recognized that the pathogenesis of MM involves
complex changes in both the myeloma cells and their surrounding bone marrow
microenvironment. Therefore, tremendous efforts have been made to understand and
identify the underlying pathophysiologic phenomena of MM, especially the interplay
between the myeloma cells and the supportive BM microenvironment. Several factors
47
account for the importance of the BM microenvironment in the support of the growth,
survival, and homing of myeloma cells.
1.5.1.1 Bone as a Repository for Growth Factors
The bone is a large repository for immobilized growth factors, including transforming
growth factor β, insulin-like growth factors I and II, bone morphogenetic proteins, and
calcium (Hauschka et al., 1986). These immobilized growth factors are released and
activated during bone resorption (Pfeilschifter and Mundy, 1987). In fact, bone resorption
is enhanced in MM due to stimulation of osteoclastogenesis, which is mediated by
interactions between nuclear factor-κB (RANK) on the surface of osteoclastic lineage
cells and RANK-L produced by BM stromal cells and osteoblasts (Abe, 2011; Roodman,
2004). In addition, decrease in osteoprotegerin, a decoy receptor for RANK-L in BM
stromal cells that acts to inhibit RANK signaling, further promotes the stimulation of
osteoclastogenesis in MM (Abe, 2011). These coordinated processes result in osteolytic
bone lesions, and there is evidence to suggest that osteolytic bone lesions are mediated by
osteoclasts rather than tumor cells in bone metastasis (Taube et al., 1994). Besides
RANK-L, myeloma cells can stimulate the release of an osteoclastogenic C-C chemokine
called macrophage inflammatory protein (MIP), specifically MIP-1α and MIP-1β. It was
documented in previous studies that the concentrations of MIP-1α in the BM plasma were
higher in patients with advanced stages of MM (Choi et al., 2000), and that most primary
MM cells from patients with multiple osteolytic lesions secreted MIP-1β that mediate
osteoclastogenic activity in vitro (Abe et al., 2002). Furthermore, upregulation of other
48
osteoclastogenic factors including interleukin (IL)-6, tumor necrosis factor (TNF) α,
PTH-rP, vascular endothelial growth factor (VEGF), and SDF-1α has also been reported
(Zannettino et al., 2005; Abe, 2011; Mitsiades et al., 2004), some of which are also
known to be autocrine/paracrine survival factors for the MM cells (Singhal and Mehta,
2006). The homing of MM cells to the BM is mediated by SDF-1α, which interacts with
its receptor CXCR4 on MM cells to induce motility, internalization of CXCR4, and
cytoskeletal rearrangement (Hideshima et al., 2002; Hideshima et al., 2007). Studies
using specific anti-CXCR4 antibodies and CXCR4 inhibitors have demonstrated the
inhibition of MM cell migration in vitro (Alsayed et al., 2002), suggesting the importance
of SDF-1α and CXCR4 interaction in regulating the homing of MM cells in the BM
microenvironment.
1.5.1.2 Cell-Cell and Cell-ECM Interactions
The BM microenvironment comprises various stromal cell components and ECM
molecules that can interact with MM cells to activate a pleiotropic cascade of
proliferative or anti-apoptotic signaling pathways, including PI3K/Akt/mTOR/p70S6K,
IKK-α/NF-κB, Ras/Raf/MAPK, and JAK/STAT3 (Mitsiades et al., 2004). Other
proliferative or anti-apoptotic effectors include upregulation of cyclin D, caspase
inhibitors, anti-apoptotic Bcl-2 family members, and increased activity of the proteasome
(Mitsiades et al., 2004). These molecular events are triggered either directly via cell-
ECM interactions or cell-cell interactions between myeloma cells and BM stromal cells,
vascular endothelial cells, osteoclasts, or other cellular components in the BM
49
microenvironment; or indirectly, via cytokines or growth factors released by BM stromal
cells or MM cells (Mitsiades et al., 2004). MM cells can also interact with osteoblasts to
suppress bone formation. Specifically, MM cells inhibit osteoblastic differentiation
through the secretion of Dickkopf-1 (DKK-1), an inhibitor of the Wnt canonical signaling
pathway (Tian et al., 2003). In addition, previous studies have documented the
constitutive expression of very late antigen VLA-4 and its corresponding ligand, vascular
cell adhesion molecule (VCAM)-1 on MM cells and BM cells, respectively (Drew et al.,
1996; Sanz-Rodríguez et al., 1999; Luque et al., 1998; Barker et al., 1992). Moreover, it
was depicted that cell-cell adhesion mediated by VLA-4 and VCAM-1 enhances MM-
induced osteoclastogenesis, and that the activity was not blocked by neutralizing
antibodies to known osteoclastogenic cytokines such as IL-1, IL-6, TNF, or PTHr-P
(Michigami et al., 2000). However, stimulation of MM cell growth by VLA-4 and
VCAM-1 interactions was inhibited by a monoclonal antibody to α
4
integrin, further
confirming the supporting role of BM stromal cells in MM cell growth and suggesting the
preferential localization of MM cells in the BM (Mori et al., 2004). In addition to cell
growth and survival, interactions between MM cells and the BM microenvironment also
contribute to resistance to chemotherapy. Myeloma cells that overexpress VLA-4 was
shown to have increased cell-ECM adhesion to fibronectin and enhanced drug resistance
to doxorubicin and melphalan (Damiano et al., 1999), supporting the concept of cell
adhesion-mediated drug resistance. Based on this concept, it is anticipated that
developing chemotherapeutics targeting CAMs on MM cells can be an effective strategy
for inhibiting the pathophysiology of drug resistant MM cells.
50
1.5.1.3 Angiogenic Interactions
Similar to osteoclastogenesis, angiogenesis is enhanced in the BM in MM. In fact, MM
was the first hematological malignancy in which a significant correlation of angiogenesis
with prognosis and survival was shown (Jakob et al., 2006), where angiogenesis is
enhanced in the BM in parallel with tumor progression (Kumar et al., 2004; Bhatti et al.,
2006). This was depicted by Vacca et al., who reported that serum-free conditioned
media from the BM plasma cells of patients with active MM had a significantly higher in
vitro and in vivo angiogenic activity than samples from patients with non-active MM or
monoclonal gammopathy of undetermined significance (MGUS) (Vacca et al., 1999). In
fact, the progression of MM can be illustrated as a vicious cycle that involves bone
destruction mediated by an overly active population of osteoclasts that can secrete pro-
angiogenic factor to enhance angiogenesis. First of all, MM cells and BM stromal cells
secrete angiogenic factors including VEGF and basic fibroblast growth factor (bFGF) to
enhance angiogenesis (Jakob et al., 2006; Corre et al., 2007). Osteoclasts constitutively
secrete a large amount of the pro-angiogenic factor osteopontin, a ligand for α
V
β
3
integrin
expressed on vascular endothelial cell surface, which cooperates with VEGF secreted
from MM cells to enhance angiogenesis (Tanaka et al., 2007). When vascular endothelial
cells are stimulated with VEGF and OPN, IL-8 production is induced to stimulate
osteoclastogenesis (Tanaka et al., 2007). Osteoclasts also produce matrix
metalloproteinase (MMP)-9, which has been demonstrated to be responsible for
osteoclast-induced angiogenesis (Cackowski et al., 2010). Taken together, these parallel
51
osteoclastogenic and angiogenic processes form a vicious cycle to drive the progression
of MM.
1.5.2 Localization of MM cells in the BM
MM cells are known to be closely interacting with osteoclasts in the BM to propagate the
pathophysiology of the disease, including the formation of osteolytic bone lesions and
hypercalcemia. Osteoclasts accumulate only on bone-resorbing surfaces adjacent to
myeloma cells, and the numbers do not increase in bones not involved in tumor
progression (Roodman, 2001). Based on this observation, it is likely that myeloma cells
are found to be localized in the endosteal region of the BM, next to the bone-resorbing
osteoclasts. A recent study using the U-266 human MM cell line injected into NSG mice
identified the specific spatial localization of myeloma cells in the BM. In this study,
engraftment of the injected GFP
+
U-266 cells were detected in the BM, but not in the
spleen or liver, and that the cells were primarily observed at the metaphyseal region of
the BM endosteum (Iriuchishima et al., 2012). Further immunohistochemical analysis
revealed that the GFP
+
U-266 cells formed a complex with osteoblasts and osteoclasts in
the BM endosteum (Iriuchishima et al., 2012), suggesting that the myeloma niche is
composed of osteoblasts and osteoclasts. In addition, there was an induction of
angiogenesis in the BM by the U-266 cells, which is in accordance with the current
observation that there is a close relationship between the progression of MM, osteoclast
formation, and angiogenesis.
52
1.5.3 Evolution of Therapeutic Targets
Traditionally, MM patients were treated with the alkylating agents, melphalan and
cyclophosphamide, and corticosteroids, and the median survival was about 3 years
(Singhal and Mehta, 2006). Recent advances in chemotherapy have brought forth three
new therapeutic agents with significant anti-myeloma activity, including the proteasome
inhibitor bortezomib and the immunomodulatory drugs such as thalidomide and
lenalidomide (Singhal and Mehta, 2006; Anderson, 2011). These new therapeutic agents
target and disrupt the MM cells in the BM microenvironment to improve the efficacy of
the treatment against tumor progression through processes including induction of
apoptosis, inhibition of DNA damage repair, down-regulation of adhesion molecules on
MM cells, and stimulation of anti-angiogenic factors (Anderson, 2011). Although MM
remains largely incurable, the development of new chemotherapeutic agents and the
application of high-dose chemotherapy and stem cell transplantation in the past decade,
the median patient survival has doubled from 3-4 to 7-8 years (Kumar et al., 2008;
Anderson, 2011). Because of the heterogeneity of MM, oncogenomic studies for
characterizing the underlying molecular pathogenesis, coupled with models of MM cells
in the BM microenvironment to define the role of cellular interactions in conferring
tumor cell growth, survival, and drug resistance, are becoming increasingly useful for
identifying and developing immune-based therapies including monoclonal antibodies,
immunotoxins targeting MM cells and cytokines, and novel vaccine strategies (Anderson,
2011).
53
Chapter 2: Pharmacologic Modulation of the Calcium-Sensing
Receptor Enhances Hematopoietic Stem Cell Lodgment in the
Adult Bone Marrow
2.1 Abstract
The ability of HSCs to undergo self-renewal is partly regulated by external signals
originating from the stem cell niche. Our previous studies using HSCs obtained from the
FL of mice deficient for the CaR have shown the crucial role of this receptor in HSC
lodgment and engraftment in the BM endosteal niche. Using a CaR agonist, Cinacalcet,
we assessed the effects of stimulating the CaR on the function of murine HSCs. Our
results demonstrate that CaR stimulation increases primitive hematopoietic cell activity in
vitro, including growth in stromal cell co-cultures, adhesion to ECM such as collagen I
and fibronectin, and migration towards the chemotactic stimulus, SDF-1α. Receptor
stimulation also led to augmented in vivo homing, CXCR4-mediated lodgment at the
endosteal niche, and engraftment capabilities. These mechanisms by which stimulating
the CaR dictates preferential localization of HSCs in the BM endosteal niche provide
additional insights into the fundamental interrelationship between the stem cell and its
niche. These studies also have implications in the area of clinical stem cell
transplantation, where ex vivo modulation of the CaR may be envisioned as a strategy to
enhance HSC engraftment in the BM.
54
2.2 Introduction
In the adult hematopoietic system, HSCs reside in a specific anatomic location in the BM
known as the stem cell niche (Schofield, 1978). The signaling cues originating from the
stem cell niche serve as instructions for the HSCs to undergo either self-renewal or
differentiation for the maintenance of the hematopoietic system in the individual. In the
clinical transplantation setting, interactions between the transplanted HSCs and the stem
cell niche are essential in determining the clinical outcome of the transplantation (Ballen,
2007). In order for successful engraftment and re-establishment of hematopoiesis to occur
in the recipient, the transplanted HSCs must first migrate from the peripheral circulation
to the BM through transendothelial migration, a process known as homing (Lapidot et al.,
2005). Homing is a fairly rapid process that occurs in within the first 24 to 48 hours
following injection of the cells and results in transient retention of the hematopoietic cells
(Lapidot et al., 2005). Following homing is a process called lodgment when
hematopoietic cells localize to a specific BM microenvironment called the stem cell niche
through mechanisms involving activation of adhesive interactions (Wolf, 1974; Lapidot
et al., 2005). However, although mature, specialized hematopoietic cells can home to the
BM, only HSCs homed to the stem cell niche can initiate long-term repopulation. Finally,
HSCs proliferate in the stem cell niche to repopulate the hematopoietic system of the pre-
conditioned recipient in a process called engraftment. This process can be further divided
into short-term and long-term engraftment. Short-term engraftment ranges from weeks to
a few months and is initiated by differentiating progenitors, whereas long-term
55
engraftment ranges from many months to years and is mediated by HSCs that have
successfully homed and lodged in the stem cell niche (Nilsson and Simmons, 2004).
The process of HSC lodgment following transplantation is believed to be regulated in
part by CAMs expressed on the cellular surface. Previous studies have demonstrated the
importance of cell surface molecules such as α
4
integrins (Priestley et al., 2006),
hyaluronic acid (Nilsson et al., 2003), or stem cell factor (Driessen et al., 2003) and
osteopontin (Grassinger et al., 2009)
in retaining HSCs in the BM endosteal region
through interactions with stem cell niche components. However, studies on the
identification of the molecules that dictate lodgment are complicated by the vast array of
CAMs that are expressed on HSCs and the broad range of potential ligands that are
expressed on the stromal cells with possible overlapping functions (Simmons et al, 1997).
Because the endosteal surface of the bone is known to have a high concentration of Ca
2+
ions that reaches as high as 40mM underneath resorbing osteoclasts (Silver et al., 1988),
we hypothesized that lodgment of HSC in the endosteal region of the BM is specifically
dictated by this unique feature. Our previous work has shown the role of the CaR, a G-
protein coupled receptor that plays a key role in the regulation of extracellular calcium
homeostasis (Hofer and Brown, 2003; House et al., 1997), in HSC lodgment and
engraftment in the BM, where HSCs deficient for the CaR lose their ability to lodge at
the endosteal surface of bone, leading to defective engraftment (Adams et al., 2006).
56
In this study we wished to examine the precise cellular and molecular mechanisms that
dictate CaR-mediated HSC lodgment in the adult BM stem cell niche. By stimulating the
activity of CaR using a CaR agonist, underlying mechanisms of HSC lodgment in the
BM endosteal region following transplantation can be elucidated, providing additional
insights for the fundamental interrelationship between the stem cell niche and stem cell
fate. Cinacalcet, a calcimimetic compound clinically approved to be used as a treatment
for secondary hyperparathyroidism, acts as a positive allosteric modulator of CaR to
increase the sensitivity of the receptor to activation by extracellular Ca
2+
ions. This allows
modulation of the functional CaR without altering the level of extracellular Ca
2+
concentration (Nemeth, 2004). Using in vitro and in vivo model systems, we assessed the
effects of Cinacalcet treatment on the function of primitive hematopoietic cells.
Specifically, we tested the hypothesis that ex vivo stimulation of the CaR on HSCs can
lead to enhanced BM homing, lodgment, and engraftment in vivo, thereby also providing
therapeutic implications pertinent to clinical stem cell transplantation.
57
2.3 Materials and Methods
2.3.1 Animals
Six- to eight-week-old male C57Bl/6 and B6.SJL mice (Taconic Farms Inc, Oxnard, CA,
USA) were obtained and used in accordance with the University of Southern California
Institutional Animal Care and Use Committee (IACUC) guidelines. Mice were housed in
sterilized microisolator cages and received autoclaved food and water ad libitum.
2.3.2 Euthanasia
Mice were sacrificed by placing them in a standard CO
2
chamber attached a pressurized
CO
2
tank and exposing them to the CO
2
gas for approximately 5 minutes to attain
complete asphyxia, narcosis, complete unconsciousness, and death. To ensure death of
the mice, cervical dislocation was used as the secondary method.
2.3.3 Bone Harvesting
Sacrificed mice were pinned down to the dissection board and doused in 70% ethanol.
All the surgical tools were sterilized using 70% ethanol. The fur and skin from each hind-
limb were removed by lifting the skin at the base with tweezers and cutting away the skin
across the thigh and down to the ankle. Muscles were removed from the entire limb so
that the bones are completely exposed. The entire limb was removed by cutting above the
hip joint. The cleaned bones were placed in minimum essential medium alpha (α-MEM)
supplemented with 10% fetal bovine serum (FBS) and 1X penicillin-streptomycin (P/S)
58
(all from Mediatech Inc., Manassas, VA, USA). The two ends of the bones were cut
using a scalpel (Becton Dickinson, Franklin Lakes, NJ, USA), and the BM was flushed
out using a 1ml syringe attached to a 25-gauge needle (Becton Dickinson). To remove
any remaining bone fragments or hair, the BM solution was filtered using a 70µm cell
strainer (Becton Dickinson).
2.3.4 Cinacalcet Preparation and Cell Treatment
Cinacalcet (30mg) (Amgen, Thousand Oaks, CA, USA) was dissolved in 95% ethanol
(EMD Chemicals, Inc., Gibbstown, NJ, USA), filtered through a 0.2µm Acrodisc
®
syringe filter (Pall Corporation, Port Washington, NY, USA), and diluted to 1mM stocks.
The stocks were stored at -20°C. Cells were treated with either 2.5µM Cinacalcet or
ethanol as the vehicle control in α-MEM supplemented with 10% FBS and P/S in a 37°C
water bath for 90 minutes, protected from light.
2.3.5 Colony Forming Unit-Culture (CFU-C) Assay
Bone marrow mononuclear cells (BM MNC) were obtained from the hind-limbs of
C57Bl/6 mice and treated with Cinacalcet or ethanol as the vehicle control as described
above. The treated cells were then resuspended in MethoCult
®
GF M3434 (STEMCELL
Technologies, Vancouver, BC, Canada) and cultured at 37°C in a humidified, 5% CO
2
atmosphere as followed. First, MethoCult
®
GF M3434 medium containing FBS, bovine
serum albumin (BSA), recombinant human (rh)-insulin, human transferrin (iron-
saturated), 2-mercaptoethanol (2βME), recombinant murine (rm)-SCF, rm IL-3, rh IL-6,
59
rh erythropoietin was defrosted at room temperature and vortexed well. The treated cells
were resuspended in 0.3ml of α-MEM supplemented with 10% FBS and P/S, then added
to the MethoCult
®
GF M3434 medium to yield a cell density of 20,000 cells/1.1ml in a
15ml tube (VWR International, Radnor, PA, USA). The tubes containing the cells were
then vortexed and let stand until all the bubbles had risen to the top surface
(approximately 5 minutes). A 3ml syringe attached to 18-gauge needle (Becton
Dickinson) was then used to plate out the cells in duplicates in a volume of 1.1ml per
well in a 6-well plate pre-marked with gridlines (VWR International). Distilled water was
added to the empty wells in the middle of the plate (Figure 2.1). The plates were wrapped
in a plastic bag and incubated at 37°C in a humidified, 5% CO
2
atmosphere for 7 days
before they were scored for the total number of hematopoietic colonies formed in each
well. The total number of CFU-Cs was determined as the sum of the number of colony
forming unit-macrophage (CFU-M), burst forming unit-erythroid (BFU-E), and colony
forming unit-granulocyte (CFU-G) colonies (Figure 2.1)
60
Figure 2.1 CFU-C assay. The plate layout is shown on the left. Representative pictures of
hematopoietic colonies observed on day 7 are shown on the right. The CFU-C assay measures
hematopoietic progenitor cell number or activity in vitro.
2.3.6 Fluorescence Activated Cell Sorting (FACS)
BM MNCs were stained in 200µl of 1X phosphate buffered saline (PBS; Mediatech Inc.)
with appropriate volumes of primary anti-mouse antibodies (Table 2.1) (all from Becton
Dickinson) for 15 minutes on ice, protected from light. Following primary antibody
incubation, cells were washed with 5ml of 1X PBS, centrifuged at 400g for 5 minutes,
and resuspended in 200µl of 1X PBS. Next, the resuspended cells were stained with 3µl
of PE-Cy5 streptavidin (Becton Dickinson) as the secondary antibody for 15 minutes on
ice, protected from light.
Control
Cinacalcet
Cinacalcet
Control Water
Water
CFU-M
CFU-G BFU-E
Day 7: CFU-C Scoring
Plate layout
61
Finally, the stained cells were resuspended in 0.5-1ml of 1X PBS, then Lin
-
c-Kit
+
Sca-
1
+
Flk-2
-
(LKS
+
F
-
) cells, Lin
-
c-Kit
+
Sca-1
+
Flk-2
+
(LKS
+
F
+
) cells, and Lin
-
c-Kit
+
Sca-1
-
Flk-
2
+
(LKS
-
F
+
) cells were sorted using the FACSAria or Aria II flow cytometer (Becton
Dickinson). The gating schemes are shown in Figure 2.2 below.
Finally, the stained cells were resuspended in 0.5-1ml of 1X PBS, then Lin
-
c-Kit
+
Sca-
1
+
Flk-2
-
(LKS
+
F
-
) cells, Lin
-
c-Kit
+
Sca-1
+
Flk-2
+
(LKS
+
F
+
) cells, and Lin
-
c-Kit
+
Sca-1
-
Flk-
Primary Antibody Antigen Recognized Clone Conjugate
B220
B-Cell Specific CD45R
Isoform
RA3-6B2
Biotin
(Biotinylated
Lineage Panel)
CD3e T-Cell Receptor (TCR) 145-2C11
CD11b
Macrophage-1 (Mac-1)
Antigen
M1/70
Gr-1
Myeloid Differentiation
Antigen
RB6-8C5
TER119 Ter-119 Antigen TER-119
CD117 (c-Kit) Stem Cell Factor 2B8
Allophycocyanin
[APC]
CD135 (Flk-2) Fetal Liver Kinase-2 A2F10.1 Phycoerythrin [PE]
Ly-6A/E (Sca-1) Stem Cell Antigen-1 E13-161.7
Fluorescein
isothiocyanate
[FITC]
Table 2.1 Antibody list. The primary antibodies used for FACS are listed.
62
2
+
(LKS
-
F
+
) cells were sorted using the FACSAria or Aria II flow cytometer (Becton
Dickinson). The gating schemes are shown in Figure 2.2 below.
Figure 2.2 Gating schemes for FACS. Long-term HSCs (LT-HSCs) are defined as LKS
+
F
-
,
short-term HSCs (ST-HSCs) are defined as LKS
+
F
+
, and multi-potent progenitors (MPPs) are
defined as LKS
-
F
+
.
2.3.7 Cobblestone Area Forming Cell (CAFC) Assay
The Lin
-
c-Kit
+
(LK) sub-population was sorted from BM MNCs as described above and
treated with 2.5µM Cinacalcet or ethanol as the vehicle control. Following treatment,
LKS
+
F
-
cells, LKS
+
F
+
cells, and LKS
-
F
+
cells were sorted and seeded in serial dilutions
onto a confluent layer of irradiated OP9 stromal cells in a cell culture treated 96-well
plate (Becton Dickinson) (Figure 2.3). The OP9 cells were cultured in α-MEM
supplemented with 10% FBS and P/S and irradiated at 35 Grays (Gy) 24 hours before use
for the CAFC assay. The irradiated OP9 cells were seeded at 20,000 cells/well in a 96-
well plate. The outer wells of the 96-well plate contained distilled water to prevent drying
of the cell cultures. The seeding procedure for the various hematopoietic progenitor and
stem cell populations was performed using the FACSAria or Aria II flow cytometer with
63
the appropriate plate adapter. The following cell numbers of each population were
seeded:
LKS
+
F
-
(LT-HSC): 3, 6, 12, 24, and 48
LKS
+
F
+
(ST-HSC): 6, 12, 24, 50, and 100
LKS
-
F
+
(MPP): 15, 30, 60, 120, and 180
The co-cultures were maintained in α-MEM supplemented with 10% FBS and P/S
(Mediatech Inc.) in a humidified atmosphere at 37°C in a humidified, 5% CO
2
atmosphere. Half of the culture medium was replaced with fresh medium every week.
The presence of CAFCs underneath the OP9 stromal layer was scored on week 5, and the
frequency of CAFCs was calculated using the L-Calc software (STEMCELL
Technologies).
64
Figure 2.3 CAFC assay. Schematics showing the seeding of hematopoietic progenitor and stem
cells onto an irradiated OP9 stromal layer. The presence of CAFCs underneath the OP9 stromal
layer, as shown in the circled area, is scored on week 5. The CAFC assay measures primitive
hematopoietic cell number or activity in vitro.
2.3.8 Calcium Flux Assay
LK cells stained with all lineage and stem cell antibodies were sorted from BM MNCs
and treated with Cinacalcet at the following dosages: 0.625µM, 1.25µM, and 2.5µM. The
treated cells were then incubated with 2µg/ml indo-1 (Molecular Probes, Carlsbad, CA,
USA) in 1ml of 1X PBS for 30 minutes in a 37°C water bath, protected from light.
Calcium flux was measured using the LSR II flow cytometer (Becton Dickinson)
equipped with a UV laser, a violet bandpass filter centered at 390±30nm, and a blue
bandpass filter centered at 530±30nm. The calcium flux response was determined by a
ratio of violet (short) to blue (long) wavelengths. The instrument setup and cellular
loading was checked by adding 2µg/ml of ionomycin (Invitrogen, Carlsbad, CA, USA) to
OP9 stromal
layer
35Gy
Radiation
MPPs, ST-HSCs,
LT-HSCs
(Control or Cinacalcet)
Week 5: CAFC
Scoring
65
indo-1-loaded cells suspended in Hank’s Balanced Salt Solution (HBSS) containing
calcium so that maximum flux response could be observed.
For the measurement of CaR stimulation by Cinacalcet treatment, the treated cells were
resuspended in approximately 300µl of Ca
2+
-free PBS. The baseline response is measured
and recorded for about 20 seconds, then 1.5mM of CaCl
2
(Sigma-Aldrich) was quickly
added to the cells to induce calcium flux. The response was measured for 5 to 10 minutes.
For the measurement of SDF-1α-induced calcium flux response, the treated cells were
resuspended in approximately 300µl of HBSS containing calcium (Invitrogen). The
baseline response is measured and recorded for about 20 seconds, then 100ng/ml of SDF-
1α (PeproTech Inc., Rocky Hill, NJ, USA) was quickly added as the stimulus. The
response was measured for about 5 minutes. In these experiments, the FlowJo software
(TreeStar, Stanford, CA, USA) was used to analyze the calcium flux response in the
LKS
+
F
-
, LKS
+
F
+
, and LKS
-
F
+
sub-populations following treatment.
2.3.9 Chemotaxis Assay
Cinacalcet-treated or control cells (10
5
) suspended in 150µl α-MEM supplemented with
10% FBS and P/S were seeded on the upper transwell insert (5µm pore size) in a 12-well
plate (Corning, Inc., Corning, NY, USA). The bottom well was added with 500µl of
medium containing 100ng/ml murine SDF-1α (Peprotech, Inc., Rocky Hill, NJ). Cell
migration was allowed to continue for 3 hours at 37°C in a humidified, 5% CO
2
66
atmosphere. To control for nonspecific cell migration, chemokinesis of the cells was
measured by 1) allowing cells to migrate towards medium without SDF-1α and 2)
suspending cells in medium containing SDF-1α and allowing the cells to migrate towards
SDF-1α. Cells were harvested from the lower well and counted on a hemacytometer. The
relative numbers of the different stem and progenitor cell sub-populations were
subsequently analyzed by flow cytometry.
2.3.10 Cell Adhesion Assay
LKS
+
F
-
, LKS
+
F
+
and LKS
-
F
+
cells were sorted from BM MNCs and treated with
Cinacalcet as described above. Cells (5x10
2
) were added to wells coated with fibronectin
(10µg/ml) or collagen I (50µg/ml) (both from Becton Dickinson) in cell culture treated
96-well plates (Becton Dickinson) and incubated for 3 hours at 37°C in a humidified, 5%
CO
2
atmosphere. To control for nonspecific binding, adhesion to 1% BSA (Sigma-
Aldrich) was quantified. Non-adherent cells were washed off five times with 1X PBS,
and adherent cells were visually counted under a microscope.
2.3.11 Cell Cycle Analysis
To stain for DNA content, cells were incubated with 10µg/ml Hoechst 33342 (Sigma-
Aldrich) and 25µg/ml of verapamil at 37°C for 45 minutes, then stained with lineage and
stem cell antibodies as described above, except rat anti-mouse Flk-2 (eBioscience, Inc.,
San Diego, CA, USA) was used as the primary antibody and goat anti-rat IgG F(ab’)2-
APC-Cy7 (Santa Cruz Biotechnology, Santa Cruz, CA, USA) was used as the secondary
67
antibody. The stained cells were resuspended and fixed in 10% neutral buffered formalin
(EMD Chemicals, Inc.) and incubated at 4°C overnight, wrapped in foil. To stain for
RNA content, pyronin Y (Polysciences Inc, Warrington, PA) was added to the cells at a
final concentration of 0.75µg/ml and incubated at 4°C for 30 minutes wrapped in foil.
Cell cycle status was examined using the LSR II flow cytometer. Both pyronin Y and
Hoechst were analyzed on a linear scale. The gating scheme for cell cycle analysis is
shown in Figure 2.4.
Figure 2.4 Cell cycle analysis. A representative flow plot demonstrating the gating scheme is
shown. Each phase of the cell cycle is defined as followed. G
0
= Pyronin Y
-
/Hoechst
-
(Q3 in the
plot); G
1
= Pyronin Y
+
/Hoechst
-
(Q1 in the plot); S/G
2
/M = Pyronin Y
+
/Hoechst
+
(Q2 in the plot).
2.3.12 Apoptosis Assay
BM MNCs were stained with lineage and Sca-1 antibodies as described above. Following
Cinacalcet treatment, the cells were then stained with 7-Amino-Actinomycin (7-AAD)
and PE Annexin V (both from Becton Dickinson) according to manufacturer’s
instructions as followed. Annexin V binding buffer (10X) was first prepared with 0.1M
G
0
G
1
S/G
2
/M
68
HEPES (pH 7.4), 1.4M NaCl, and 25mM CaCl
2
in distilled water and 0.2µm sterile
filtered. Prior to staining cells, 1X working solution of the binding buffer was made by
diluting the 10X concentrate 1:10 with distilled water. Cells were stained with 7-AAD
and PE Annexin V at a concentration of 5:100 in Annexin V binding buffer (1X). Using
the LSR II flow cytometer, the percentage of apoptotic LKS
+
cells following treatment
was determined as 7-AAD negative and PE Annexin V positive.
2.3.13 Quantitative Reverse Transcription-Polymerase Chain Reaction (RT-PCR)
Total RNA was extracted using the RNA Miniprep Kit (Stratagene, La Jolla, CA, USA)
as followed. To lyse the cells, 0.7μl of 2βME was added to 100μl of lysis buffer for each
sample of ≤5×10
5
cells. Then, 100μl of the lysis buffer-2βME mixture was added to each
cell sample and vortexed vigorously until homogenized. An equal volume of 70% ethanol
was added to the cell lysate and mixed thoroughly by vortexing for 5 seconds. The
mixture was then transferred to an RNA-binding spin cup seated in a 2ml receptacle tube
and spun down in a microcentrifuge at maximum speed for 60 seconds. The spin cup was
retained and the filtrate was discarded. Next, 600μl of 1X low-salt wash buffer was added
to the cells and spun down for 60 seconds at maximum speed. The spin cup was retained
and the filtrate was discarded. The spin cup was placed in a new collection tube and spun
down one more time for 2 minutes at maximum speed to dry the fiber matrix. For each
sample, 5µl of reconstituted RNase-free DNase I was gently mixed with 25μl of DNase
digestion buffer and then directly added onto the fiber matrix of the spin cup. The
samples were then incubated at 37°C for 15 minutes. Then, 500μl of 1X high-salt wash
69
buffer was added to the samples and spun down for 60 seconds at maximum speed. Once
again, the spin cup was retained and the filtrate was discarded. The samples were washed
again twice, using 600μl and 300μl of 1X low-salt wash buffer for 60 seconds and 2
minutes respectively. The spin cup was then transferred to a 1.5ml collection tube, and
30µl pre-warmed elution buffer was directly added onto the fiber matrix. The samples
were incubated for 2 minutes at room temperature and finally spun down for 60 seconds
at maximum speed. The purified RNA in the elution buffer in was stored in -80°C for
long-term storage.
The purified RNA was reverse-transcribed into cDNA using the SuperScript VILO
cDNA synthesis kit (Invitrogen) in accordance with the manufacturer’s instructions. The
steps are as followed. For each single reaction, the following components were combined
in a PCR tube on ice:
5X VILO™ Reaction Mix 4µl
10X SuperScript
®
Enzyme Mix 2µl
RNA 10-14µl
DEPC-treated water bring up to 20µl
The reaction was performed using an Eppendorf AG 22331 Mastercycler machine
(Eppendorf, Hamburg, Germany). The thermal cycling conditions to generate cDNA
were as followed: 10 minutes at 25°C followed by 60 minutes 42°C. To terminate the
70
reaction the tubes were then incubated at 85°C for 5 minutes. The cDNA samples were
stored at -20°C until use.
To quantify the expression of car (Mm00443375_m1), cxcr4 (Mm01292123_m1), and
hprt1 (Mm00446968_m1), Taqman Gene Expression Assays primers (Applied
Biosystems, Branchburg, New Jersey, USA) and probe sets (Roche Diagnostics,
Mannheim, Germany) were used. The primers are intron-spanning to ensure that the PCR
products are not the result of amplifying contaminating genomic DNA segments. For
each gene, the reaction mix (10µl/reaction) was prepared and added to each well of a
384-well reaction plate (Eppendorf) on ice by pipetting 0.5µl of 20X TaqMan Gene
Expression Assay primers and 5µl of 2X FastStart Universal Probe Master (Rox), then
adding 4.5µl of the cDNA templates. Standard curves in these experiments were created
using total mouse kidney RNA obtained from C57Bl/6 mice. The kidney RNA was
reverse transcribed into cDNA and serially diluted for use in the following final
quantities: 50, 10, 2, 0.08, and 0.016ng. The 384-well plate was then sealed with a clear
adhesive film, centrifuged at 400g for five minutes, and loaded into the 7900HT real-time
PCR system (Applied Biosystems). The thermocycling conditions are detailed as follows:
50°C for 2 minutes
Initial denaturation: 95°C for 10 minutes
Denaturation: 95°C for 15 seconds
Annealing/Extension: 60°C for 60 seconds
40 cycles
71
No template control was used to assess for reagent contamination or primer-dimers.
2.3.14 Adhesion Molecule Expression
BM MNCs were treated with Cinacalcet or ethanol as the vehicle control, then stained
with lineage and Sca-1 antibodies as described above, along with 3µl of PE anti-mouse
CXCR4 (Becton Dickinson), CD49d, or CD62L antibodies (both from eBioscience, Inc.)
for CXCR4, α
4
β
1
integrins, and L-selectins, respectively. The expression levels of each
adhesion molecule on the LKS
+
cells were measured on the LSR II flow cytometer and
analyzed using FlowJo.
2.3.15 Immunohistochemistry
Tibias were dissected from C57Bl/6 mice and fixed in 10% neutral buffered formalin
overnight at 4°C. The bones were then decalcified with 20% EDTA over a two-week
period and were processed and embedded in paraffin using standard histological
procedures. Whole bone sections were cut into 5um thick and incubated with anti-
collagen I goal polyclonal IgG (Abcam, Cambridge, MA, USA) overnight at 4°C. Slides
were then blotted with donkey-anti-goat secondary antibody (Santa Cruz Biotechnology,
Santa Cruz, CA, USA) for 30 minutes at room temperature. The Vectastain ABC kit
(Vector Laboratories, Burlingame, CA, USA) was applied to the slides and incubated for
30 minutes at room temperature. The slides were then rinsed with PBS-Tween, and
Alkaline Phosphatase Vector Substrate was added to the blots. The slides were then
incubated in the dark for 10 minutes. Upon color formation, the slides were rinsed with
72
distilled water and then counterstained with Methyl Green. The slides were then cover-
slipped using Cytoseal XYL mounting medium (Richard-Allan Scientific, Kalamazoo,
MI, USA) and examined using a Nikon Eclipse 50i upright microscope (Nikon
Instruments, Inc., Melville, NY, USA).
2.3.16 In Vivo Homing
LK cells were sorted using a FACSAria or Aria II flow cytometer as described above and
treated with Cinacalcet or ethanol as the vehicle control. In order to track the cells in vivo
after co-injection of Cinacalcet-treated and control cells into the same mouse recipient,
LK cells (~5x10
5
to 1x10
6
) were labeled with a green fluorescent dye, either
carboxyfluorescein diacetate succinimidyl ester (CFSE) or 3,3'-
dioctadecyloxacarbocyanine (DiO), and a red fluorescent dye either
seminaphtharhodafluor (SNARF-1) carboxylic acid acetate succinimidyl ester or 1,1'-
dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine (DiI) (all from Invitrogen), according
to the procedures as followed.
For cell labeling with DiO or DiI, first the cells to be labeled were resuspended in 1ml of
1X PBS. Then, 5µl of DiO or DiI from the Vybrant
®
Multicolor Cell-Labeling Kit
(Invitrogen) was added to the cell suspension and mixed well. The cells were incubated at
37°C for 20 minutes in the dark, then centrifuged at 400g for 5 minutes at 37°C. Pre-
warmed α-MEM supplemented with 10% FBS and P/S was added to the cells, then the
cells were incubated at 37°C for 10 minutes in the dark for recovery. Finally, the cells
73
were centrifuged at 400g for 5 minutes and resuspended in approximately 0.3ml of 1X
PBS before injection.
For cell labeling with CFSE or SNARF-1, first the cells to be labeled were resuspended
in 1ml of 1X PBS. Then, 50µl of the 100µM CFSE or SNARF-1 stock was added to the
resuspended cells to yield a 5µM working solution. The cells were incubated for 15
minutes at 37°C in the dark. Next, the cells were centrifuged for 5 minutes at 400g,
resuspended in 1ml of pre-warmed growth medium, and incubated for 30 minutes at
37°C in the dark. Finally, the cells were resuspended in 0.3ml of 1X PBS before
injection.
Labeled cells from both groups were co-injected into the tail vein of a non-irradiated
C57Bl/6 mouse. Mice were then sacrificed after 16 hours, and the number of labeled cells
was measured in the BM and spleen through the detection of CFSE
+
and SNARF-1
+
cells
or DiO
+
and DiI
+
cells by flow cytometry.
2.3.17 In Vivo Lodgment
LK cells were labeled with 5µM CFSE or 5µM SNARF-1 cell-labeling solutions and
injected into the tail vein of non-irradiated C57Bl/6 mice, as described above. To block
CXCR4 function during lodgment, cells were treated with both 2.5µM Cinacalcet and
10µM AMD3100 (Sigma-Aldrich, St. Louis, MO, USA) or ethanol as the vehicle control
in the same fashion as described above. Tibias and femurs were dissected from the
74
recipient mice 16 hours after injection, decalcified for 3 days in Immunocal (Decal
Chemical Corporation, Tallman, NY, USA), and embedded in paraffin blocks after
processing. Bone sections of 5µm thickness were cut and mounted with Vectashield
containing 4’,6-diamidino-2-phenylindole (DAPI; Vector Laboratories). To assess the
lodgment of injected cells to the endosteal niche, the number of CFSE
+
(green) or
SNARF-1
+
(red) cells within two cell diameters from the endosteal surface were counted
on a total of 40-60 femoral and tibial sections per mouse. Each experiment was
performed with 1-3 mouse recipients.
2.3.18 Competitive Repopulation Assay
LKS
+
F
-
cells were sorted using a FACSAria or Aria II flow cytometer and then treated
with 2.5µM Cinacalcet or ethanol as the vehicle control. Following treatment, 450
LKS
+
F
-
cells (CD45.1) and 225,000 BM MNCs (CD45.2) were co-injected into the tail
vein of C57Bl/6 mice that were lethally irradiated at 9Gy approximately 24 hours before
transplantation. Engraftment levels and multilineage reconstitution were measured in
peripheral blood samples obtained from the tail of recipients starting week 4. Before
staining, 50µl of blood were added to 50µl of 1X PBS in polystyrene flow tubes (Becton
Dickinson), then PE anti-mouse CD45.1, FITC anti-mouse CD45.2, APC anti-mouse
CD3e, PE-Cy7 anti-mouse CD11b, and biotin anti-mouse B220 antibodies (3µl each) (all
from eBioscience) were added to stain the peripheral blood samples. The cells were
stained for 15 minutes on ice in the dark. Next, the cells were washed with 3ml of 1X
PBS, then centrifuged at 400g for 5 minutes. The cells were resuspended in 200µl of 1X
75
PBS and stained with 3µl of PE-Cy5 streptavidin (Becton Dickinson) for 15 minutes on
ice in the dark. After staining, the red blood cells were lysed in 1ml of 1X FACS lysing
solution (Becton Dickinson) for 1 minute at room temperature. The samples were then
fixed in 300µl of 10% neutral buffered formalin and analyzed by flow cytometry using
the LSR II cell flow cytometer. The data were analyzed using FlowJo.
2.3.19 Statistical Analysis
Comparison of experimental groups was performed using the paired or unpaired two-
tailed Student’s t-test as appropriate for the data set. A p-value of <0.05 was considered
significant.
76
2.4 Results
2.4.1 Cinacalcet treatment of primitive hematopoietic cells augments signaling
through the CaR.
We first wished to confirm that primitive murine hematopoietic cells express functional
CaR and that signaling through the receptor could be augmented by Cinacalcet treatment.
Cinacalcet has high selectivity for CaR and does not interact with a number of other G-
protein coupled receptors and mGluRs that share significant homology with CaR
(Nagano, 2006). In addition, because Cinacalcet is an allosteric modulator, it possesses
another level of specificity by being able to selectively propagate responses only in the
tissues where the physiological, endogenous agonist is active (Christopoulos, 2002). In
these experiments, primitive hematopoietic cells treated with Cinacalcet were subjected
to calcium flux analysis following stimulation with extracellular Ca
2+
ions. Analysis of
the more primitive LKS
+
F
-
subset or the more differentiated LKS
+
F
+
or LKS
-
F
+
subsets
of cells demonstrated maximal enhanced calcium flux upon stimulation of the CaR with
2.5μM Cinacalcet (Figure 2.5A), but not at lower concentrations (Figure 2.5B). These
effects were not attributable to augmentation of receptor expression, as quantitative RT-
PCR demonstrated that the treatment with Cinacalcet did not alter mRNA levels (Figure
2.5C). Altogether, these data demonstrated that Cinacalcet treatment enhances CaR
signaling on primitive hematopoietic cells without affecting receptor mRNA expression.
77
Figure 2.5 Effects of Cinacalcet treatment on CaR signaling and expression. (A) LK cells were
loaded with indo-1 dye (2µg/ml) as a fluorescent probe to mark intracellular Ca
2+
concentrations. CaCl
2
(1.5mM) was added as an exogenous source of Ca
2+
ions to induce a
response. The response in LKS
+
F
-
, LKS
+
F
+
, and LKS
-
F
+
sub-populations was then analyzed using
FlowJo software. Arrow indicates the addition of CaCl
2
stimulus. Green, control; red, Cinacalcet
(n=4 from 4 independent experiments). (B) A representative figure for calcium flux response at
limiting dosages is shown. LK cells were treated with Cinacalcet at dosages 0.625µM and
1.25µM, or with ethanol as the vehicle control. Calcium flux response is measured after the
addition of 1.5mM CaCl
2
as the stimulus. Cinacalcet treatment does not act in a dose dependent
manner. Arrow indicates the addition of CaCl
2
stinmulus. Orange, control; red, 0.625µM; blue,
1.25µM (n=4 from 2 independent experiments). (C) Total RNA was extracted from LK cells from
both groups. The expression level of car was measured using quantitative RT-PCR, with hprt1 as
the housekeeping gene. CaR mRNA expression was not affected by Cinacalcet treatment (n=8
from 5 independent experiments; error bars represent s.e.m.).
0
0.0005
0.001
0.0015
0.002
0.0025
0.003
0.0035
Control Cinacalcet
CaR/HPRT1
Time (seconds)
Indo-1 ratio
LKS
+
F
-
LKS
+
F
+
LKS
-
F
+
Time (seconds)
Indo-1 ratio
C
B
A
78
2.4.2 Cinacalcet treatment enhances in vitro primitive cell growth on stromal cell
layers without alteration of differentiation potential.
To address the question of whether Cinacalcet treatment alters the differentiation
potential of hematopoietic progenitor cells, the functional CFU-C assay was performed.
These data demonstrated that Cinacalcet treatment did not cause cellular toxicity or
inhibit differentiation potential of hematopoietic progenitor cells, as these cells were able
to form colonies comparable to the control group (Figure 2.6A). To assess whether more
primitive hematopoietic cell activity was altered by Cinacalcet treatment, the functional
cobblestone area-forming cell (CAFC) assay was performed. Compared to the control
group, CAFC activity was significantly higher in the Cinacalcet treatment group, as
evident in the LKS
+
F
-
sub-population (p<0.05) (Figure 2.6B). On the other hand,
Cinacalcet treatment did not appear to have any significant effects on CAFC activity in
the LKS
+
F
+
or LKS
-
F
+
sub-population (Figure 2.6B). These data indicate that with CaR
stimulation, primitive hematopoietic cells are more capable of being maintained in co-
culture, resulting in a higher frequency of CAFCs after the 5-week co-culture.
79
Figure 2.6 Effects of CaR stimulation on hematopoietic progenitor and primitive cell activity in
vitro. (A) BM MNCs treated with Cinacalcet or ethanol control were assessed for in vitro growth
potential using the CFU-C assay (n=10 from 5 independent experiments). (B) LKS
+
F
-
, LKS
+
F
+
and LKS
-
F
+
cells were seeded on a confluent stromal layer of supportive OP9 cells in serial
dilutions and cultured at 33°C and 5% CO
2
. Cobblestone areas were scored on Week 5 (*p<0.05,
n=9 for LKS
+
F
-
; n=7 for LKS
+
F
+
; n=3 for LKS
-
F
+
from 5 independent experiments; error bars
represent s.e.m.).
0
70
140
210
280
350
Control Cinacalcet
CFU-Cs/10
5
cells seeded
0
1.5
3
4.5
6
7.5
9
Control Cinacalcet
CAFCs/10
2
cells
LKS
+
F
-
0
0.1
0.2
0.3
0.4
0.5
0.6
Control Cinacalcet
CAFCs/10
2
cells
LKS
+
F
+
0
0.05
0.1
0.15
0.2
0.25
0.3
Control Cinacalcet
CAFCs/10
2
cells
LKS
-
F
+
*
A
B
80
2.4.3 Cinacalcet treatment augments adhesion of hematopoietic stem and progenitor
cells to ECM molecules.
Multiple different mechanisms may explain the increase in CAFC frequencies with CaR
stimulation, including altered cell cycle status, enhanced cell survival, or augmented cell
adhesion to ECM molecules. Specific analysis of each of these indicated that there was
no significant alteration in the number of cells residing in G
0
, G
1
, or S/G
2
/M phase of the
cell cycle immediately following ex vivo Cinacalcet treatment (Figure 2.7A). In addition,
there was no change in the percentage of apoptotic cells after Cinacalcet treatment
(Figure 2.7B), suggesting that CaR stimulation does not alter cell survival. Previous
immunofluorescence studies on the distribution of ECM molecules in the BM have
shown that two prominent molecules located in the BM to be collagen I and fibronectin,
with collagen I present in the endosteal region and fibronectin in the central marrow
region (Nilsson et al., 1998). Testing the ability of primitive hematopoietic cells to adhere
specifically to these ECM molecules, we found that although the control primitive
hematopoietic cells were able to bind to the cell culture plates, the addition of collagen I
or fibronectin did not enhance the binding capabilities of these cells. CaR stimulation, on
the other hand, significantly enhanced LKS
+
F
-
cell adhesion to collagen I, but not to
fibronectin (p<0.01) (Figure 2.7C). Furthermore, CaR stimulation slightly enhanced
LKS
+
F
+
cell adhesion to collagen I (Figure 2.7C). In contrast, CaR stimulation
significantly enhanced LKS
-
F
+
cell adhesion to fibronectin, but not to collagen I (p<0.01)
(Figure 2.7C). These results demonstrate that the effect of CaR stimulation was to
augment cell adhesion specifically to collagen I or fibronectin, which was distinctive
81
according to both the cell population and the ECM components expressed in the adult
BM.
Figure 2.7 CaR stimulation does not alter cell cycle status, or cell survival, but significantly
increases HSC adhesion to collagen I. (A) Cell cycle profiles for LKS
+
F
-
, LKS
+
F
+
, and LKS
-
F
+
sub-populations (n=4 from 2 independent experiments). (B) Percentage of LKS
+
cells undergoing
apoptosis following treatment. The percentage of apoptotic cells was determined as 7-AAD
negative and PE Annexin V positive (n=3 from 3 independent experiments). The bracketed area
represents the apoptotic population (C) Adhesion of LKS
+
F
-
, LKS
+
F
+
, and LKS
-
F
+
cells to
fibronectin and collagen I. Cells were allowed to adhere to wells coated with fibronectin and
collagen I for 3 hours at 37°C and 5% CO
2
. Bovine serum albumin (BSA, 1%) was used as a
control for nonspecific binding (**p<0.01, n=8 from 3 independent experiments; error bars
represent s.e.m.).
0
10
20
30
40
50
60
70
80
90
100
Percentage (%)
LKS
+
F
+
0
10
20
30
40
50
60
70
80
90
100
Percentage (%)
LKS
-
F
+
Control
Cinacalcet
0
10
20
30
40
50
60
70
80
90
100
Percentage (%)
LKS
+
F
-
A
82
Figure 2.7 Continued
However, it is important to note that adhesion to collagen I alone does not provide
specificity in the localization of primitive hematopoietic cells in the endosteal region of
the BM, as histological assessment showed that collagen I is expressed ubiquitously
C
B
0
0.5
1
1.5
2
2.5
Control Cinacalcet
Apoptosis (%)
7-AAD
Annexin V
Control
Cinacalcet
0
10
20
30
40
50
60
70
80
Adhesion (%)
LKS
+
F
-
**
0
10
20
30
40
50
60
70
80
Adhesion (%)
LKS
+
F
+
0
10
20
30
40
50
60
70
80
Adhesion (%)
LKS
-
F
+
Control
Cinacalcet
**
83
along the surface of the bone (Figure 2.8). Therefore, collagen I may only play a part in
the specific localization of primitive hematopoietic cells in the endosteal niche.
Figure 2.8 Collagen I is expressed ubiquitously along the bone. A tibial section stained with
anti-mouse collagen I is shown in the middle at 4x magnification. A picture of the same tibial
section is shown on the right at 20x magnification, zooming in on the endosteal region. Negative
control is shown on the left. Pink, collagen I; blue, nucleus.
2.4.4 Cinacalcet treatment augments primitive hematopoietic cell activity in vivo.
We next wished to investigate the effects of CaR stimulation on hematopoietic stem and
progenitor cell function in vivo using models of homing, lodgment, and engraftment. For
our homing studies, purified LK cells were differentially labeled with fluorescent dyes to
track the cells in vivo, and then treated with Cinacalcet or ethanol as the vehicle control.
These cells were then co-injected into the same non-irradiated mouse recipient, and the
frequency of cells homing to the BM and spleen was calculated using flow cytometric
analysis. Our results showed CaR stimulation by Cinacalcet treatment led to a significant
improvement in the ability of the injected cells to home to the BM (p<0.05) (Figure
2.9A), and a slight improvement to the spleen (Figure 2.9B). To test for the effect of
different dyes on homing efficiency, we labeled the cells with either DiI or SNARF-1
4x 4x 20x
84
(red dyes) and DiO or CFSE (green dyes) and analyzed the number of homed cells in the
recipient. We found that there was no effect on homing efficiency from the use of
different dyes to label the cells, as the same enhancement in homing was observed after
Cinacalcet treatment (Figure 2.9C). With respect to lodgment, histological assessment of
the femur and tibia showed that CaR stimulation resulted in an approximate three-fold
increase in the preferential anatomical localization of primitive hematopoietic cells at the
endosteal niche following homing (p<0.002) (Figure 2.9D), To assess whether the
increases in homing and lodgment led to increased engraftment of primitive
hematopoietic cells, we performed a competitive repopulation analysis using purified
LKS
+
F
-
cells. These data demonstrated increased engraftment of the primitive
hematopoietic cells following ex vivo Cinacalcet treatment over a 24-week period, with
significant increases in total engraftment levels starting on week 20 (p<0.05) (Figure
2.9E). In terms of the specific lineages, Cinacalcet treatment also gave rise to enhanced
multilineage reconstitution (p<0.05) (Figure 2.9E).
85
Figure 2.9 Cinacalcet treatment enhances in vivo homing, lodgment, and engraftment. (A)
Cinacalcet-treated or control LK cells were labeled with green or red membrane fluorescent
dyes, respectively. The percentage of labeled cells present in the BM after transplantation was
determined by flow cytometric analysis. Representative flow plots (left) and percentage of labeled
cells present in the BM (right) are shown (*p<0.05, n=4 from 4 independent experiments). (B)
Spleen cells from each recipient mouse were obtained, and homing was analyzed as for BM cells.
(C) The frequency of differentially labeled LK cells homing to the BM of the recipient mouse was
measured 16 hours post-injection using flow cytometric analysis. Control cells were labeled with
either DiI or SNARF-1. Cinacalcet-treated cells were labeled with either DiO or CFSE. The use
of either dye combination depicted the same enhancement in the percentage of labeled cells
homing to the BM after Cinacalcet treatment. Red dotted lines represent DiI and DiO dyes used;
blue dotted lines represent SNARF-1 and CFSE dyes used (n=4 from 4 independent experiments;
error bars represent s.e.m.). (D) Quantification of the percentage of LK cells labeled with CFSE
(Cinacalcet) or SNARF-1 (control) present in the BM within two cell diameters of the endosteal
surface in femoral and tibial sections. A representative picture of the anatomical localization of a
CFSE
+
LK cell (green) at the endosteal region and a SNARF-1
+
LK cell (red) away from the
endosteal region is shown. Cells were also stained with DAPI (blue) present in Vectashield
(***p<0.002, n=180 sections from 3 independent experiments.). (E) Competitive repopulation of
CD45.1 donor LKS
+
F
-
cells (Cinacalcet or control) and CD45.2 BM MNCs was analyzed by flow
cytometric analysis on peripheral blood samples obtained from the recipient mice. The
contribution of CD45.1 donor LKS
+
F
-
cells to multilineage reconstitution is also depicted
(*p<0.05, n=5; error bars represent s.e.m.).
0
0.005
0.01
0.015
0.02
0.025
0.03
0.035
0.04
% labeled cells in the BM
Control Cinacalcet
*
A
86
Figure 2.9 Continued
C
D
0
5
10
15
20
25
30
35
40
% labeled cells
Femur
Tibia
***
***
0
0.02
0.04
0.06
0.08
0.1
0.12
0.14
0.16
% labeled cells in the spleen
Control Cinacalcet
B
0
0.005
0.01
0.015
0.02
0.025
0.03
0.035
0.04
% labeled cells in the BM
Control Cinacalcet
87
Figure 2.9 Continued
2.4.5 The effects of CaR stimulation on primitive hematopoietic cells are mediated
through the CXCR4 signaling pathway.
We reasoned that the enhancement in BM homing and lodgment following Cinacalcet
treatment may be an effect of an altered response to chemotactic agents produced by the
BM stroma. Previous studies on the migratory response of murine HSCs have shown that
these cells migrate only towards the chemokine, SDF-1α, in vitro
(Wright et al., 2002).
Therefore, calcium flux assays in response to SDF-1α stimulus were conducted. Flow
cytometric analysis demonstrated that all hematopoietic stem and progenitor cells treated
with Cinacalcet had an enhanced, sustained calcium flux response to SDF-1α compared
* *
0
20
40
60
80
4 8 12 16 20 24
% CD45.1
Weeks
Total Engraftment
0
10
20
30
40
50
60
70
4 8 12 16 20 24
% CD45.1
Weeks
B Lymphoid Engraftment
Control
Cinacalcet
0
20
40
60
80
100
4 8 12 16 20 24
% CD45.1
Weeks
Myeloid Engraftment
0
20
40
60
80
4 8 12 16 20 24
% CD45.1
Weeks
T Lymphoid Engraftment
Control
Cinacalcet
* *
* *
*
* *
E
88
to the control (Figure 2.10A). Chemotaxis assays were then used to determine if
migration to SDF-1α was also enhanced following Cinacalcet treatment. With CaR
stimulation by Cinacalcet treatment, there were significant increases in the chemotactic
response to SDF-1α in all three sub-populations of cells, especially in the LKS
+
F
-
subset
(p<0.002) (Figure 2.10B). CXCR4 is the chemokine receptor for SDF-1α and its role in
mediating the retention of HSCs in the BM has been documented previously using
transgenic CXCR4
-/-
chimeric mice
(Foudi et al., 2006). To investigate whether the
increase in chemotactic response to SDF-1α was due to increased CXCR4 expression
after Cinacalcet treatment, CXCR4 mRNA and cell surface expression levels were
examined. Our results demonstrated that CaR stimulation by Cinacalcet treatment did not
induce CXCR4 mRNA or cell surface expression levels (Figure 2.10C,D). To further
investigate the effect of Cinacalcet treatment on the expression of adhesion molecules
important in interactions between HSCs and their niche, the expression levels of α
4
β
1
integrins and L-selectins on the cell surface were also examined. We found no alteration
in the expression levels of these adhesion molecules on the cell surface following
Cinacalcet treatment (Figure 2.10E,F). Altogether, these results suggest that with CaR
stimulation, CXCR4 signaling is enhanced without any alterations in the expression
levels of CXCR4, α
4
β
1
integrins, and L-selectins. Finally, to examine whether the effects
of CaR stimulation could be blocked by the CXCR4 antagonist, AMD3100, LK cells
were treated with both Cinacalcet and AMD3100 and co-injected with control treated
cells into the same recipients. Our results showed that when we stimulated CaR, but
blocked CXCR4, the enhancement in cell lodgment was eliminated (Figure 2.10G). This
89
suggests that the effects of CaR stimulation on hematopoietic stem and progenitor cells
are mediated in part through the CXCR4 signaling pathway.
Figure 2.10 CaR stimulation enhances CXCR4 signaling and cell migration towards SDF-1α,
but does not alter the expression of adhesion molecules. (A) Calcium flux assays in response to
100ng/ml SDF-1α were performed using purified LK cells. The response in LKS
+
F
-
, LKS
+
F
+
and
LKS
-
F
+
sub-populations was then analyzed using FlowJo software. Arrow indicates the addition
of SDF-1α stimulus. Green, control; red, Cinacalcet treated (n=3 from 3 independent
experiments). (B) In vitro chemotaxis assay of LKS
+
F
-
, LKS
+
F
+
and LKS
-
F
+
cells to 100ng/ml
SDF-1α following a 3-hour incubation. Blue columns represent chemokinesis controls, red
columns represent chemotaxis (***p<0.002, *p<0.05, n=3). (C) Total RNA was extracted from
LK cells from both groups. The expression level of cxcr4 was measured using quantitative RT-
PCR, with hprt1 as the housekeeping gene. CXCR4 mRNA expression was not affected by
Cinacalcet treatment (n=6 from 3 independent experiments; error bars represent s.e.m.). The
expression levels of CXCR4, α
4
β
1
integrins, and L-selectins were measured using flow cytometry
with PE-conjugated (D) CXCR4, (E) CD49d, and (F) CD62L antibodies, respectively. The
histograms for gating on the PE positive cell population is shown on the left, and the percentages
of cells expressing the corresponding adhesion molecules are shown on the right. Green, control;
red, cinacalcet (n=6 for CXCR4 from 4 independent experiments; n=5 for CD49d and CD62L
from 5 independent experiments; error bars represent s.e.m.). (G) In vivo lodgment showing the
percentages of injected LK cells localized at the endosteal region after ex vivo Cinacalcet and
AMD3100 treatment (n=160 sections from 2 independent experiments; error bars represent
s.e.m.).
LKS
+
F
-
LKS
+
F
+
LKS
-
F
+
Indo-1 ratio
Time (seconds)
(seconds)
A
90
Figure 2.10 Continued
0
20
40
60
80
100
120
140
160
Control Cinacalcet
CXCR4/HPRT1
0
5
10
15
20
25
30
Migration (%)
LKS
+
F
-
0
5
10
15
20
25
Migration (%)
LKS
-
F
+
– – + + + +
– + – + – +
SDF-1α
Cinacalcet
0
5
10
15
20
25
Migration (%)
LKS
+
F
+
– – + + + +
– + – + – +
SDF-1α
Cinacalcet
– – + + + +
– + – + – +
SDF-1α
Cinacalcet
***
*
*
B
C
91
Figure 2.10 Continued
F
0
0.04
0.08
0.12
0.16
0.2
Control Cinacalcet
% CXCR4
+
cells
0
5
10
15
20
25
30
Control Cinacalcet
% CD49d
+
cells
0
10
20
30
40
50
60
70
Control Cinacalcet
% CD62L
+
cells
D
E
92
Figure 2.10 Continued
0
5
10
15
20
25
30
35
40
Control Cinacalcet + AMD3100
% labeled cells
Femur
Tibia
G
93
2.5 Discussion
Here we demonstrated that enhancements in cell adhesion and CXCR4 signaling
associated with CaR stimulation promote homing, lodgment, and engraftment of
transplanted hematopoietic stem and progenitor cells. Specifically, the results indicated
that with CaR stimulation, HSC adhesion to collagen I, one of the major ECM molecules
released by cells of the osteoblastic lineage, was increased. Furthermore, with CaR
stimulation, HSCs became more responsive to SDF-1α, resulting in enhanced migration
towards this chemotactic agent present in the BM. A model of these coordinated
interactions is shown in Figure 2.11.
Figure 2.11 A proposed model for the role of CaR in HSC lodgment in vivo. Active bone
remodeling releases Ca
2+
ions into the endosteal region of the BM. HSCs arriving at the
endosteal region are able to sense the released Ca
2+
ions through the CaR. When the CaR is
activated by Ca
2+
ions, CXCR4 signaling is activated, which is known to be involved in cell
survival and the retention of HSCs in the BM. Furthermore, activation of the CaR also enhances
HSC adhesion to major ECM components in the adult BM, such as collagen I. This complex
interplay of intracellular signaling dictates the fate of the transplanted HSCs.
SDF-1α
Ca
2+
Stromal cell
Endosteum
Osteoblast
Osteoclast
Collagen I
CaR
CXCR4
HSC
Integrin
+
+
Bone
[Ca
2+
]
94
However, Cinacalcet treatment did not affect cell proliferation or the number of cells
undergoing apoptosis, suggesting that CaR stimulation does not affect the cell cycle
status of HSCs or their survival. Also, CaR stimulation by Cinacalcet treatment did not
alter the expression of the chemokine receptor CXCR4 and cell adhesion molecules such
as α
4
β
1
integrins and L-selectins. Interestingly, CaR stimulation appeared to have the
most significant effect on the most primitive LKS
+
F
-
sub-population compared to its
more mature counterparts in hematopoietic lineage commitment.
CaR was previously shown to be an important player in the regulation of HSC lodgment
and engraftment following transplantation, where CaR
-/-
HSCs were found to have
impaired ability to lodge in the endosteal surface of the bone, leading to defective
engraftment (Adams et al., 2006). This defect in CaR
-/-
HSCs may have been attributable
to a reduction in their ability to adhere to collagen I in the endosteal region. Collagen I is
mainly secreted by osteoblasts in the endosteal region, and several studies have shown
osteoblasts to be a key cellular component of the HSC niche (Zhang et al., 2003; Calvi et
al., 2003; Visnjic et al., 2004). In addition, a previous immunofluorescence study has
shown collagen I to be distributed non-uniformly throughout the endosteal region of the
murine BM, with some areas of the region exhibiting very high levels of expression while
others having much lower levels, such as the central marrow region (Nilsson et al., 1998).
Because collagen I is mainly secreted by osteoblasts, the effect of CaR stimulation in
enhancing adhesive interactions between LKS
+
F
-
cells and collagen I further confirms the
importance of osteoblasts in supporting the endosteal HSC niche. Furthermore, by
95
modulating the activity of CaR such that the receptor becomes more sensitive to
activation by Ca
2+
ions present in the endosteal region of the adult BM, this study
provides new observations on the specific adhesive interaction between collagen I and
LKS
+
F
-
cells in retaining these cells in the endosteal niche following transplantation.
Altogether, these results seem to indicate that both Ca
2+
ions released by active bone
remodeling and collagen I secreted by osteoblasts in the endosteal region of the BM play
important roles in HSC lodgment following transplantation, and that the CaR may be a
crucial player in this process that ultimately dictates the clinical outcome of the
transplantation. Since these molecules are specifically expressed in the adult bone, this
would further support our hypothesis that CaR allows for the preferential localization of
the HSCs in the adult BM (Adams et al., 2006).
Regulation of homing and lodgment of transplanted HSCs may involve crosstalk between
CaR and CXCR4 signaling pathways. In adult hematopoiesis, CXCR4, the receptor for
CXCL12 or SDF-1α, is known to play important roles in cell migration, proliferation, and
survival (Aiuti et al., 1997; Kahn et al., 2004).
In relevance to transplantation, when
CXCR4 is absent, retention of hematopoietic progenitors within the BM is impaired,
resulting in defective hematopoietic reconstitution as shown by a study on CXCR4
-/-
chimeric mice
(Foudi et al., 2006). Results from this study have indicated that CXCR4
signaling is elevated with CaR stimulation, leading to enhanced retention of HSCs in the
BM following transplantation. In fact, when CXCR4 is blocked by its antagonist
AMD3100, the enhancement in cell lodgment after CaR stimulation is eliminated. This
96
suggests that the CXCR4 signaling pathway may be an important mediator in the
regulation of hematopoietic stem and progenitor cell function by the CaR. In addition,
with CaR stimulation, the ability of LKS
+
F
-
cells to repopulate the BM and contribute to
hematopoiesis in the recipient mice is enhanced. However, the underlying mediator that
serves as the link between the CaR and CXCR4 signaling pathways is unknown.
Under homeostasis, HSCs reside very close to the endosteal surface of the bone (Lord et
al., 1975; Gong, 1978; Nilsson et al., 2001), where active bone remodeling occurs
through tightly regulated interactions between osteoblasts and osteoclasts, resulting in the
release of Ca
2+
ions from the bone into the endosteal region. Localization of HSCs in the
BM following transplantation has been one of the main focuses in the study of HSC niche
biology. Because HSCs are able to sense and respond to fluctuations in Ca
2+
concentrations through the CaR, the receptor might have a crucial role in the specifying
the localization of HSCs in the BM. By stimulating the CaR expressed on HSCs with the
clinically approved allosteric agonist, Cinacalcet, we have identified some of the
underlying mechanisms dictating the preferential localization of transplanted HSCs in the
endosteal region of the BM, thereby providing additional insights into HSC niche
biology. These mechanisms include activation of CXCR4 signaling and enhancement in
adhesion to key ECM components including collagen I and fibronectin that are expressed
in the adult BM, following CaR activation by extracellular Ca
2+
ions. Furthermore,
clinical implications of this study include the use of pharmacologic agents such as
97
calcimimetics to modulate the CaR activity ex vivo, thus enhancing HSC activity and
improving lodgment and engraftment of transplanted HSCs in vivo.
98
Chapter 3: Calcium-Sensing Receptor-Mediated Regulation of
Cell Proliferation and Interactions with the Bone Marrow
Microenvironment in Multiple Myeloma
3.1 Abstract
In our previous studies investigating the role of the CaR in normal hematopoiesis, we
showed the expression of the CaR on HSCs and the crucial role of the receptor in the
localization of HSCs within the endosteal region of the BM, where the stem cell niche is
located. However, the functional role of the CaR in pathologic hematopoiesis is poorly
understood. One hematological malignancy in which the CaR potentially plays an
important role is MM. MM is a B-cell malignancy characterized by the outgrowth of
plasma cells in the BM, with accompanying bone destruction involving bone resorption
due to excessively active osteoclasts. This leads to heightened levels of extracellular
calcium. To investigate the functional roles of the CaR in MM, we used a genetic
approach by performing a series of loss of function and gain of function studies. We used
lentiviral shRNAmir technology to downregulate the CaR on MM cells and discovered an
impairment of cell proliferation, specific cell migration towards SDF-1α, and cell
adhesion to fibronectin in vitro. In contrast, overexpression of CaR using a lentiviral
vector resulted in enhanced cell proliferation associated with increased cell cycle entry
into the S phase. Although specific cell migration towards SDF-1α was enhanced with
CaR overexpression, there was a decrease in cell adhesion to fibronectin, suggesting that
the CaR is partially involved in mediating adhesive interactions of MM cells and the BM
ECM. More importantly, the CaR plays a chemoprotective role in MM cells, as CaR
99
knockdown sensitizes MM cells to bortezomib treatment while CaR overexpression
protects MM cells from bortezomib treatment. We also generated the SCID-hu model of
MM by first implanting human fetal bone fragment subcutaneously into SCID mice and
then injecting MM cells directly into the human BM microenvironment. We showed that
the human MM cells can be reproducibly propagated in this human BM
microenvironment. More importantly, these human MM cells do not migrate to the
mouse BM, suggesting a preferential localization of human MM cells in the human BM
microenvironment. Collectively, these data suggest that the CaR play a crucial role in
mediating interactions between MM cells and the BM microenvironment and can
potentially be targeted for chemotherapeutic treatments in MM.
100
3.2 Introduction
MM is a B-cell malignancy characterized by the outgrowth of plasma cells in the BM,
with accompanying bone destruction involving osteolytic bone lesions due to the
accumulation of excessively active osteoclasts adjacent to the myeloma cells, leading to
heightened levels of extracellular calcium concentrations in the vicinity (Mundy, 1998;
Bataille et al., 1997; Roodman, 2004). In MM patients, coordinated osteoclastogenic
interactions mediated by increased secretion of RANK-L and IL-6 by BM stromal cells
and MIP-1α and MIP-1β by myeloma cells (Abe, 2011; Roodman, 2004; Taube et al.,
1994; Callander and Roodman, 2001; Han et al., 2001) altogether act to promote
osteoclastic bone resorption, resulting in hypercalcemia in about one-third of patients
with advanced stages of MM (Oyajobi, 2007).
Calcium is involved in the regulation of various physiological functions in different
biological systems, including blotting clotting, bone formation, and most importantly,
calcium homeostasis (Saidak et al., 2009b). In fact, extracellular calcium levels are
tightly maintained within a narrow range of 1.1 to 1.3mmol/L in the peripheral blood
(Dvorak et al., 2004; Maurer et al., 1996). In non-resorbing bone surfaces, extracellular
calcium concentration is ~2mmol/L (Berger et al., 2001), but can reach as high as 8 to
40mmol/L near bone resorbing osteoclasts (Silver et al., 1988). Because bone resorption
is increased in MM due to activation of osteoclastogenesis, and that osteoclasts
accumulate on bone resorbing surfaces adjacent to MM cells (Mundy, 1998), exposure of
MM cells to high calcium concentrations near bone lesions could contribute to the
101
pathophysiology of MM. One previous study showed that MM cells possess the capacity
to sense these changes in calcium concentrations through the CaR, a G-protein coupled
receptor that plays a key role in extracellular calcium homeostasis. The study
demonstrated that functional properties of MM cells can be modulated through the CaR
to contribute to the pathophysiology of MM, including increased cell proliferation and
expansion of myeloma cell mass (Yamagutchi et al., 2002).
In our previous studies investigating the role of the CaR in normal hematopoiesis, we
showed the expression of CaR on HSCs and the crucial role of the receptor in the
localization of HSCs within the endosteal region of the BM, where the stem cell niche is
located (Adams et al., 2006; Lam et al., 2011). Since MM cells are derived from
hematopoietic precursors, and interactions of the MM cells with the BM
microenvironment are known to be important for the growth, survival, and homing of
MM cells, we investigated the role of CaR in propagating the physiologic phenomena of
MM using a genetic approach. Specifically, we generated lentiviral vectors and
performed a series of gain of function and loss of function studies to study the role of
CaR in various cellular functions, including cell growth, migration, adhesion, and
survival. In addition, in order to address the involvement of CaR in the localization of
MM cells in vivo, we generated the SCID-hu mouse model of human MM by implanting
human fetal bones subcutaneously into SCID mice. The SCID-hu mouse model is unique
as both human hematopoietic cells and the human BM microenvironment are engrafted in
102
the mouse, which allows for the study of the localization of MM cells in the BM
microenvironment.
103
3.3 Materials and Methods
3.3.1 Cell Culture
Three human MM cell lines: U-266, RPMI-8226, and NCI-H929 MM (kind gifts from
Dr. Vinod Pullarkat, USC Stem Cell Transplantation Center, Los Angeles, CA, USA)
were used in this study. The sources of each cell line are depicted in the following table:
Cell line Source Reference
U-266 Peripheral blood of a 53-year-old male with
plasmacytoma
Nilsson et al., 1970
RPMI-8226 Peripheral blood of a 61-year-old male with
plasmacytoma
Gazdar et al., 1986
NCI-H929 BM of a 62-year-old female with
plasmacytoma
Matsuoka et al., 1967
Table 3.1 Information in regards to the sources from which the human MM cell lines were
derived. The three human MM cell lines were derived from the peripheral blood or the BM of
MM patients.
The three cell lines were cultured and maintained at the specified cell density and growth
medium as depicted in the following table:
104
Cell line Growth medium Cell density (cells/ml)
U-266 RPMI 1640 + 10% FBS + P/S 3x10
5
– 2x10
6
RPMI-8226 RPMI 1640 + 10% FBS + P/S 5x10
5
– 2x10
6
NCI-H929 RPMI 1640 + 10% FBS + P/S + 0.05mM
2βME
3x10
5
– 2x10
6
RPMI 1640 = RPMI 1640, 1X, with L-glutamine (Mediatech, Inc., Manassas, VA, USA)
FBS = Fetal bovine serum (Mediatech, Inc.)
P/S = Penicillin-streptomycin (Mediatech, Inc.)
2βME = β-Mercaptoethanol (Sigma-Aldrich, St. Louis, MO, USA)
Table 3.2 Culture conditions for the three human MM cell lines. The three human MM cell
lines were cultured using the appropriate growth medium and conditions as specified.
2βME was prepared at 0.1M by adding 21µl of concentrated 2βME into 3ml of 1X PBS
(Mediatech, Inc.) and passing through a 0.2µm filter (Pall Corporation, Newquay,
Cornwall, UK). The 0.1M 2βME stock is stored at 4°C for up to 3 weeks. The volume of
0.1M 2βME required for dosing is 1:2000 of the total medium volume. The medium is re-
dosed with 0.1M 2βME at 1:4000 of the total remaining medium volume every 3 weeks.
3.3.2 MM Patient Sample
BM aspirate was obtained from a 42-year-old male patient with MM. The patient had
hypercalcemia, elevated paraprotein levels, and multiple lytic lesions leading to bone
pain. BM smear morphological analysis revealed marked plasmacytosis varying from
mature to immature forms. The patient sample was processed as followed. First, the
105
sample was transferred to a 50ml tube (VWR International), and enough RPMI1640
medium was added to the sample to bring the total volume up to 35ml. Next, 15ml of
Ficoll-paque
®
PLUS medium (GE Healthcare, San Ramon, CA, USA) was added slowly
to the bottom of the suspension to avoid mixing, and the sample was centrifuged at
2000rpm for 20 minutes without brake. Finally, the interphase containing the
mononuclear cell layer was carefully aspirated and transferred to a new tube for
subsequent assays.
3.3.3 Animals
Six- to eight-week-old male C.B-17 scid/scid mice (Taconic Farms Inc, Oxnard, CA,
USA) were obtained and used in accordance with the University of Southern California
Institutional Animal Care and Use Committee (IACUC) guidelines. Mice were housed in
sterilized microisolator cages and received autoclaved food and water ad libitum. To
generate the SCID-hu mouse model, human fetal long bones (femurs, tibias, and humeri)
of 19 gestational weeks were obtained (Novogenix Laboratories LLC, Los Angeles, CA,
USA) and dissected into approximately 5x5x10mm fragments using a scalpel (Becton
Dickinson, Franklin Lakes, NJ, USA). The SCID mice were first anesthetized with
isoflurane (Phoenix Pharmaceuticals, Inc., St. Joseph, MO), then a 1cm incision was
made horizontally on the upper back of the mice after cleaning the area with a sterile
alcohol prep pad (Professional Disposables International, Inc., Orangeburg, NY, USA).
The connective tissue under the skin was loosened to create a subcutaneous pocket big
enough to accommodate the bone fragment. Then, the human fetal bone fragment was
106
implanted into the subcutaneous pocket, and the skin incision was closed by adding the
tissue adhesive, Vetbond™ (3M Animal Care Products, St. Paul, MN, USA), next to the
skin incision and covering the incision with the adjacent skin. The mice were kept warm
as they recovered from the anesthesia and monitored after the surgery.
3.3.4 Cell Growth
Cells (9x10
4
) were seeded in a 48-well plate in 0.3ml of growth medium in triplicates and
cultured at 37°C in a humidified, 5% CO
2
atmosphere. On days 2, 4, 6, and 8, cell counts
were taken by taking 10µl of cells and adding 10µl of 0.4% trypan blue solution
(AMRESCO, Solon, Ohio, USA) to the cells for viability. Viable cells were counted on a
hemacytometer using the Nikon Eclipse TS100 microscope (Nikon Instruments, Inc.,
Melville, NY, USA). The cell cultures were maintained at 0.3ml of total volume by
replenishing depleted growth medium.
3.3.5 Immunocytochemistry
Poly-L-lysine solution, 0.1% w/v (Sigma-Aldrich), was diluted 1:10 in deionized water.
Microscope slides (VWR International, Radnor, PA, USA) were coated with the diluted
poly-L-lysine solution for 30 minutes. The slides were then allowed to dry in an oven set
at 60°C for 1 hour. A drop of MM cells (~1x10
6
cells) was added to the end of a poly-L-
lysine coated slide and spread using the beveled edge of a clean microscope slide. The
slides were then allowed to air dry for 30 minutes. The cells on the slides were fixed in
4% paraformaldehyde (Alfa Aesar, Ward Hill, MA, USA) that was diluted in 1X PBS for
107
10 minutes at room temperature, then washed three times in 1X PBS for 5 minutes per
wash. Next, the slides were incubated in the blocking solution, 5% donkey serum (Sigma-
Aldrich) and 0.1% BSA (Sigma-Aldrich) in 1X PBS, for 30 minutes at room temperature,
then incubated overnight at 4°C in the primary antibody, mouse anti-human CaR
monoclonal antibody (clone: 5C10, ADD) (Pierce Antibodies, Rockford, IL, USA),
diluted to 2µg/ml in the blocking solution. Before adding the secondary antibody, the
slides were washed five times in 1X PBS for 3 minutes per wash. Then, the secondary
antibody, Alexa Fluor
®
594 donkey anti-mouse IgG (Molecular Probes, Carlsbad, CA,
USA), diluted to 4µg/ml in the blocking solution, was added to the slides and incubated
for 1 hour at room temperature. The slides were washed five times in 1X PBS for 3
minutes per wash. Finally, the stained cells were mounted with Vectashield containing
4’,6-diamidino-2-phenylindole (DAPI), coverslipped, (Vector Laboratories, Burlingame,
CA, USA), and visualized under the LSM5 PASCAL Axio upright confocal laser
scanning microscope (Carl Zeiss MicroImaging, Inc., Thornwood, NY, USA) with a UV
diode laser (405nm) and a HeNe (543nm) laser. A long pass (LP) filter, LP561, and a
band pass (BP) filter, BP420-480, were used to visualize the fluorescence from DAPI and
Alexa Fluor
®
594, respectively.
3.3.6 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-
2H-tetrazolium (MTS) Assay – Cell Proliferation
Cells were seeded in a 96-well plate (Becton Dickinson, Franklin Lakes, NJ, USA) at
2x10
5
cells/ml in 100µl of cell culture medium in triplicates. Absorbance readings
(490nm for assay; 660nm for reference) were measured on the SpectraMax M5
108
microplate reader (Molecular Devices, Downingtown, PA, USA) after adding 20µl of
MTS/PMS solution from the CellTiter 96
®
AQ
ueous
Non-Radioactive Cell Proliferation
Assay kit (Promega Corporation, Madison, WI, USA) and incubating the plate for 3
hours at 37°C in a humidified, 5% CO
2
atmosphere. The plate was wrapped in aluminum
foil to prevent exposure to light. Standard curves were created by seeding the cells at the
following cell densities (cells/ml): 1x10
5
, 2x10
5
, 3x10
5
, 5x10
5
, 7x10
5
, 1x10
6
, 2x10
6
, and
3x10
6
. Background readings were obtained from wells containing the same volume of
medium but without cells.
3.3.7 MTS Assay – Chemosensitivity
Cells were seeded in a 96-well plate (Becton Dickinson) at 1x10
6
cells/ml in 100µl of
growth medium in triplicates. Bortezomib (Millenium Pharmaceuticals, Inc., Cambridge,
MA, USA) was added to the cell cultures at the following dosages (nM): 0, 2, 5, 10, 20,
50. The cells were cultured in the presence of bortezomib overnight at 37°C in a
humidified, 5% CO
2
atmosphere. Absorbance readings were taken the next day by
following the same procedure as described above.
3.3.8 Transfer Vector – Loss of Function
Glycerol stocks for the pGIPZ lentiviral shRNAmir constructs (6 clones) targeting human
CaR were purchased (Open Biosystems, Lafayette, CO, USA). The pGIPZ lentiviral
shRNAmir constructs contain the turboGFP reporter driven by the CMV promoter to
track shRNAmir expression (Figure 3.1).
109
Figure 3.1 Diagram of pGIPZ lentiviral vector carrying CaR shRNAmir. The pGIPZ lentiviral
vector used for all loss of function studies is shown.
The glycerol stock was inoculated into a 100x25mm petri dish (VWR International)
containing solidified LB agar (EMD Chemicals, Inc., Gibbstown, NJ, USA)
supplemented with 100µg/ml of carbenicillin (EMD Chemicals, Inc.). The petri dish was
incubated for approximately 16 hours at 37°C. Individual colonies were picked using an
inoculating loop and transferred to 3ml of LB broth (EMD Chemicals, Inc.)
supplemented with 100µg/ml of carbenicillin. These individual colonies were cultured at
37°C for approximately 8 hours with vigorous shaking (225rpm), then transferred to a
larger culture vessel containing 250ml of LB broth supplemented with 100µg/ml of
carbenicillin. The larger cultures were incubated at 37°C for approximately 16 hours with
vigorous shaking, then the plasmids were extracted using the Plasmid Maxi kit
(QIAGEN, Valencia, CA, USA). For all loss of function studies, the GFP non-silencing
vector, which carries a sequence that has been verified to contain no homology to known
mammalian genes, was used as the control.
3.3.9 Transfer Vector – Gain of Function
pcDNA3 carrying the cDNA for human CaR (a kind gift from Dr. Edward Brown,
Brigham and Women’s Hospital, Boston, MA, USA) was sequenced, and primers
GIP Z
110
designed to flank the cDNA were used to specifically amplify the CaR cDNA using
polymerase chain reaction (PCR). Phusion DNA polymerase (Fermentas, Glen Burnie,
MD, USA) was used to perform the PCR reaction. A FLAG tag (5’-
CTTGTCATCGTCGTCCTTGTAGTC-3’) DNA sequence was added to the reverse
primer in between the cDNA and the stop codon such that the tag would be located on the
C-terminus of the CaR. In addition, BclI restriction sites were added to flank the CaR-
FLAG insert. The design of the primers (Integrated DNA Technologies, Inc., Coralville,
IA) for PCR amplication of the CaR is shown below:
Primer name Sequence
Forward hCaR-BclI insert 5’ – GGC CGC TGA TCA ATG GCA TTT TAT
AGC TGC – 3’
Reverse hCaR-FLAG-BclI
insert
5’ – TCT CCT TGA TCA TTA CTT GTC ATC GTC
GTC CTT GTA GTC TGA ATT CAC TAC GTT – 3’
Blue = 6bp corresponding to pcDNA3
Red = BclI sequence
Orange = Start codon (ATG) or stop codon (TTA)
Green = 15bp corresponding to CaR
Black = FLAG tag
Table 3.3 Primers for subcloning. The sequences of the forward and reverse primers used for
generating the CaR-FLAG insert are shown.
The PCR amplification product was run on a 0.8% agarose gel (VWR International) in
1X TAE buffer (EMD Chemicals, Inc.) next to the 1kb DNA ladder (New England
111
BioLabs, Inc.), and the band at ~3kb was cut from the gel and recovered using the
Zymoclean™ Gel DNA Recovery Kit (Zymo Research Corporation, Irvine, CA, USA).
The recovered PCR amplication product was then digested with BclI (New England
BioLabs, Inc., Ipswich, MA, USA) and cleaned using the DNA Clean & Concentrator™
kit (Zymo Research Corporation). The FUIGW lentiviral vector was used as the
backbone for cloning. The vector contains an internal ribosome entry site (IRES)-GFP
cassette downstream of the ubiquitin C promoter to allow visualization of the infected
cells. CaR-FLAG was cloned into the BamHI site on the FUIGW lentiviral vector as
followed. First, the FUIGW vector was digested with the BamHI restriction enzyme for 3
hours at 37°C and dephosphorylated with Antarctic Phosphatase (New England BioLabs,
Inc.) for 1 hour at 37°C. Next, the Antarctic Phosphatase was heat inactivated for 10
minutes at 65°C. Finally, the CaR-FLAG insert was ligated with 50ng of linearized and
dephosphorylated FUIGW vector at a molar ratio of 3:1 (insert:vector) using the T4 DNA
ligase in rapid ligation buffer (both from Fermentas). The ligation product was
transformed into DH5α as described above. The linearized and dephosphorylated FUIGW
vector was also transformed as the negative control. Colonies were picked and PCR
screened using 1µl of bacterial culture as the template with primers directed against the
CaR-FLAG insert. The primers were designed using the Primer3 online software (Rozen
and Skaletsky, 2000). The PCR amplification products were run on a 0.8% agarose gel
with the 100bp DNA ladder as the molecular weight marker (New England BioLabs,
Inc.), and the correct product size should be 434bp. The sequences of the primers used for
the screens are depicted in the table as follows:
112
Primer name Sequence
Forward hCaR right primer 5’ – AAG GGG GAC ATT ATC CTT GG – 3’
Reverse hCaR left primer 5’ – AGT CTG CTG GAG GAG GCA TA – 3’
Table 3.4 Primers for PCR screens. The primers used for PCR screening of positive clones are
shown.
Positive clones were cultured in LB broth supplemented with 100µg/ml of ampicillin, and
plasmids were extracted using the Plasmid Mini kit (QIAGEN). The plasmids were
digested with EcoRI for 3 hours at 37°C and ran on a 0.8% agarose gel in 1X TAE buffer
next to the undigested plasmids. The FUIGW vector is 10627bp, and the CaR-FLAG
insert is 3267bp, totaling 13894bp for the positive clones. After EcoRI digestion, positive
clones should yield bands at 11613bp, 1976bp, 258bp, and 46bp. The 1kb DNA ladder
was used as the molecular weight marker. Positive clones were sent for DNA sequencing
(Genewiz, Inc., La Jolla, CA, USA). For sequencing, 1000ng of template DNA and
25pmol of primer were suspended in 15µl total volume. The primers used for sequencing
are depicted in the table as follows:
113
Primer name Sequence
Primer_CaR cDNA back 5’ – AGG AAA CAG GTC TGC AAG GA – 3’
Primer_CaR cDNA middle 1 5’ – AGC CCC TCA TCA AGG AGA TT – 3’
Primer_CaR cDNA middle 2 5’ – CTG GTG TTT GAG GCC AAG AT – 3’
FUIGW-hCaR-FLAG left primer 5’ – CAC CCG TTC TGT TGG CTT AT – 3’
Primer_CaR cDNA middle 2’ 5’ – TAT GCC TCC TCC AGC AGA CT – 3’
Primer_CaR cDNA middle 3 5’ – GGC TCC ATC GTG TTT AAG GA – 3’
Table 3.5 Primers for DNA sequencing. The primers used for sequencing the CaR-FLAG insert
in FUIGW after subcloning are shown.
The FUIGW-CaR-FLAG overexpression lentiviral vector is shown in the diagram below:
Figure 3.2 Diagram of the FUIGW lentiviral vector carrying the CaR-FLAG insert. The
FUIGW lentiviral vector used for all gain of function studies is shown.
For all gain of function studies, the empty FUIGW lentiviral vector was used as the
control.
3.3.10 Packaging and Envelope Vectors
The 2
nd
generation packaging and envelope vectors, psPAX2 and pMD2.G, respectively,
were used to generate lentiviral vectors in 293T cells. The psPAX2 vector contains the
gag, pol, rev, and tat genes to package the viral particle, while the pMD2.G vector
provides a VSV-G envelope, which broadens the host range and stabilizes the viral
IRES CaR-FLAG GFP FUIGW
Ubi-C
114
particle produced. The vectors were transformed into DH5α competent cells (Invitrogen,
Grand Island, NY, USA) by incubating on ice for 30 minutes, then heat shocked for 30
seconds at 42°C. SOC medium (Invitrogen) was added to the transformation products and
incubated for 1 hour at 37°C in a rotating platform set at 225rpm. Bacterial cultures were
then prepared as described above. For preparation of plasmids for transfection, the
cultures were grown in LB broth supplemented with 100µg/ml of ampicillin (Acros
Organics, Geel, Belgium). The plasmids were extracted as described above.
3.3.11 Generating High-Titer Lentiviral Vectors
Lentiviral vectors were generated by transient transfection in 293T cells using the
jetPRIME™ DNA and siRNA transfection reagent (Polyplus, Illkirch, France). 293T
cells were grown to approximately 80% confluency in 17ml of Dulbecco’s modified
eagle medium (DMEM) supplemented with 10% FBS and penicillin-streptomycin (all
from Mediatech, Inc.) in a 145x20mm cell culture dish (Greiner Bio-One, Monroe, NC).
The cells were transfected with 3µg of transfer vector, 5µg of psPAX2, and 2µg of
pMD2.G. The transfection efficiency was determined by the percentage of cells
expressing GFP, visualized using the Zeiss AX10 Observer microscope (Carl Zeiss
MicroImaging, Inc.) attached to the X-Cite® Series 120 excitation system for
fluorescence illumination (Lumen Dynamics, Ontario, Canada). Viral supernatants were
harvested 48 and 72 hours after transfection and centrifuged at 3000rpm for 20 minutes at
4°C to remove cell debris. Then, the viral supernatants were passed through a 0.45µm
filter (Pall Corporation). The viral supernatants were concentrated in the Ultra-Clear™
115
centrifuge tubes (Beckman Coulter, Inc., Brea, CA, USA) by ultracentrifugation at
28,000rpm for 2 hours using the Optima L-100 XP ultracentrifuge (Beckman Coulter,
Inc.). The concentrated lentiviral particles were divided into 5-fold serial dilutions and
titered on 5x10
4
293T cells plated the day before. The viral titers were analyzed 48 hours
later based on the expression of turbo GFP using flow cytometry. The dilution containing
1-10% turbo GFP positive cells were used to determine the viral titer. The viral titer was
calculated using the formula below:
(5x10
4
) x (% GFP positive) x (Dilution factor)
Viral volume
3.3.12 Transduction of Cells
U-266 cells were resuspended in 1ml of growth medium then transduced with the
concentrated lentiviral particles at a multiplicity of infection (MOI) of 10 in the presence
of 4µg/ml polybrene (Sigma-Aldrich). The transduction was performed in a 15ml
centrifuge tube (VWR International) by spinning the cells with the lentiviral particles at
800g for 30 minutes at 32°C. Then, the cells were resuspended in the lentivirus-
containing medium and cultured at 5x10
5
cells/ml overnight at 37°C in a humidified, 5%
CO
2
atmosphere. The cells were replenished with fresh complete growth medium the
next day and cultured for an additional 72 hours before sorting.
= TU/ml
116
3.3.13 Flow Cytometry
Transduced cells were suspended in 1X PBS and sorted using the FACS Aria or Aria II
flow cytometer (Becton Dickinson) based on the expression of GFP. Cells were stained
with 100ng/ml of DAPI (Sigma-Aldrich) immediately before sorting for viability. The
DAPI
-
GFP
+
population was collected.
3.3.14 Apoptosis Assay
Annexin V binding buffer (10X) was prepared with 0.1M HEPES (pH 7.4), 1.4M NaCl,
and 25mM CaCl
2
in distilled water and 0.2µm sterile filtered. Prior to staining cells, 1X
working solution of the binding buffer was made by diluting the 10X concentrate 1:10
with distilled water. Cells were stained with 7-AAD and PE Annexin V at a concentration
of 5:100 (both from Becton Dickinson) according to manufacturer’s instructions in
Annexin V binding buffer (1X). Using the LSR II flow cytometer, the percentage of
apoptotic cells with or without bortezomib treatment was determined as 7-AAD negative
and PE Annexin V positive. A total of 50,000 events were recorded for analysis.
3.3.15 Quantitative Reverse Transcription-Polymerase Chain Reaction (RT-PCR)
Total RNA was extracted from cells by lysing with 0.5-1ml of TRIzol
®
Reagent
(Invitrogen) and purified using the Direct-zol™ RNA MiniPrep kit (Zymo Research
Corporation). Speficially for the purification of RNA, 1 volume of 100% ethanol (EMD
Chemicals, Inc.) was added to each volume of sample homogenate in TRizol
®
Reagent.
Then, the mixture was loaded into a Zymo-Spin™ IIC column in collection tube and
117
centrifuged at 12,000g for 1 minute. Next, 400µl of Direct-zol™ RNA PreWash was
added to the column and centrifuged at 12,000g for 1 minute. This step was repeated. The
column was then washed with 700µl of RNA Wash Buffer and centrifuged at 12,000g for
1 minute. The column was transferred to an autoclaved 1.7ml micro-centrifuge tube
(VWR International), and 30-50µl of nuclease-free water (Molecular Biologicals
International, Inc., Irvine, CA, USA) was added directly to the column matrix to elute the
RNA. 0.5-1µg of RNA was reverse-transcribed into cDNA using the Maxima
®
First
Strand cDNA Synthesis kit (Fermentas) in accordance with the manufacturer’s
instructions. The reaction conditions are as followed. First, the sample was incubated for
10 min at 25°C, then followed by 30 minutes at 30°C. Finally, the reaction was
terminated by heating at 85°C for 5 minutes. To quantify the expression of car
(Hs01047795_m1) and gapdh (Hs00266705_g1) as the housekeeping gene, Taqman
Gene Expression Assays primers (Applied Biosystems, Foster City, CA) and Maxima
®
Probe/ROX qPCR Master Mix (Fermentas) were used. The primers are intron-spanning
to ensure that the PCR products are not the result of amplifying contaminating genomic
DNA segments. The total PCR volume was 20µl, and 2µl of cDNA (10% of the final
PCR volume) was used per reaction. The thermocycling conditions are detailed below:
50°C for 2 minutes
Initial denaturation: 95°C for 10 minutes
Denaturation: 95°C for 15 seconds
Annealing/Extension: 60°C for 60 seconds
40 cycles
118
No template control was used to assess for reagent contamination or primer-dimers. Gene
expression levels were quantified using the 7900HT real-time PCR system (Applied
Biosystems). Relative expression levels were calculated using the Delta-Delta Ct method.
3.3.16 Chemotaxis Assay
Cells (1-5x10
5
) suspended in 150µl growth medium were seeded on the upper transwell
insert (5µm pore size) in a 12-well plate (Corning, Inc., Corning, NY, USA). The bottom
well was added with 500µl of medium containing 100ng/ml murine SDF-1α (Peprotech,
Inc., Rocky Hill, NJ). Cell migration was allowed to continue for 3 hours at 37°C in a
humidified, 5% CO
2
atmosphere. To control for nonspecific cell migration, chemokinesis
of the cells was measured by 1) allowing cells to migrate towards medium without SDF-
1α and 2) suspending cells in medium containing SDF-1α and allowing the cells to
migrate towards SDF-1α. Cells were harvested from the lower well and counted on a
hemacytometer.
3.3.17 Cell Adhesion Assay
Non-cell culture treated 96-well plates were coated with fibronectin (10µg/ml) and
collagen I (50µg/ml) overnight at 4°C (both from Becton Dickinson). The coated wells
were washed twice with 1X PBS, then blocked with 1% BSA for 30 minutes at room
temperature. Cells (10
4
) were seeded on coated wells and allowed to adhere for 3 hours at
37°C in a humidified, 5% CO
2
atmosphere. To control for non-specific binding, cells
119
were also seeded on wells coated with 1% BSA. Non-adherent cells were washed off five
times with 1X PBS, and adherent cells were counted under a microscope.
3.3.18 Cell Cycle Analysis
Cells (1x10
6
cells/ml) suspended in growth medium were stained with 10µg/ml Hoechst
33342 (Sigma-Aldrich) for DNA content and treated with 25µg/ml verapamil
hydrochloride concurrently (Sigma-Aldrich) at 37°C for 45 minutes wrapped in foil.
Cells were then washed with growth medium and resuspended in 10% neutral buffered
formalin (EMD Chemicals, Inc.) Cell cycle status was examined using the LSR II flow
cytometer.
3.3.19 Cell Surface Expression of CXCR4
Cells (5x10
5
) were stained with PE anti-human CXCR4 antibody (pre-diluted for use at
20µl/test) (Becton Dickinson) in 200µl of 1X PBS for 15 minutes on ice in the dark. For
control, cells (5x10
5
) were stained with PE Rat IgG
2b
isotype control (Becton Dickinson)
following the same procedure. After staining, the cells were washed with 5ml of 1X PBS
and resuspended in 300µl of 1X PBS for flow cytometric analysis on the LSR II.
3.3.20 Cell Labeling and Injections into SCID-hu Mice
Cells (2.5x10
6
) were labeled with 5µM of CFSE, a green fluorescent dye, or 5µM
SNARF, a red fluorescent dye (both from Invitrogen) as followed. First, the cells to be
labeled were resuspended in 1ml of 1X PBS. Then, 50µl of the 100µM CFSE or SNARF
120
stock was added to the resuspended cells to yield a 5µM working solution. The cells were
incubated for 15 minutes at 37°C in the dark. Next, the cells were centrifuged for 5
minutes at 400g, resuspended in 1ml of pre-warmed growth medium, and incubated for
30 minutes at 37°C in the dark. Finally, the cells were resuspended in 0.3ml of 1X PBS
before injection into the tail vein or 10µl of 1X PBS before injection into the human fetal
bone fragment.
For direct injection of labeled cells into the human fetal bone fragment, a Hamilton
syringe (705SN) (Hamilton Company, Reno, NV, USA) was first cleaned 3 times with
70% ethanol followed by 3 times with 1X PBS, then the labeled cells were aspirated into
the syringe. The SCID-hu mouse recipient was anesthetized with isoflurane as described
above, then the human fetal fragment was located using the thumb and forefinger. The
area for injection was cleaned with a sterile alcohol prep pad. The needle was inserted at
the base of the human fetal bone fragment along the length of the bone, then the cells
were injected slowly. After injection, the area was cleaned again with a sterile alcohol
prep pad.
For injection of labeled cells into the tail vein of a SCID-hu mouse, the recipient mouse
was first warmed under a heat lamp for 5-10 minutes to induce vasodilation. Meanwhile,
a sterile syringe with a 27-gauge needle attached (Becton Dickinson) was used to aspirate
the labeled cells for injection. Next, the recipient mouse was placed in a restrainer, and its
tail was stabilized between the thumb and forefinger of the hand that will not be
121
manipulating the syringe. The tail was cleaned with a sterile alcohol prep pad. With the
tail under tension, the needle was inserted near the middle part of the tail, bevel up,
approximately parallel to the vein located on either side of the tail. Once the entire length
of the needle was inserted into the vein, the cells were injected in a slow, fluid motion.
After injection, pressure was applied on the area of injection with a sterile alcohol prep
pad for approximately 30 seconds.
3.3.21 Immunohistochemistry
Tissues were fixed in 10% neutral buffered formalin overnight at 4°C. After fixation, the
tissues were washed with 1X PBS three times for 5 minutes per wash, then the tumor
tissue was suspended in 30% sucrose (EMD Chemicals, Inc.) and placed in 4°C
overnight. The human fetal bone was decalcified for one week using Immunocal (Decal
Chemical Corporation). The tissues were processed using the Tissue-Tec Vip-6 (Sakura
Finetek USA Inc., Torance, CA, USA) and embedded in paraffin following standard
histological procedures. Tissue sections were cut with a microtome at 5μm and used for
staining. The paraffin embedded sections were prepared for staining as follows. First, the
slides were baked at 58°C for 1 hour, then deparaffnized/rehydrated manually as depicted
below:
Xylene: 5 minutes, 2x
100% Ethanol: 5 minutes, 2x
95% Ethanol: 5 minutes, 2x
122
The slides were bathed in distilled water after the procedure. Antigen retrieval was
performed in Target Retrieval Solution (Dako North America, Inc., Carpinteria, CA,
USA) using the 2100 Retriever (PickCells Laboratories, Amsterdam, The Netherlands)
for 10 minutes. The slides were allowed to cool, then rinsed with distilled water. Next,
the slides were bathed in 1xPBS with 0.5% Tween 20 (Teknova, Hollister, CA, USA) for
5 minutes, then the EnVision+ System-HRP (DAB) kit (Dako North America, Inc.) was
used to stain for GFP according to manufacturer’s instructions as followed. First, enough
Peroxidase Block was applied to cover the specimens for 5 minutes, then the slides were
rinsed gently with distilled water and bathed in 1X PBS for 5 minutes. Excess buffer was
tapped off, and primary rabbit anti-turboGFP antibody (Evrogen, Moscow, Russia) was
applied at a concentration of 1:5000 (diluted from 1mg/ml stock in 0.05M Tris-HCl,
pH7.4 containing 1% BSA). For negative control, diluent containing no primary antibody
was applied to the specimens. The slides were incubated with the primary antibody at
4°C overnight. Next, the slides were rinsed once with 1X PBS and bathed in 1X PBS for
5 minutes. The Peroxidase Labeled Polymer was applied to the slides to cover the
specimen, then the slides were incubated at room temperature for 30 minutes.
Meanwhile, the Substrate-Chromogen Solution was prepared by adding one drop of
Liquid DAB+ Chromogen to 1ml of Substrate Buffer and mixed well. The slides were
rinsed as before, then enough of the prepared Substrate-Chromogen Solution was added
to cover the specimens for 5 minutes. The solution was removed, and the slides were
bathed in distilled water. Next, hematoxylin (EMD Chemicals, Inc.) counterstaining was
performed as depicted in the following:
123
Hematoxylin: 1 minute
Distilled water: rinse 3 times
0.25% Acid alcohol: 1 dip
Distilled water: bathe
0.037M Ammonium hydroxide (Sigma-Aldrich): 10 dips
Distilled water: bathe
The stained slides were dehydrated in 95% ethanol for 2 minutes, followed by 100%
ethanol for 2 minutes and finally xylene for 2 minutes. The slides were mounted with
Cytoseal XYL mounting medium (Thermo Scientific) and coverslipped. The slides were
examined using a Nikon Eclipse 50i upright microscope (Nikon Instruments, Inc.).
3.3.22 Statistical analysis
Comparison of experimental groups was performed using the unpaired two-tailed
Student’s t-test as appropriate for the data set. A p-value of <0.05 was considered
significant.
124
3.4 Results
3.4.1 The CaR is expressed in various human MM cell lines and a patient BM
sample.
A previous study documented the expression of the CaR in three human MM cell lines,
U-266, RPMI-8226, and IM-9 (Yamagutchi et al., 2001). To verify the expression of the
CaR in human MM cells, we investigated the presence of the receptor in three human
MM cell lines, U-266, RPMI-8226, and NCI-H929, as well as a BM sample obtained
from a patient with MM, BM5004. Quantitative RT-PCR revealed the expression of the
CaR on the mRNA level (Figure 3.3).
Figure 3.3 Expression of CaR mRNA in U-266, RPMI-8226, NCI-H929, and a BM sample
from a patient with MM. Total RNA was extracted and cDNA was synthesized. Using real-time
qPCR, the expression levels of car were measured (n=2-4 from 1 to 2 independent experiments).
Data represent relative expression levels normalized to gapdh and calculated using the Delta-
Delta Ct method. Data represent the mean ± s.e.m.
To examine the cell surface expression of the CaR, we performed immunocytochemistry
on non-permeabilized cells using a monoclonal antibody specific for the CaR.
125
Immunocytochemistry revealed the cell surface of CaR in the three human MM cell lines
(Figure 3.4).
Figure 3.4 Immunocytochemistry of U-266, RPMI-8226, and NCI-H929 cells for cell surface
CaR. Non-permeabilized MM cells were stained with a primary CaR-specific monoclonal
antibody (ADD) and a secondary Alexa Fluor
®
594 antibody, mounted with Vectashield
containing DAPI, and visualized using a confocal microscope. The photomicrographs were taken
at a magnification of 63x.
3.4.2 Reduced expression of CaR impairs MM cell growth in vitro.
In a previous study, it was shown that CaR agonists stimulated the proliferation of human
MM cells (Yamagutchi, et al., 2001). We first decided to further investigate the
involvement of the CaR in mediating MM cell growth in vitro. Using the pGIPZ
lentiviral shRNAmir construct, we were able to confirm CaR knockdown on the mRNA
level compared to the non-silencing control (Figure 3.5A). To monitor the ability of these
cells to grow in vitro, we performed an in vitro cell growth assay over a period of 8 days.
With CaR knockdown, there was a significant reduction in cell growth in vitro (Figure
3.5B). The significant difference in cell number on day 8 was also observed using
fluorescent microscopy (Figure 3.5C).
C aR / D API 63x C aR / D API 63x C aR / D API 63x
U- 266 R P MI- 8226 N C I- H92 9
126
Figure 3.5 Effects of CaR reduction on MM cell growth in vitro. (A) Total RNA was extracted
from U-266 cells transduced with pGIPZ non-silencing control vectors and pGIPZ CaR
shRNAmir vectors. The RNA was reverse-transcribed into cDNA, and the relative expression of
CaR was analyzed using real-time PCR (n=8 from 4 independent experiments). (B) 9x10
4
U-266
cells were seeded in growth medium, and cell counts were taken on days 2, 4, 6, and 8 (n=6 from
2 independent experiments). (C) The photomicrographs were taken at a magnification of 10x on
day 8 of the cell growth assay using a fluorescent microscope. Data represent the mean ±
s.e.m.*p<0.05 compared to the non-silencing control.
3.4.3 CaR overexpression enhances MM cell proliferation in vitro.
To further investigate the involvement of the CaR in the proliferation of human MM
cells, we used the FUIGW lentiviral construct to overexpress the CaR in MM cells.
Overexpression of the CaR was confirmed using quantitative RT-PCR (Figure 3.6A) and
A B
C
GFP non-silencing control CaR shRNA
Day 8:
10x 10x
127
immunocytochemistry for cell surface CaR (Figure 3.6C). To determine the effects of
CaR overexpression on MM cell proliferation, we performed the colorimetric MTS assay
on over a period of 9 days. With CaR overexpression, we discovered that there was an
increase in cell proliferation on day 4 compared to the empty vector control, but as the
cells became confluent and the nutrients in the growth medium were depleted, the
difference diminished.
Figure 3.6 Effects of CaR overexpression on MM cell proliferation in vitro. (A) Total RNA was
extracted from U-266 cells transduced with FUIGW empty control vectors and FUIGW-CaR-
FLAG overexpression vectors. The RNA was reverse-transcribed into cDNA, and the relative
expression of CaR was analyzed using real-time PCR (n=3-5 from 2 independent experiments).
(B) U-266 cells were seeded in a 96-well plate day 0 at a cell density of 2x10
5
cells/ml. After the
addition of MTS/PMS solution, the cells were incubated for 3 hours at 37°C in a humidified, 5%
CO
2
atmosphere, absorbance readings were taken. Absorbance readings were also taken on days
2, 4, 6, and 9 (n=3-6 from 2 independent experiments) to monitor cell proliferation. (C) Non-
permeabilized U-266 cells transduced with FUIGW empty vectors or FUIGW-CaR-FLAG
overexpression vectors were stained with a primary CaR-specific monoclonal antibody (ADD)
and a secondary Alexa Fluor® 594 antibody, mounted with Vectashield containing DAPI, and
visualized using a confocal microscope. The photomicrographs were taken at a magnification of
63x. Data represent the mean ± s.e.m.*p<0.05 and ***p<0.00005 compared to the empty vector
control group.
A B
128
Figure 3.6 Continued
3.4.4 CaR overexpression does not lead to a decrease in apoptosis but a shift of cell
cycle status from the G
1
phase to the S phase.
Two underlying mechanisms that could lead to enhanced cell proliferation are increased
cell survival or entry into the S phase of the cell cycle. To investigate the mechanisms
that drive cell proliferation in CaR overexpressing MM cells, we performed the apoptosis
assay and analyzed the cell cycle status. Using 7-AAD and Annexin V staining, we
observed no difference in the apoptotic status of MM cells with CaR overexpression
(Figure 3.7A, B). In contrast, cell cycle analysis revealed a decrease in G
1
phase with a
concomitant increase in S phase in MM cells when the CaR is overexpressed (Figure
3.7C). Taken together, the enhancement of MM cell proliferation with CaR
overexpression is a result of increased exit from G
1
phase and entry into S phase of the
cell cycle.
C
63x C aR / D API 63x
Control
C aR / D API
CaR
129
Figure 3.7 Effects of CaR overexpression on apoptosis and cell cycle status. (A) Representative
flow cytometry plots depicting the gating schemes for cells undergoing apoptosis. U-266 cells
from the empty vector control group or the CaR overexpression group were stained with 7-AAD
and PE Annexin V. The apoptotic population is defined as 7-AAD
-
Annexin V
+
. (B) Quantification
of apoptotic cells (n=3). (C) Cell cycle status was determined by Hoechst 33342 staining
followed by flow cytometric analysis (n=2-3). Data represent the mean ± s.e.m.
3.4.5 Reduced expression of CaR decreases cell migration towards SDF-1α in vitro
and impairs cell adhesion to major ECM molecules in the BM.
The BM microenvironment is a complex microenvironment that contains both cellular
and non-cellular components that interact with MM cells to propagate the disease. It is
clear that the homing of both normal and malignant BM cells is mediated by the SDF-
GFP control
CaR
Annexin V PE-A
7 AAD-A
Annexin V PE-A
7 AAD-A
B
A
1.5%
1.7%
C
130
1/CXCR4 axis (Dar et al., 2005; Tavor et al., 2004). In addition, MM cells express an
array of integrins, including VLA-4, that serve as major anchors mediating physical
interactions of MM cells and the ECM of the BM microenvironment (Uchiyama et al.,
1992). In fact, the BM ECM of MM patients were characterized to contain fibronectin
and collagen I (Kibler et al., 1998; Tancred et al., 2009), which MM cells can adhere to
via cell surface integrins. To examine the functional effects of reduced expression of CaR
on cell migration towards SDF-1α and cell adhesion to fibronectin and collagen I, we
performed the chemotaxis assay and the cell adhesion assay. With CaR knockdown, there
was a significant decrease in the specific migration of MM cells towards SDF-1α (Figure
3.8A). In addition, associated with CaR knockdown was a significant impairment in the
ability of MM cells to adhere to fibronectin (Figure 3.8B).
131
Figure 3.8 Effects of CaR knockdown on cell migration and cell adhesion. (A) Chemotaxis
assay of U-266 cells migrating towards SDF-1α in vitro. Cells were seeded onto the upper
transwell insert and allowed to migrate towards 100ng/ml of SDF-1α for 3 hours at 37°C in a
humidified, 5% CO
2
atmosphere. The number of migrated cells on the bottom well was counted.
Chemokinesis was performed to measure spontaneous, nonspecific cell movements. The
chemotactic index was calculated as the number of cells migrated in response to SDF-1α divided
by the number of spontaneously migrated cells (n=3). (B) Cell adhesion to fibronectin and
collagen I. Cells were allowed to adhere to 10µg/ml of fibronectin and 50µg/ml of collagen I for
3 hours at 37°C in a humidified, 5% CO
2
atmosphere. Cells that did not adhere were washed off
with 1X PBS. Cells that adhered were counted under a microscope. Cell adhesion to 1% BSA was
used as a control for nonspecific binding (n=3). Data represent the mean ± s.e.m.*p<0.05 and
**p<0.005 compared to the non-silencing control group.
3.4.6 CaR overexpression increases cell migration towards SDF-1α in vitro but
impairs cell adhesion to major ECM molecules in the BM.
To further investigate the functional role of CaR in cell migration and cell adhesion, we
assessed the effects that overexpression of CaR has on MM cells in their ability to
migrate towards SDF-1α and adhere to fibronectin and collagen I, similar to the loss of
function studies. With CaR overexpression, there was an increase in specific cell
migration towards SDF-1α in vitro (Figure 3.9A). Interestingly, CaR overexpression
A
B
132
impairs the ability of MM cells to adhere to fibronectin (Figure 3.9B). Collectively, the
loss of function and gain of function studies on cell migration and cell adhesion suggest
that CaR positively regulates MM cell migration towards SDF-1α in vitro. On the other
hand, CaR is necessary for cell adhesion to fibronectin in MM, but the precise
mechanism for the involvement of CaR in cell adhesion is unclear.
Figure 3.9 Effects of CaR overexpression on cell migration and cell adhesion. (A) Chemotaxis
assay of U-266 cells migrating towards SDF-1α in vitro (n=6 from 2 independent experiments).
(B) Cell adhesion to fibronectin and collagen I (n=3-6 from 2 independent experiments). Data
represent the mean ± s.e.m.
3.4.7 With CaR overexpression, the enhancement in specific cell migration towards
SDF-1α is not a result of increased cell surface expression of CXCR4.
From the chemotaxis assay, we discovered that CaR overexpression led to enhanced
specific cell migration towards SDF-1α. To determine if this effect was associated with
an increased cell surface expression of CXCR4, we performed cell surface staining with
A B
133
an antibody specific for human CXCR4 and analyzed the expression using flow
cytometry. We found that with CaR overexpression, there was no alteration in the cell
surface expression of CXCR4 (Figure 3.10), suggesting that the increase in cell migration
towards SDF-1α was not due to increased CXCR4 expression on the cell surface.
Figure 3.10 Cell surface expression of CXCR4. Representative flow cytometry histogram
demonstrating the cell surface expression of CXCR4 on MM cells. U-266 cells from the empty
vector control group or the CaR overexpression group were stained with PE anti-human CXCR4
antibody. For the isotype control, the cells were stained with PE IgG
2b
antibody (n=2).
3.4.8 CaR plays a chemoprotective role in MM cells.
The oncogenic properties of the CaR have been reported in studies on bone metastatic
cancers, such as breast and prostate cancers (Mihai et al., 2006; Liao et al., 2006). To
determine if the CaR has a chemoprotective function in MM, we cultured CaR
knockdown MM cells in the presence of bortezomib, a proteasome inhibitor used
clinically to treat MM patients. Our results revealed that with CaR knockdown, there was
increased cell death associated with an increase in apoptotic status compared to the non-
134
silencing control (Figure 3.11A, B). In contrast, when the CaR is overexpressed, the MM
cells became less chemosensitive to high dosages of bortezomib (Figure 3.11C). Taken
together, these results suggest that the CaR plays a chemoprotective role in MM cells.
Figure 3.11 Chemosensitivity of MM cells in the presence of bortezomib. (A) 9x10
4
U-266 cells
from the non-silencing control group or the CaR knockdown group were put into culture on day 0
in the presence of 10nM bortezomib. Viable cells were counted using trypan blue exclusion on
days 2, 4, 6, and 8 (n=6 from 2 independent experiments). (B) Apoptosis assay using 7-AAD and
Annexin V staining. U-266 cells from the non-silencing control group or the CaR knockdown
group were treated with bortezomib for 1 day, and the percentage of apoptotic cells were
measured using flow cytometry (n=3). (C) Chemosensitivity of MM cells overexpressing the CaR.
U-266 cells from the empty vector control group or the CaR overexpression group were treated
with 0, 2, 5, 10, 20, 50ng of bortezomib for 1 day, and the percentage of viable cells were
measured using the MTS assay (n=3). Data represent the mean ± s.e.m.*p<0.05 and **p<0.005
compared to the non-silencing control group.
A B
135
Figure 3.11 Continued
3.4.9 The SCID-hu model of MM is a relevant model for the study of human MM.
The importance of the BM microenvironment in the pathogenesis of MM is increasingly
recognized. MM cells grow and expand almost exclusively in the BM, which reinforces
the importance of the BM microenvironment in supporting MM cell growth and survival.
In fact, recent development of chemotherapeutic agents including bortezomib has been
focused on targeting MM cells in their BM microenvironment (Hideshima et al., 2007;
Laubach et al., 2010; Palumbo and Anderson, 2011). However, one major drawback to
studying the mechanisms responsible for MM has been a lack of suitable animal models
of the human disease that replicate the complexity of tumor-host cell interactions. In
order to generate a mouse model for the study of human MM, we first generated the
SCID-hu model by subcutaneously implanting human fetal bone fragments into SCID
mice and waiting for at least 4 weeks for vascularization to occur (Figure 3.12A). Next,
to determine the proper method for introducing MM cells into the human BM
microenvironment, we labeled the MM cells with CFSE or SNARF, then injected the
C
136
CFSE-labeled cells directly into the human fetal bone fragment and the SNARF-labeled
cells into the tail vein of the same SCID-hu mouse recipient. The reverse was also
performed to eliminate any effects from the cell labeling. The recipient SCID-hu mouse
was sacrificed approximately 16 hours later to examine the presence of labeled cells
using flow cytometry. As expected, cells that were injected directly into the human fetal
bone fragment were detectable by flow cytometry (Figure 3.12B,C). However, cells that
were injected into the tail vein of the SCID-hu mouse recipient were not detectable in the
human fetal bone fragment (Figure 3.12B,C). Taken together, these results suggest that in
order to introduce MM cells into the human BM microenvironment, the cells have to be
injected directly into the human bone fragment. It is also interesting to note that the
labeled cells were not detectable in the mouse BM or the spleen of the SCID-hu mouse
recipient (Figure 3.12D), suggesting that the labeled cells preferentially localize and
retain in the human BM microenvironment within the 16-hour timeframe.
137
Figure 3.12 Introducing MM cells into the SCID-hu mouse model. (A) Generating the SCID-hu
mouse model. A human fetal bone (FB) is cut into ~5x5x10mm fragments and implanted
subcutaneously (s.c.) into the upper back of a SCID mouse, as shown in the diagram. After a
minimum of 4 weeks, the mouse was sacrificed, and vascularization was observed as depicted in
the photographs. The photograph on the right is zoomed in on the human FB fragment. (B)
CFSE-labeled cells were injected directly into the implanted human FB fragment while the
SNARF-labeled cells were injected into the tail vein of a SCID-hu mouse recipient as shown in
the diagrams on the left. The mouse was sacrificed ~16 hours after injection, and the human FB
fragment was obtained for flow cytometric analysis for the presence of CFSE
+
or SNARF
+
cells.
(C) Same procedure as depicted in (B), except that SNARF-labeled cells were injected directly
into the human FB fragment while CFSE-labeled cells were injected into the tail vein of a SCID-
hu mouse recipient. (D)Representative flow cytometry plots for detecting the presence of CFSE
+
or SNARF
+
cells in the mouse BM and spleen of a SCID-hu recipient.
≥ 4 weeks for vascularization
Human
FB
s.c.
SCID
A
Human FB 1
SCID-hu
CFSE-U-266
SNARF-U-266
B
138
Figure 3.12 Continued
3.4.10 The SCID-hu model can be adapted to study homing and localization of MM
cells.
In a previous study, an in vivo model was developed for the study of MM localization
and growth within the human BM microenvironment using SCID-hu mice implanted with
bilateral human fetal bone grafts (Urashima et al., 1997). This in vivo model was found
to harbor the hallmarks of human MM, including tumor cell growth accompanied with
secretion of monoclonal human Ig and human IL-6 in sera of tumor bearing mice
Human FB 2
SCID-hu
SNARF-U-266
CFSE-U-266
C
Mouse BM Mouse spleen
D
139
(Urashima et al., 1997). To verify that this SCID-hu model can be adapted to study
homing and localization of MM cells, we have generated SCID-hu mice by implanting
human fetal bone fragments bilaterally onto the back of the SCID mice and assessing
tumor cell growth and localization (Figure 3.13A). Using this in vivo model, we were
able to observe human myeloma cell growth in the human fetal bone implant in SCID-hu
mice (Figure 3.13B). Interestingly, the injected human MM cells did not home to the
mouse BM, although they are detectable in the peripheral blood circulation (Figure
3.13C). Furthermore, using immunohistochemistry, we were able to detect the presence
of GFP
+
cells in the contralateral human fetal bone implant, suggesting migration or
metastasis of the injected MM cells to the contralateral human fetal bone fragment. Taken
together, these findings suggest that the SCID-hu model is an excellent model for
studying the homing and localization of human MM cells in a human BM
microenvironment.
140
Figure 3.13 Adapting the SCID-hu model for the study of homing and localization of MM
cells. (A) Schematics for generating the SCID-hu model of MM with bilateral human fetal bone
implants. A SCID mouse was subcutaneously implanted with bilateral human fetal bone
fragments of approximately 5x5x10mm each. After a minimum of 4 weeks post-surgery, the SCID-
hu mouse was exposed to a sublethal dose of 400 centi-Grays (cGy) of irradiation. To introduce
MM cells into the human BM microenvironment of the SCID-hu mouse, 7x10
4
GFP
+
U-266 cells
were injected directly into the marrow cavity of the left fetal bone implant. The SCID-hu mouse
was sacrificed at week 8 due to tumor burden. (B) Assessment of tumor growth and metastasis.
Photographs in the top panel depict the tumor growth on week 8 in the left human fetal bone
implant. Fluorescence microscopy revealed that the tumor was GFP
+
, indicating monoclonal
tumor cell growth from the injected GFP
+
U-266 cells. Photographs in the bottom panel depict
immunohistochemistry for GFP
+
cells in the tumor. Negative control staining is shown on the left.
Photographs were taken at 20x magnification. (C) Flow cytometric analysis revealed the
presence GFP
+
cells in peripheral blood circulation, but not in mouse BM.
Immunohistochemistry performed on the right fetal bone implant revealed the presence of GFP
+
cells. Negative control staining is shown on the left. Arrowheads point to the GFP
+
cells.
Photographs were taken at 20x magnification.
≥ 4 weeks
400cGy
7x10
4
GFP
+
U-266 cells
SCID-hu SCID-hu SCID-hu
A
141
Figure 3.13 Continued
B
Week 8 tumor
GFP Merged Bright field
142
Figure 3.13 Continued
C
Human FB
143
3.5 Discussion
The oncogenic role of the CaR has been implicated in a range of bone metastatic cancers
such as breast and prostate cancers, both of which, in advanced stages, preferentially
localize to bones displaying high rates of bone turnover (Schneider et al., 2005; Cicek
and Oursler, 2006). Although the CaR plays a similar role in bone destructive MM as
reported by a previous study, which demonstrated that exposure of MM cells to agonists
of the receptor enhances cell proliferation in vitro (Yamagutchi et al., 2002), the precise
underlying mechanisms were not clearly defined. In addition, because the importance of
the BM microenvironment is increasingly recognized in the pathogenesis of MM, it is
imperative to incorporate this cell extrinsic perspective into the study of MM
pathophysiology. The BM microenvironment of MM has a distinctively high level of
calcium owing to a concerted effort of BM stromal cells and MM cells in providing
stimulatory signals for osteoclastogenesis, resulting in excessive bone resorption that
leads to osteolytic bone lesions. In the present study, we first verified the expression of
the CaR in three human MM cell lines, U-266, RPMI-8226, and NCI-H929, that allows
MM cells to sense the locally high level of calcium to mediate functional changes. We
confirmed the expression of the CaR in the human MM cell lines on both the mRNA and
protein level. In addition, to provide clinical relevance, we also showed that the
expression of the receptor on the mRNA level in a BM sample obtained from a MM
patient with marked plasmacytosis. To further investigate the functional roles of the CaR
in MM, we generated two lentiviral vectors for conducting loss of function and gain of
function studies. In our loss of function studies, we demonstrated that with CaR
144
knockdown there was decreased cell proliferation in vitro. In contrast, we observed
enhanced cell proliferation in vitro associated with increased cell cycle entry into the S
phase when the CaR was overexpressed in our gain of function studies. However, the
CaR did not appear to play a major role in cell survival in MM cells, as CaR
overexpression did not reduce apoptosis. Taken together, these functional studies support
the role of the CaR in modulating MM cell proliferation.
In MM, the BM microenvironment provides a fertile soil with favorable conditions not
only for the growth of the cancer cells, but also for the homing of cancer cells to the BM.
In fact, compelling evidence is accumulating that cancer cells may employ several
mechanisms involving chemokine-chemokine receptor axes during their metastasis to the
bone, and that these mechanisms also regulate the trafficking of normal cells (Kucia et
al., 2004). The most extensively studied chemokine in the migration of bone metastatic
cancer cells is SDF-1α. In MM, SDF-1α secreted by BM stromal cells regulates the
migration and homing of MM cells to the BM microenvironment by interacting with its
receptor, CXCR4, expressed on myeloma cells (Alsayed et al., 2002; Hideshima et al.,
2002; Hideshima et al., 2007). In fact, upregulation of SDF-1α has been reported in the
BM of MM patients (Zannettino et al., 2005), potentially to confine the myeloma cells to
the BM and to prevent further trafficking of the cells. On the other hand, when myeloma
cells from patients were mobilized with cyclophosphamide and granulocyte-macrophage
colony-stimulating factor (GM-CSF), a decrease in the surface expression of CXCR4 on
myeloma cells compared to pre-mobilization BM specimens was observed (Gazitt and
145
Akay, 2004). To investigate the role of the CaR in the regulation of MM cell migration,
we performed CaR knockdown and overexpression using the lentiviral vectors generated
and assessed the migratory response of MM cell to SDF-1α in vitro. We discovered that
when the CaR was downregulated, there was a significant decrease in MM cell migration
towards specifically SDF-1α, while the vice versa was observed when the CaR was
overexpressed. However, these functional changes in MM cell migration when CaR
expression was altered did not correlate directly with cell surface expression of CXCR4,
which suggests that receptor expression alone is not an adequate parameter to predict the
functionality of the receptor. In fact, large variations were reported in CXCR4 expression
in MM cells, which were demonstrated to be inconsistent with the migratory response of
MM cells to SDF-1α (Möller et al., 2003). Since binding of chemokines to their receptors
causes a characteristic increase in cytosolic calcium that can be measured using an
intracellular calcium flux assay, it remains to be investigated whether the changes
observed in MM cell migration with CaR modulation are correlated with changes in
intracellular calcium mobilization.
In addition to providing an abundance of chemokines for the homing of MM cells, the
BM microenvironment also contains ECM components such as fibronectin and collagens,
which interact with a wide range of integrins expressed on MM cells. Integrins control
cell functions by binding to ECM molecules to transmit signals from the ECM to the
cells, as well as conveying intracellular signals to the extracellular environment through
changes in affinity for ECM molecules. Using immunohistochemistry, previous studies
146
have characterized that the BM ECM of MM patients contain fibronectin, collagen I,
collagen IV, laminin, and tenascin (Kibler et al., 1998; Tancred et al., 2009), which MM
cells can adhere to via cell surface integrins involving predominantly the α
4
, α
5
, α
v
, and
the β
1
subunits (Uchiyama et al., 1992; Van Riet et al., 1991; Pellat-Deceunynck et al.,
1995; Sanz-Rodriguez et al., 1999). Besides mediating bidirectional signaling,
interactions between integrins and ECM molecules can also confer drug resistance. A
previous study demonstrated that in drug-resistant malignant plasma cells, α
4
β
1
integrin,
or VLA-4, was overexpressed (Damiano et al., 1999). These cells were also shown to
have increased cell-ECM adhesion to fibronectin and enhanced drug resistance to
doxorubicin and melphalan (Damiano et al., 1999). The CaR was recently identified as a
key protein interaction partner with β
1
integrins in rat medullary thyroid carcinoma cells
(Tharmalingam et al., 2011), where functional coupling of the receptor to β
1
integrins
promoted cellular adhesion and migration in cancer cells. However, the role of the CaR in
modulating cell adhesion in MM cells was unclear. Here we demonstrated for the first
time using a genetic approach that the CaR is involved in the adhesion of MM cells to
fibronectin. It remains to be demonstrated if the CaR also mediates cell adhesion in MM
cells through functional coupling with β
1
integrins. Interestingly, the CaR does not appear
to mediate MM cell adhesion to collagen I. Fibronectin and collagen I were shown to be
expressed in the central marrow region as well as the endosteal region of the BM, but
fibronectin was more highly expressed than collagen I in bone sections from MM patients
with high level plasmacytosis (Tancred et al., 2009). The relevance of the spatial
147
localization of ECM molecules to the distinctive pattern in cell adhesion mediated by
CaR remains to be investigated.
Drug development is now focused on targeting MM cells in the BM microenvironment.
One pathologic characteristic triggered by the binding of MM cells to the BM stroma is
the upregulation of the ubiquitin proteasome cascade at both the gene transcript and
activity levels (McMillin et al., 2010). As a result, this makes the proteasome a favorable
target for treating MM. Bortezomib is a proteasome inhibitor that targets MM cell in the
BM microenvironment and was rapidly adapted into clinical practice for the treatment of
relapsed and refractory MM. The range of its therapeutic effects include abrogation of
tumor growth and survival, downregulation of adhesion molecules on cancer cells and the
BM, and induction of apoptosis in cancer cells and osteoclasts while promoting
osteoblast differentiation (Anderson, 2011; Chauhan et al., 2008; Mukherjee et al., 2008).
In the present study, we examined the effect of CaR modulation on MM cell sensitivity to
bortezomib treatment in vitro. We found that the CaR played a chemoprotective role in
MM cells, as CaR knockdown sensitized the cells to bortezomib treatment while CaR
overexpression protected the cells to bortezomib treatment. Mechanistically, we
demonstrated that the decrease in cell viability when CaR knockdown cells were treated
with bortezomib was due to an increase in apoptosis. Collectively, these results suggest
that the CaR can potentially be targeted to sensitize MM cells to bortezomib treatment,
thereby improving the clinical outcome of MM patients.
148
Multiple myeloma represents a malignant disease in which the non-malignant accessory
cellular and non-cellular components in the BM microenvironment are extensively
involved to propagate the pathologic phenomena of the disease. In order to investigate the
mechanisms responsible for the pathogenesis of MM, an in vivo model that recapitulates
the complexity of tumor-host cell interactions is necessary. Here, we generated the SCID-
hu model of MM by first implanting human fetal bone fragment subcutaneously into
SCID mice and then injecting MM cells directly into the human BM microenvironment.
This in vivo model is a unique model because both human hematopoietic cells and the
human hematopoietic microenvironment are engrafted in the mouse, enabling the study
of human cell-human microenvironment interactions in a xenogeneic model. We showed
that the human MM cells can be reproducibly propagated in the human BM
microenvironment in SCID-hu mice. More importantly, these human MM cells do not
migrate to the mouse BM, suggesting a preferential localization of human MM cells in
the human BM microenvironment and reinforcing the relevance of the SCID-hu model in
the study of MM. Our previous studies on the functional role of the CaR in normal
hematopoiesis unveiled the importance of the receptor in regulating the localization of
HSCs in the endosteal BM stem cell niche (Adams et al., 2006; Lam et al., 2011). Using
the SCID-hu model of MM, we can potentially examine the functional role of the CaR in
regulating the localization of myeloma cells in the BM microenvironment. In particular,
using bilateral implants, we can study the functional effect of CaR expression on the
metastasis of MM cells from one human fetal bone fragment to the other. Furthermore, it
has been reported that various cellular components of the human BM in the SCID-hu
149
model can be manipulated by exogenously providing human hematopoietic growth
factors (Kyoizumi et al., 1993). Taking this advantage of the SCID-hu model, we can
potentially modulate tumor-host cell interactions in the human BM microenvironment,
assess the pathophysiology of MM in response to these modulations, and validate the
efficacy of various chemotherapeutic agents in the treatment of MM.
150
Chapter 4: The Calcium-Sensing Receptor Specifies a Distinct
Cell Population in the Human Hematopoietic System
4.1 Abstract
Previous studies demonstrated the crucial role of the CaR in specifying murine HSC
lodgment in the endosteal BM niche following transplantation. By using pharmacologic
modulation to stimulate the CaR, we further identified the underlying mechanisms
involved in this process, including activation of the CXCR4 signaling pathway and
increased HSC-collagen I binding. In our current study, we wished to investigate the role
of the CaR on primitive human hematopoietic cells by first analyzing the expression of
CaR in human BM, UCB, MPB, and FL. We found that the percentage of CaR
+
cells in
the primitive human CD34
+
CD38
-
cell population was markedly different depending on
the tissue source, with the highest percentage found in UCB, followed by BM, FL, and
then MPB. Interestingly, our in vitro functional studies demonstrated that the CD34
+
CaR
-
cell population was superior in its ability to form CFU-GM and BFU-E colonies and had
an overall higher CFU-C frequency than the CD34
+
CaR
+
cell population. The
CD34
+
CaR
-
cell population also had a significantly higher CAFC frequency than the
CD34
+
CaR
+
cell population, indicating a higher primitive cell activity in vitro. Cell cycle
analysis revealed that the CD34
+
CD38
-
CaR
-
cell population was significantly more
quiescent compared to the CD34
+
CD38
-
CaR
+
cell population. However, when human FL
CD34
+
CaR
+
and CD34
+
CaR
-
cell populations were assessed for their engraftment
potential, both cell populations displayed comparable engraftment levels in NSG mouse
recipients. This study begins to provide a novel understanding both of the role of the CaR
151
in specifying a distinct stem cell population in the human hematopoietic system and the
functional differences between HSCs obtained from the BM, UCB, MPB, and FL. In
addition, the results of these studies may have implications in the development of HSC-
based therapies using various sources of HSCs.
152
4.2 Introduction
The intrinsic characteristic that defines a functional human HSC resides in its migration
and repopulation potential upon transplantation into pre-conditioned recipients. Despite
species-dependent differences, human and mouse BM share common architectural
structures that facilitate HSC migration and repopulation in the BM (Lapidot et al.,
2005). Studies on the functions of human HSCs are often hampered by a suitable
xenogeneic model or limited by the availability of human tissue sources, which can be
derived from the UCB, BM, MPB, or FL. Nonetheless, regulators involved in the
homing, lodgment, and engraftment of murine HSCs have also been identified to play
similar roles in human HSCs after transplantation.
Studies on the homing of human HSCs to the BM have identified the importance of the
CXCR4/SDF-1α axis as an important regulator. It was discovered that the chemokine
SDF-1α was expressed by immature human osteoblasts lining the endosteal region and in
the BM endothelium (Imai et al., 1999), specifically by endothelial cells lining the small
and large vessels, including peri-arterial regions as well as the blood capillaries of the
bone (Ponomaryov et al., 2000). In addition, SDF-1 expression could be induced in
NOD/SCID recipients by conditioning with DNA-damaging agents such as ionizing
irradiation, cyclophosphamide, and 5-fluorouracil, to enhance human CD34
+
cell
migration and repopulation potentials (Ponomaryov et al., 2000). Another study
demonstrated that human CD34
+
CD38
-/low
cells migrated to SDF-1 in vitro in a CXCR4-
153
dependent manner, and that treatment of human cells with antibodies to CXCR4
prevented engraftment (Peled et al., 1999).
Besides the CXCR4/SDF-1 axis, cell adhesion molecules also mediate the homing,
lodgment, and engraftment of transplanted human HSCs. Neutralizing antibodies against
the integrin VLA-4 blocks homing of human CD34
+
cells to the BM of fetal sheep
(Zanjani et al., 1999). VLA-4, VLA-5, and LFA-1 were demonstrated to be the
participating players in the homing of human CD34
+
cells to the BM and spleen of
NOD/SCID mice and thus essential for BM repopulation (Kollet et al., 2001; van der Loo
et al., 1998; Peled et al., 2000). Moreover, SDF-1 was shown to activate VLA-4, VLA-5,
and LFA-1 on human CD34
+
cells to support cell adhesion to fibronectin and
transendothelial migration (Peled et al., 2000).
Our previous studies have shown that the CaR expressed on murine HSCs regulates cell
lodgment and engraftment in the endosteal niche following transplantation (Adams et al.,
2006; Lam et al., 2011). Although the mechanisms of CaR regulation of murine HSC
functions have been identified, the role of the receptor in human HSC functions is still
unknown. Therefore, in this study, we isolated human HSCs from various hematopoietic
tissues based on the expression of the CaR and examined functional differences between
the CaR
+
and CaR
-
cell populations. Specifically, we performed immunophenotypic
analysis for the expression of cell surface CaR in the human BM, UCB, MPB, and FL
HSCs and measured hematopoietic stem and progenitor cell activity in vitro as well as the
154
engraftment potential in vivo. This study will allow us to begin to define the role of the
CaR in human HSCs and provide us a novel understanding on the regulation of human
HSC functions by the CaR.
155
4.3 Materials and Methods
4.3.1 Tissue Sources and Immunomagnetic Enrichment
Fresh UCB, BM, and MPB MNCs (~100x10
6
cells) from healthy donors were shipped in
a 50ml conical tube containing Dulbecco’s PBS with 0.5% BSA and 2mM EDTA with a
cold gel pack (AllCells, LLC, Emeryville, CA, USA). Cells were centrifuged at 400g for
5 minutes, then resuspended in 300µl of 1X PBS. Next, 100µl of FcR blocking reagent
and 100µl of CD34 microbeads were added (both from Miltenyi Biotec, Auburn, CA,
USA), then the cells were incubated at 4°C for 30 minutes. Meanwhile, a MACS MS
column with a 30µm cell strainer on top was placed in the magnetic MACS Separator (all
from Miltenyi Biotec), and 1ml of 1X PBS was applied to the column for rinsing. The
flow-through was discarded. When the incubation step was over, the cell suspension was
washed with 5ml of 1X PBS, centrifuged at 400g for 5 minutes, and resuspended in 1ml
of 1X PBS. The cell suspension was applied to the column, and the flow-through
containing unlabeled cells was collected. The column was then washed twice with 1ml of
1X PBS, and the flow through was also collected. The column was removed from the
separator and placed on a clean 15ml tube. Then, 2ml of 1X PBS was added to the
column, and the magnetically labeled cells were flushed out by firmly plunging the
plunger into the column. These cells contained the enriched containing the enriched
CD34
+
cell population. Frozen human FL (15-24 weeks old) CD34
+
cells (kind gifts from
Dr. Paula Cannon, USC Keck School of Medicine, Los Angeles, CA, USA) was quickly
156
thawed over a 37°C water bath and transferred to a 15ml tube with 13ml of 1X PBS. The
cells were then centrifuged down at 400g for 5 minutes to remove the supernatant.
4.3.2 Flow Cytometry
The anti-human CaR monoclonal antibody (clone: 5C10, ADD) was labeled using the
Zenon
®
Alexa Fluor
®
647 Mouse IgG
2a
Labeling Kit (Molecular Probes) according to
manufacturer’s instructions as follows. First, 1µg of the antibody was incubated with 5µl
of the Zenon
®
mouse IgG
2a
labeling reagent (Component A) for 5 minutes at room
temperature, protected from light. Next, 5µl of the blocking reagent (Component B) was
added to the reaction mixture and incubated for 5 minutes at room temperature, protected
from light. The antibody complexes were used within 30 minutes. The enriched human
CD34
+
cell population was resuspended in 200µl of 1X PBS, then stained with anti-
human FITC CD34 and PE-Cy7 CD38 antibodies (3µl each) (both from Becton
Dickinson), as well as the entire volume of the CaR antibody complexes (11µl) for 15
minutes on ice, protected from light. The stained cells were washed with 5ml of 1X PBS,
centrifuged down at 400g for 5 minutes, and resuspended in 0.5ml of 1X PBS for
immunophenotypic analysis and sorting using the FACSAria or Aria II flow cytometer.
4.3.3 CFU-C Assay
Sorted human CD34
+
CaR
-
and CD34
+
CaR
+
cells from the various tissue sources were
resuspended in 0.3ml of α-MEM, then added to 3ml of the MethoCult
®
H4435
methylcellulose medium (STEMCELL Technologies) to yield cell density of 750
157
cells/3.3ml in a 15ml tube. The MethoCult
®
GF
+
H4435 methylcellulose medium
contains FBS, BSA, 2-mercaptoethanol, rh SCF, rh G-CSF, rh granulocyte macrophage
colony-stimulating factor (GM-CSF), rh IL-3, rh IL-6, and rh erythropoietin. The tubes
containing the cells were then vortexed and let stand until all the bubbles had risen to the
top surface (approximately 5 minutes). A 3ml syringe attached to 18-gauge needle was
then used to plate out the cells in duplicates in a volume of 1.1ml per well (250
cells/well) in a 6-well plate pre-marked with gridlines (VWR International). Distilled
water was added to the empty wells in the middle of the plate. The plates were wrapped
in a plastic bag and incubated at 37°C in a humidified, 5% CO
2
atmosphere for 14 days
before they were scored for the number of colony forming unit-granulocyte macrophage
(CFU-GM) and BFU-E colonies.
4.3.4 CAFC assay
Sorted human CD34
+
CaR
-
and CD34
+
CaR
+
cells from the various tissue sources were
seeded in serial dilutions onto a confluent layer of OP9 stromal cells (irradiated at 35Gy
and seeded a day before) in a cell culture treated 96-well plate.
CD34
+
CaR
-
: 60, 125, 250, 500, 1000
CD34
+
CaR
+
: 60, 125, 250, 500
The co-cultures were maintained in α-MEM supplemented with 10% FBS and P/S
(Mediatech Inc.) in a humidified atmosphere at 37°C in a humidified, 5% CO
2
158
atmosphere. Half of the culture medium was replaced with fresh medium every week.
The presence of CAFCs underneath the OP9 stromal layer was scored on week 5, and the
frequency of CAFCs was calculated using the L-Calc software.
4.3.4 Cell Cycle Analysis
MNCs from the various sources were stained with 10µg/ml Hoechst 33342 for DNA and
25µg/ml of verapamil at 37°C for 45 minutes, then stained with anti-human FITC CD34,
PE-Cy7 CD38, and Zenon
®
Alexa Fluor
®
647-labeled CaR antibodies as described
above. The stained cells were resuspended and fixed in 10% neutral buffered formalin
and incubated at 4°C overnight, wrapped in foil. To stain for RNA content, pyronin Y
was added to the cells at a final concentration of 0.75µg/ml and incubated at 4°C for 30
minutes wrapped in foil. Cell cycle status was examined using the LSR II flow cytometer.
Both pyronin Y and Hoechst were analyzed on a linear scale.
4.3.5 Engraftment
Immunodeficient NSG mice were housed in sterilized microisolator cages and received
autoclaved food and water ad libitum, in accordance with the University of Southern
California IACUC guidelines. Newborn NSG pups (P1) were used as subjects for the
engraftment studies. First, the pups were irradiated at 1.5Gy in a clean chamber lined
with paper towels approximately 4 hours prior to transplantation. Following irradiation,
the pups were placed with the nursing mother as soon as possible. Human FL CD34
+
CaR
-
and CD34
+
CaR
+
cells were sorted as described above. Next, the sorted cells were
159
resuspended at ~2,500 cells/25µl, and each pup was injected with 25µl of cells
intrahepatically using a clean Hamilton syringe as follows. First, the pup was grasped by
the shoulders and back of the neck with the thumb and forefinger, with the belly side up.
With the free hand, the syringe was held in a way that the index finger was on the end of
the syringe plunger. The needle was inserted subcutaneously at a 10° angle above the
sternum and continued towards the liver, which was visible under the right costal margin.
The needle was carefully inserted into the liver, and the cells were slowly injected. The
syringe was washed three times with 1X PBS, three times with ethanol, then three times
with 1X PBS again before the next injection. Following transplantation, the pups were
returned to the nursing mother as soon as possible. After 3 weeks, the pups were weaned.
For analysis of human cell engraftment, approximately 50µl of blood was collected into a
microcentrifuge tube using retro-orbital sampling. The red blood cells were allowed to
settle to the bottom of the tube, and the leukocyte-containing upper fraction was
transferred to fresh polystyrene flow tubes. Next, enough 1X PBS was added to the
sample to bring the total volume up to 100µl. Then, the sample was stained with anti-
human PE-Cy5 CD45, FITC CD19, PE CD33 (all from Becton Dickinson), and APC
CD3 (eBioscience) antibodies on ice for 15 minutes, protected from light. After staining,
3ml of 1X PBS was used to wash the cells, and the cells were centrifuged at 400g for 5
minutes. The red blood cells were lysed in 1ml of 1X FACS lysing solution for 1 minute
at room temperature. The samples were then fixed in 300µl of 10% neutral buffered
formalin and analyzed by flow cytometry using the LSR II cell flow cytometer. The data
were analyzed using FlowJo.
160
4.4 Results
4.4.1 The expression level of cell surface CaR is distinctive among the human
hematopoietic tissues.
In the research and clinical setting, FL, UCB, BM, and MPB have been used as sources
of HSCs for either xenogeneic transplantation studies or for therapeutic applications.
However, differences in the time for hematopoietic recovery with the use of each source
have been documented (Bensinger et al., 1995; Körbling et al., 1995; Smith and Wagner,
2009), which could be attributed to differences in the functional properties of the HSCs.
In order to avoid any biases on the selection of tissue sources for our study which might
potentially interfere with the analysis of HSC functions, we decided to first examine the
cell surface expression of the CaR in all four human HSC tissue sources.
Immunophenotypic analysis revealed that the CaR was expressed at different levels on
the surface of primitive CD34
+
CD38
-
cells, with the highest expression level in UCB and
the lowest in MPB (Figure 4.1).
161
Figure 4.1 Expression levels of cell surface CaR in human BM, UCB, MPB, and FL.
Hematopoietic cells from the human BM, UCB, MPB, and FL were stained with anti-human
FITC CD34, anti-human PE-Cy7 CD38, and anti-human Zenon
®
-labeled CaR antibodies. The
stained cells were analyzed for the expression of the CaR using flow cytometry. Representative
flow plots are shown on the left, demonstrating the gating schemes used. The percentages of
CaR
+
cells within the primitive CD34
+
CD38
-
cell population from the different cell sources are
shown on the right (n=1-3 from 1 to 3 independent experiments; error bars represent s.e.m.).
4.4.2 The CD34
+
CaR
-
cell population has superior hematopoietic progenitor activity
in vitro compared to the CD34
+
CaR
+
cell population in human hematopoietic
tissues.
We next investigated hematopoietic progenitor frequency and differentiation potential in
the CD34
+
CaR
-
and CD34
+
CaR
+
cell populations from the different tissue sources by
assessing their ability to differentiate into BFU-E and CFU-GM colonies in vitro.
Interestingly, the CD34
+
CaR
-
cell population had higher BFU-E and CFU-GM
frequencies than the CD34
+
CaR
+
cell population in all four tissue sources (Figure 4.2),
suggesting that cell surface expression of CaR marks a cell population with less
progenitor activity. Furthermore, FL had the lowest CFU-C frequency compared to BM,
UCB, and MPB, suggesting a cell intrinsic difference in FL progenitor activity.
0
5
10
15
20
25
30
35
40
Percentage (%)
BM UCB MPB FL
162
Figure 4.2 Hematopoietic progenitor activities in human BM, UCB, MPB, and FL as
measured by CFU-C frequency. CD34
+
CaR
-
and CD34
+
CaR
+
cells were seeded in MethoCult
®
GF
+
H4435 methylcellulose medium and cultured for two weeks at 37°C in a humidified, 5% CO
2
atmosphere. The number of BFU-E and CFU-GM colonies formed after the two-week culture
period was scored (n=4-6 from 1 to 3 independent experiments; error bars represent s.e.m.).
4.4.3 The CD34
+
CaR
-
cell population has higher primitive hematopoietic cell activity
in vitro than the CD34
+
CaR
+
cell population in human hematopoietic tissues.
To address the question of whether a difference in primitive hematopoietic cell activity in
vitro exists in the CD34
+
CaR
-
and CD34
+
CaR
+
cell populations, the functional CAFC
assay was performed. Compared to the CD34
+
CaR
+
cell population, the CD34
+
CaR
-
cell
population had a higher week 5 CAFC frequency in vitro (Figure 4.3), translating into
higher primitive hematopoietic cell activity. This was observed for all four human
0
5
10
15
20
25
30
35
40
CD34+CaR- CD34+CaR+
CFU-Cs/250 cells
FL
BFU-E
CFU-GM
0
5
10
15
20
25
30
35
40
CD34+CaR- CD34+CaR+
CFU-Cs/250 cells
BM
0
5
10
15
20
25
30
35
40
CD34+CaR- CD34+CaR+
CFU-Cs/250 cells
UCB
BFU-E
CFU-GM
0
5
10
15
20
25
30
35
40
CD34+CaR- CD34+CaR+
CFU-Cs/250 cells
MPB
163
hematopoietic tissues. Interestingly, BM CD34
+
CaR
-
cells had the highest overall CAFC
frequency compared to the other human hematopoetic tissues (Figure 4.3), suggesting
that the BM CD34
+
cell population has the highest primitive hematopoietic cell activity.
Figure 4.3 Primitive hematopoietic cell activities in human BM, UCB, MPB, and FL.
CD34
+
CaR
-
and CD34
+
CaR
+
cells were seeded on a confluent stromal layer of supportive OP9
cells in serial dilutions and cultured at 37°C in a humidified, 5% CO
2
atmosphere. The presence
of cobblestone areas was scored on Week 5 (n=3-6 from 2 or 3 independent experiments; error
bars represent s.e.m.).
4.4.4 Cell cycle analysis reveals a loss of quiescence in the primitive CD34
+
CD38
-
CaR
+
cell population.
To determine if cell surface expression of the CaR in the human hematopoietic tissues
specifies a distinct cell population with an altered cell cycle profile, cell cycle analysis
was performed. The results revealed that the CD34
+
CD38
-
CaR
-
cell population was
164
largely residing in the G
0
phase of the cell cycle in human BM, UCB, and MPB, while
the majority of the CD34
+
CD38
-
CaR
+
cell population were actively cycling (Figure 4.4).
Interestingly, there was no difference in the percentage of primitive CD34
+
CD38
-
cells in
the G
0
phase based on the expression of the CaR, suggesting an intrinsic difference in the
regulation of cell cycle entry in human FL HSCs comparing to other human
hematopoietic tissues.
Figure 4.4 Cell cycle analysis based on the expression of cell surface CaR in the primitive
CD34
+
CD38
-
cell population. Representative flow cytometry plots depicting the gating schemes
for each phase of the cell cycle are shown on the left. The percentage of CD34
+
CD38
-
CaR
-
and
CD34
+
CD38
-
CaR
+
cells in the G
0
phase is plotted on the right (n=1-3 from 1 to 3 independent
experiments; error bars represent s.e.m.).
4.4.5 Human FL CD34
+
CaR
-
and CD34
+
CaR
+
cells display similar engraftment
potential in NSG mouse recipients.
A previous study demonstrated that human HSCs exhibiting cell cycle entry by
transitioning from the G
0
to G
1
phase of the cell cycle have an impaired stem cell
function as demarcated by their decreased ability to engraft in xenogeneic recipients
0
20
40
60
80
100
% G
0
BM UCB MPB FL
CD34
+
CD38
-
CaR
-
CD34
+
CD38
-
CaR
+
Hoechst 33342
Pyronin Y
G
1
G
0
S/G
2
/M
G
0
S/G
2
/M G
1
CD34
+
CD38
-
CaR
+
CD34
+
CD38
-
CaR
-
165
(Gothot et al., 1998). To investigate any difference in engraftment potential, human FL
CD34
+
CaR
-
and CD34
+
CaR
+
cells were transplanted into irradiated NSG mouse
recipients, and the contribution of human cells to hematopoiesis was monitored for up to
13 weeks. Surprisingly, despite the functional difference and stem and progenitor activity
in vitro, human FL CD34
+
CaR
-
and CD34
+
CaR
+
cells displayed similar engraftment
potentials, as depicted by immunophenotypic analysis of the peripheral blood as well as
the BM (Figure 4.5A, B). Furthermore, the cells were able to contribute to multilineage
engraftment in the NSG mouse recipients (Figure 4.5C).
Figure 4.5 The engraftment potential of human FL CD34
+
CaR
-
and CD34
+
CaR
+
cell
populations in NSG mouse recipients. (A) The percentages of human CD45
+
cells in the
peripheral blood of NSG mouse recipients were examined on weeks 8, 12, and 13 after
transplantation. (B) The NSG mouse recipients were sacrificed on week 13, and the percentages
of human CD45
+
cells in the BM were examined. (C) The contribution of human CD45
+
cells in
the reconstitution of B lymphoid, T lymphoid, and myeloid lineages were measured in the BM on
week 13 (n=3-4; error bars represent s.e.m.).
A
166
Figure 4.5 Continued
B
C
167
4.5 Discussion
In our current study, we provided a novel characterization of distinct human stem and
progenitor cell populations based on the cell surface expression of the CaR. In addition,
we provided a direct comparison of the functional properties of the four most widely
investigated sources of human HSCs, including the BM, UCB, MPB, and FL, which was
lacking in previous studies. We first showed that the CaR was expressed at distinctive
levels on the cell surface of primitive CD34
+
CD38
-
cells in the BM, UCB, MPB, and FL,
with the highest expression in UCB and the lowest expression in MPB. The difference in
expression levels might be ontogeny-related, as these human hematopoietic tissues were
obtained from adults and fetuses during development. However, it is not known precisely
why primitive human HSCs from the UCB have the highest level of cell surface CaR
expression and the functional implications that this characteristic has on HSCs derived
from the UCB. On the other hand, we discovered that the CaR was expressed at very
minimal levels on the cell surface of primitive human HSCs derived the MPB. Our
previous studies on murine HSCs have identified the importance of the CaR in the
lodgment or localization of HSCs in the endosteal stem cell niche (Adams et al., 2006;
Lam et al., 2011), and that mice deficient for the CaR displayed high level of circulating
HSCs (Adams et al., 2006). Based on this, HSCs that have been mobilized from the stem
cell niche to circulate in the peripheral blood should have minimal cell surface CaR
expression, which is in accordance with our current finding.
168
Interestingly, we discovered that the CD34
+
CaR
-
cell population had higher
hematopoietic stem and progenitor cell activity in vitro than the CD34
+
CaR
+
cell
population, as exemplified by higher CFU-C and CAFC frequencies. This could be
explained by the in vitro functional assays used for measuring hematopoietic stem and
progenitor activity, which specifically assess myeloid activity in those cells. Therefore, if
the CD34
+
CaR
+
cell population was lymphoid biased, the activity would not be evident in
the in vitro functional assays performed. It is also interesting to note that the in vitro
functional differences between the two cell populations were particularly significant in
HSCs obtained from the BM, UCB, and MPB than the FL, providing supportive evidence
for intrinsic functional differences between HSCs derived from different hematopoietic
tissues. Developmental changes in CD34 expression have been documented in a previous
study on murine HSCs, where in fetal, neonatal, and younger adult hematopoietic tissues,
most of the Lin
-
c-Kit
+
HSCs express CD34 (Matsuoka et al., 2001). Whether the
difference observed in hematopoietic stem and progenitor cell activity in human BM,
UCB, MPB, and the FL can be attributed to developmental changes is still unknown.
Using cell cycle analysis, we also discovered a loss of quiescence in human HSCs with
cell surface CaR expression. A previous study on identifying the link between cell cycle
and stem cell function has reported distinct hematopoietic capabilities in CD34
+
cells
residing in the G
0
phase compared to those in the G
1
phase, where primitive LTC-IC
activity was higher in CD34
+
cells isolated in G
0
than those isolated in G
1
(Gothot et al.,
1997). Taken together, the impairment in vitro hematopoietic stem and progenitor cell
activity in the CD34
+
CaR
+
cell population could be explained by a loss of quiescence in
169
those cells. However, the underlying mechanism that leads to the impairment in
hematopoietic stem and progenitor cell activity remains to be elucidated.
At present, the NSG mouse model is considered a superior model for analysis of human
stem cell development and function because it contains a multitude of immunological
dysfunctions, including impaired cytokine production capability and functional
incompetence of T, B, and NK cells, which may lead to the high engraftment levels in the
recipients (Ito et al., 2002). Despite significant functional differences in hematopoietic
stem and progenitor cell activity observed in vitro, human FL CD34
+
CaR
-
and
CD34
+
CaR
+
cells demonstrated comparable engraftment potentials in NSG mice. In
addition, both cell populations exhibited similar multilineage reconstitution potential.
However, because the total engraftment level was very low (~1% in the BM of NSG
recipients on week 13), it is inconclusive whether there is a true functional difference
between CD34
+
CaR
-
and CD34
+
CaR
+
cells in their ability to contribute to hematopoiesis
in NSG recipients. Regardless, this study presented the first investigation of the
functional role of the CaR on human HSCs and providing a direct comparison of the
functional differences between the four most widely studied or clinically used human
hematopoietic tissue sources: BM, UCB, MPB, and FL. Moreover, these findings will be
useful in the investigation and development of HSC-directed therapies using various
hematopoietic tissue sources.
170
Chapter 5: Concluding Remarks and Future Perspectives
5.1 Concluding Remarks
The capacity to sense extracellular changes in the environment, translate these changes
into intracellular biological responses, and adapt to these changes accordingly is of
paramount importance for the maintenance of proper cellular functions in every living
organism. With the development of membrane-bound sensing systems through evolution,
primarily the G protein-coupled receptor superfamily, adaptation to changes in
environmental cues is possible. The CaR was the first ion-sensing receptor identified on
the surface of various cell types in humans that is able to bind to Ca
2+
ions to decode its
signal (Riccardi et al., 2009). Besides its key role in extracellular calcium homeostasis,
the CaR is also an integrator of multiple environmental cues for the regulation of
numerous cellular processes. One process of particular interest to us is the regulation of
HSC biology in the BM microenvironment. Under homeostatic conditions in the BM,
specifically in the endosteal region, active bone remodeling mediated by osteoblasts and
osteoclasts releases Ca
2+
ions into the vicinity. In fact, the concentration of extracellular
Ca
2+
ions can reach as high as 40mM near bone resorbing osteoclasts in the endosteal
region (Silver et al., 1998). HSCs residing in the endosteal niche express the CaR, which
allows them to sense changes in extracellular calcium concentrations. It was
demonstrated in a previous study using CaR
-/-
mice that the CaR mediates lodgment and
localization of HSCs in the BM endosteal niche, with implications that therapeutic
targeting of the CaR could enhance the engraftment outcome of HSCs in the
171
transplantation setting (Adams et al., 2006). The first study presented here emphasized
the potential of using the clinically approved calcimimetic, Cinacalcet, to
pharmacologically stimulate the CaR to improve homing, lodgment, and engraftment of
transplanted HSCs (Chapter 2). In addition, the study reported a novel signaling
connection between the CaR and CXCR4, with intracellular Ca
2+
ions as the candidate
for secondary messenger (Chapter 2). CXCR4 is perhaps the most well studied regulator
of hematopoietic cell retention in the BM (Ma et al., 1999). The findings from the present
study suggest that the concerted interplay of the CaR and CXCR4 signaling pathways
acts to promote HSC localization in the endosteal BM niche by enhancing HSC-niche
interactions.
The BM microenvironment supports the pathophysiology of bone metastatic cancers,
such as breast cancer and prostate cancer, and the CaR has been documented to play an
oncogenic role by promoting tumor growth (Mihai et al., 2006; Liao et al., 2006). A
hematological malignancy that involves devastating bone destructions driven by an
imbalance of osteoblast-osteoclast activity is MM. In MM, bone lesions are caused by
overly active bone resorbing osteoclasts, which release a massive amount of Ca
2+
ions
from the bone into the BM endosteal region. MM cells were shown to specifically engraft
in the BM endosteal region and form a complex with osteoblasts and osteoclasts
(Iriuchishima et al., 2012). The functional properties of MM cells were demonstrated to
be modulated through the CaR to contribute to the pathophysiology of MM, including
increased cell proliferation and expansion of myeloma cell mass (Yamagutchi et al.,
172
2002). However, because interactions of the MM cells with the BM microenvironment
are known to be important in the pathophysiology of the malignancy, and the underlying
mechanisms behind CaR-mediated myeloma cell growth were largely lacking in the
previous studies, we specifically investigated interactions between MM cells and the BM
microenvironment as mediated by the CaR and generated an in vivo mouse model for the
study of the localization and metastasis of human MM cells in the BM
microenvironment. The findings presented here in the study suggest that the CaR
stimulates cell cycle entry to drive cell proliferation. In addition, the mechanisms behind
CaR regulation of MM pathophysiology involved adhesive and chemotactic interactions
between MM cells and the BM microenvironment (Chapter 3). More importantly, our
study provided the first evidence that the CaR plays a chemoprotective role in MM cells,
and that targeting CaR expression can sensitize MM cells to chemotherapeutic
intervention (Chapter 3). Collectively, findings from the present study further support the
importance of understanding and identifying the interplay between cancer cells and the
supportive BM microenvironment in developing new chemotherapeutic agents for bone
metastatic cancers.
The ultimate goal is to be able to apply what we have learned in regards to the regulation
of hematopoietic cell functions by the CaR, such as lodgment and localization, into
clinical uses. However, before that can be achieved, the impending question that also
needs to be addressed is the regulation of HSC functions by the CaR in the human
hematopoietic system. In the clinical setting, there are currently three different sources of
173
HSCs for hematopoietic stem cell transplantation for the treatment of various
hematological disorders: BM, UCB, and MPB (Nishino et al., 2012). Each source is
endowed with certain clinical advantages over another that can influence the outcome of
the transplantation. Besides these three sources of HSCs, the human FL is also widely
used as a source of HSCs in the research setting. Although the CaR has been shown to be
a critical player for mediating HSC-niche interactions in mice, its role in mediating
human hematopoietic stem and progenitor activity is lacking. The study presented here
provides the first functional evidence as well as a direct comparison for the regulation of
hematopoietic stem and progenitor cell activity by the CaR in human BM, UCB, MPB,
and FL. Findings from the study support the existence of intrinsic functional differences
between HSCs derived from human BM, UCB, MPB, and FL, as there are apparent
functional differences in hematopoietic stem and progenitor activity in vitro among the
tissues (Chapter 4). Although the CaR specifies a less quiescent primitive human HSC
population with impaired hematopoietic stem and progenitor activity in vitro, the
multilineage repopulation potential of human hematopoietic stem and progenitor cells in
NSG recipients is not abrogated by the phenotypic expression of the CaR (Chapter 4).
This raises the question of the real clinical usefulness of human hematopoietic assays, as
few studies have attempted to correlate the numbers of CFU-Cs or CAFCs to the outcome
of clinical transplantation (Breems et al., 1996). Nonetheless, these findings will be
useful in the investigation and development of HSC-directed therapies using various
hematopoietic tissue sources.
174
5.2 Future Perspectives
The studies presented in this thesis on CaR-mediated regulation of hematopoietic cell
interactions with the BM microenvironment may lead to additional insights on the
following areas of stem cell research: 1) The development of hematopoiesis, 2)
Pharmacologic modulation of HSCs and the stem cell niche, and 3) Alternative
approaches to enhance HSC transplantation.
5.2.1 The Development of Hematopoiesis
During fetal development, hematopoietic stem and progenitor cells follow an established
pattern of migration from the FL to the fetal BM, where the colonization of the fetal BM
by hematopoietic progenitor cells precedes that of long-term HSCs, which occurs at
E17.5 in mice (Christensen et al., 2004). A recent study using multiplex single cell
quantitative RT-PCR analysis on genes involved in HSC migration revealed distinct
patterns of gene expression in the primitive hematopoietic cell population, defined by the
Lin
-/low
Sca-1
+
c-Kit
+
CD150
+
CD48
-
CD41
-
phenotype, during migration from the FL to the
fetal BM (Ciriza et al., 2012). In particular, the study showed that CaR expression is
upregulated specifically in E17.5 fetal BM LT-HSCs compared to E14.5 FL, E17.5 FL,
and 4-week old adult BM (Ciriza et al., 2012). Since the CaR has been demonstrated to
be important for the localization of HSCs in the BM endosteal niche (Adams et al., 2006;
Lam et al., 2011), the upregulation of CaR expression in E17.5 fetal BM could suggest
the following. First, initiation of bone ossification at E17.5 during fetal development
creates a microenvironment suitable for the maintenance of LT-HSCs, and LT-HSCs are
175
able to sense this change and migrate towards the microenvironment via the CaR.
Second, the upregulation of CaR expression aids in guiding LT-HSCs in the blood
circulation to migrate to the endosteal niche. Finally, fetal BM LT-HSCs are retained in
the microenvironment via the orchestrated actions mediated by the CaR and cell adhesion
molecules, such as VLA-4 and VLA-5, which also facilitate CXCR4/SDF-1α
engagement. As the BM develops further and reaches adulthood, the expression of CaR
returns to the initial level to allow maintenance of hematopoiesis. Although there is the
unresolved question of how these signals are intricately regulated during the
development, the findings presented here and in previous studies on the CaR nonetheless
point to the importance of the capacity to sense and adapt to changes in the environment
both during fetal development and adult life.
The aforementioned studies highlight the ongoing question of how FL hematopoiesis is
different from BM hematopoiesis, especially on the functional properties of HSCs
derived from the FL and the BM. The CaR was first demonstrated to regulate the
lodgment of HSCs using FL CaR
-/-
HSCs injected into adult mouse recipients, since
beyond that age CaR
-/-
mice become hypercalcemic and die by the age of 7-10 days
(Adams et al., 2006). However, intrinsic differences between FL HSCs and BM HSCs
have been documented, such as increased cell cycling and engraftment potential in FL
HSCs compared to BM HSCs (Fleming et al., 1993; Micklem et al., 1972; Morrison et
al., 1995; Rebel et al., 1996). Therefore, BM HSCs derived from adult mice deficient in
the CaR may be necessary to complete our understanding on the role of the CaR in HSC
176
lodgment in the adult animal, as these cells have survived through multiple
environmental changes during the development of hematopoiesis. Continuing on the
discussion of FL and BM HSCs, another relevant question to address is whether intrinsic
differences exist in the homing and lodgment potential between normal BM and FL
HSCs as well as CaR
-/-
FL and BM HSCs in a competitive setting. In order to address
these questions, a conditional CaR knockout mouse model is needed to circumvent the
problem of early death in a straight, generalized CaR knockout mouse model. In fact,
tissue-specific deletion of the CaR in the parathyroid gland, bone, and chondrocytes have
been documented (Chang et al., 2008). Therefore, it is possible generate a conditional
knockout mouse with CaR deleted specifically in hematopoietic tissues, for example,
using Mx1-Cre mediated deletion. This conditional CaR knockout model will be
beneficial in defining various functions of the CaR in both fetal and adult hematopoiesis,
and also providing additional insights into CaR-mediated HSC-niche interactions.
5.2.2 Pharmacologic Modulation of HSCs and the Stem Cell Niche
The stem cell niche, proposed by Schofield in 1978, describes an anatomical environment
that is in close association with a true stem cell to maintain the capacity of the stem cell
to undergo self-renewing divisions (Schofield, 1978). Although the identification of stem
cell niche components in mammalian systems are challenging due to their complex
anatomical structure and the rarity of stem cells, extensive progress has been made in the
last decade. Studies have demonstrated that HSCs are located near the bone surface (Lord
et al., 1975, Lambertsen and Weiss, 1984; Nilsson et al., 2001), and that osteoblasts are a
177
critical component of the stem cell niche for HSC maintenance (Calvi et al., 2003; Zhang
et al., 2003; Visnjic et al., 2004). Although osteoclasts have been suggested to play a
dispensable role in the maintenance of HSCs using three osteopetrotic mouse models
(op/op, c-Fos-deficient, and RANK-L-deficient mice) (Miyamoto et al., 2011), the
importance of the dynamic bone remodeling that releases Ca
2+
ions into the endosteal
region cannot be ignored as it could affect HSC-niche interactions mediated by the CaR
expressed on HSCs. Recently, a study that examined hematopoiesis on mice treated with
bisphosphonates, an anti-resorptive agent that inhibits osteoclasts and affects bone
remodeling, reported a decrease in proportion and absolute number of primitive
hematopoietic cells in the BM (Lymperi et al., 2011). This finding suggests that
osteoclasts participate in the regulation of the HSC niche through interactions with
osteoblasts to change the physical conformation of the BM cavity, which potentially
affects available niche space. Since the CaR anchors HSCs in the endosteal region of the
BM where the extracellular calcium concentration is the highest, this raises the possibility
of modulating osteoblast-osteoclast interactions in the niche to alter the bone remodeling
process, which would in turn affect the amount of Ca
2+
ions being released in the
endosteal region, disturbing the homeostatic balance. This could potentially either
enhance HSC lodgment in the transplantation setting or mobilize HSCs away from the
niche for therapeutic applications.
The first study presented in this thesis also demonstrated that ex vivo pharmacologic
modulation of the CaR expressed on HSCs by a calcimimetic can enhance HSC homing,
178
lodgment, and engraftment in the BM (Chapter 2). In this regard, an interesting question
to ask is whether these stepwise processes of HSC journey from the peripheral blood
circulation to the stem cell niche could be disrupted with pharmacologic modulation
using a calcilytic that blocks the CaR. We could potentially apply the knowledge
gathered from the studies using CaR modulators to alter the localization of cancer cells in
bone metastatic malignancies. The BM microenvironment was proposed many years ago
to be a fertile soil to support cancer cell growth (Paget, 1889), owing to the fact that it
contains numerous growth factors and provides proliferative cues for the cancer cells via
cell-cell or cell-ECM interactions. For instance, in MM, interactions between the
myeloma cells and osteoclasts in the BM microenvironment contribute to the
pathophysiology of the disease. In addition to promoting cell growth and survival,
interactions between the myeloma cells and the BM microenvironment also contribute to
resistance to chemotherapy. A previous study revealed that myeloma cells overexpressing
VLA-4 have increased cell-ECM adhesion to fibronectin and enhanced drug resistance to
doxorubicin and melphalan (Damiano et al., 1999). Taken together, if modulating the
CaR on myeloma cells can alter their localization in the BM microenvironment,
therapeutic targeting of the CaR using allosteric modulators can be envisioned as an
effective strategy for inhibiting the pathophysiology of myeloma cells in MM patients.
One technical drawback of tracking HSCs following transplantation into a recipient is the
lack of stable cell labeling technologies that can sustain post-acquisition tissue processing
without a loss in signal. Current studies in the laboratory are focused on overcoming this
179
technical challenge, and efforts have been made to revolutionize the traditional
fluorescent cell labeling approach by using nanoprobes developed based on the principle
of second harmonic generation (SHG). SHG nanoprobes have multiple photophysical
advantages that make them a perfect tool for cell labeling, a few of which include
reduced phototoxicity and photobleaching and enhanced signal-to-noise ratio (Pantazis et
al., 2010). Recently, it has been demonstrated that these SHG nanoprobes can be used to
label BM MNCs and allow visualization of the labeled cells in bone sections following
standard histochemical procedures (Rashidi, 2011). With the advancement of SHG
nanoprobes for labeling HSCs and visualizing them in their niche following
transplantation, it will be technically feasible to identify the cellular and non-cellular
components that are involved in HSC lodgment and localization after ex vivo treatment
with a CaR modulator. Direct visualization of the effects of various genetic mutations on
stem cell number and function in lower organisms such as the Drosophila has identified
signaling pathways involved in stem cell niche biology (Yamashita et al., 2003; Xie and
Spradling, 2000). Therefore, findings from this investigation will provide new insights on
the interacting partners of HSCs that may be involved in facilitating hematopoietic
reconstitution by donor HSCs in the recipient, and also aid in identifying the location of
all HSC niches through direct visualization.
5.2.3 Alternative Approaches to Enhance HSC Transplantation
With the increased availability of cord blood banks and the relative ease of collecting
HSCs from peripheral blood, the annual numbers of hematopoietic stem cell
180
transplantation have dramatically increased during the last decades (Appelbaum, 2007;
Gratwohl et al., 2010). However, broad application of HSC transplants to most recipients
is still limited by the low HSC number recovered from grafts (Takizawa et al., 2011). To
solve this problem, strategies to expand HSCs ex vivo have been pursued. These
strategies mainly include manipulation of intrinsic factors or supplementation of extrinsic
growth factors to achieve HSC expansion. One notable study that reported significant ex
vivo HSC expansion was retrovirus-mediated introduction of the homeobox B4 (HoxB4)
gene in HSCs, which permitted about 40-fold expansion in vitro (Antonchuk et al., 2002)
and 1000-fold expansion in vivo (Sauvageau et al., 1995). However, virus-mediated gene
transfer by itself poses the risk of insertional mutagenesis, such as activation of potential
oncogenes (Baum et al., 2004). Therefore, it would be preferable to avoid manipulation
of the HSC genome and use alternative methods such as the addition of extrinsic factors.
One major breakthrough was the use of Delta-1 protein fused with the Fc portion of
human immunoglobulin (Delta-1 Fc) to significantly increase (100-fold) the number of
human CD34
+
cord blood progenitors capable of enhanced repopulation in the BM of
NOD/SCID mice (Delaney et al., 2010; Ohishi et al., 2002). This study has been carried
forward in a phase I clinical trial, and the results demonstrated substantial reduction in
the time to neutrophil recovery (Delaney et al., 2010). However, long-term risks and
benefits remain to be evaluated. The findings presented in this thesis suggest that it is
possible to stimulate the CaR on HSCs using a clinically approved calcimimetic to
enhance homing, lodgment, and engraftment in the recipient (Chapter 2). In addition, we
have shown that the treatment imposes a change in the HSC on the functional level only,
181
instead of altering the expression level of the CaR or other molecules involved in the
migration of HSC to the BM (Chapter 2). Therefore, this strategy could allow the
functional enhancement of HSC engraftment in the transplantation setting through
improved lodgment efficiency, thereby circumventing the need to expand HSC number
for transplantation.
Another possible approach to enhance HSC transplantation is to pre-condition the patient
receiving the transplant with a calcimimetic for the CaR. In the clinical setting, HSCs are
transplanted into the patient by simple infusion of the graft into the venous system
(Appelbaum, 2007), and the transplanted HSCs migrate through the blood circulation to
home to the BM stem cell niche that supports HSC engraftment and reconstitution of the
hematopoietic system. By pre-conditioning the patient with a calcimimetic for the CaR,
HSCs transplanted into the patient may be activated by the calcimimetic present in the
blood circulation. Subsequently, the activated HSCs in the blood circulation may become
more efficient in homing to the BM and lodging in the BM stem cell niche to give rise to
multilineage hematopoietic reconstitution in the patient. On the contrary, it may also be
possible to target the CaR as a strategy to mobilize HSCs away from the BM stem cell
niche to yield MPB grafts containing high HSC numbers. In comparison to stem cell
transplant recipients that have been subjected to BM grafts, MPB grafts containing high
doses of HSCs give rise to faster hematopoietic recovery (Bensinger et al., 1995;
Körbling et al., 1995). In the clinical setting, administration of G-CSF is widely used to
mobilize HSCs into the peripheral blood, but studies have reported the side effects of this
182
approach, which includes depletion of endosteal macrophages that support osteoblast
function and inhibition of niche function through the loss of endosteal osteoblasts
required for HSC retention and self-renewal (Winkler et al., 2010; Christopher et al.,
2011). In addition, some patients and donors fail to mobilize adequate numbers of HSCs
with this standard approach. Therefore, enhanced mobilizing strategies have been
developed, such as the small synthetic reversible inhibitor of SDF-1α/CXCR4 binding,
AMD3100. The administration of both AMD3100 and G-CSF was proven to be superior
to G-CSF alone in the number of CD34
+
cells mobilized into the circulation (Flomenberg
et al., 2005). Furthermore, in two Phase III clinical studies, it was reported that the
addition of AMD3100 to G-CSF is generally safe and well tolerated in patients with non-
Hodgkin’s lymphoma (DiPersio et al., 2009; Micallef et al., 2009). Since transgenic mice
deficient in the CaR display increased number of circulating hematopoietic stem and
progenitor cells (Adams et al., 2006), the use of CaR antagonists may be envisioned as an
alternative strategy to mobilize HSCs into the blood circulation. This strategy could
potentially produce several clinical benefits, including reduced numbers of apheresis
procedures required and increased CD34
+
cell doses for transplantation. In addition,
because CaR antagonists are thought to target HSCs directly, the side effects associated
with G-CSF administration could be avoided. The feasibility of such approach in
mobilizing HSCs and the associated risks involved remain to be explored.
183
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Abstract (if available)
Abstract
Hematopoiesis occurs through complex interplay between hematopoietic stem and progenitor cells and the supportive bone marrow (BM) microenvironment. Defined by the unique ability to self-renew and differentiate into all of the necessary blood cells to support lifelong hematopoiesis, hematopoietic stem cells (HSCs) are spatially located in BM stem cell niches. It was first discovered that primitive HSCs are localized at the endosteal region of the BM. In this region, bone remodeling mediated by bone-forming osteoblasts and bone-resorbing osteoclasts dynamically alters the endosteal surface and releases Ca²⁺ ions from the bone. HSCs are able to sense extracellular Ca²⁺ concentrations through the cell surface expression of the calcium-sensing receptor (CaR). Previous studies demonstrated the crucial role of the CaR in the localization of HSCs in the endosteal BM niche, where CaR⁻/⁻ HSCs displayed stem cell autonomous defects including impaired lodgment and engraftment. We further explored the role of the CaR on HSC functions by using a pharmacologic approach to stimulate the receptor ex vivo with the clinically approved allosteric agonist, Cinacalcet. With CaR stimulaton, HSC homing, lodgment, and engraftment in the endosteal BM niche were promoted via mechanisms involving increased CXCR4 signaling and cell adhesion to major ECM molecules, suggesting that HSC-niche interactions were enhanced to support hematopoietic recovery in the recipient. Although the regulatory functions of the CaR in HSC lodgment and localization have been investigated in normal hematopoiesis, the functional role of the CaR in pathologic hematopoiesis is still poorly understood. In this regard, we used a genetic approach to study the functional role of the CaR in regulating the pathophysiology of multiple myeloma (MM), as this specific hematological malignancy involves devastating bone destruction caused by an excessively active bone-resorbing osteoclastic population, leading to heightened levels of Ca²⁺ ions in the BM endosteal region. We demonstrated that the CaR stimulates myeloma cell proliferation, with mechanisms involving adhesive interactions with the BM microenvironment and cell cycle entry. We also provided the first evidence that the CaR plays a chemoprotective role in MM, as genetic manipulations of CaR expression altered the chemosensitivity of myeloma cells to the chemotherapeutic agent, bortezomib. Finally, we explored the clinical relevance of the CaR in the specification of a functionally distinct HSC population in human hematopoietic tissues based on the cell surface expression of the receptor. Interestingly, we discovered that human HSCs displaying cell surface CaR have impaired hematopoietic stem and progenitor activity in vitro, but similar in vivo engraftment potential in xenogeneic recipients compared to human HSCs lacking cell surface CaR. Although the precise regulatory mechanisms remain to be elucidated, our findings nonetheless suggest that intrinsic functional differences exist in these human HSC populations. Collectively, these data have implications in developing novel therapeutic strategies to enhance HSC engraftment in the transplantation setting by stimulating the CaR, and also in targeting the CaR to abrogate the pathophysiology of bone metastatic cancers by disrupting the complex interplay between the cancer cells and the BM microenvironment.
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Lam, Ben S.
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Core Title
The calcium-sensing receptor in the specification of normal and malignant hematopoietic cell localization in the bone marrow microenvironment
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Keck School of Medicine
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Doctor of Philosophy
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Genetic, Molecular and Cellular Biology
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06/19/2012
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06/12/2012
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Bone Marrow,calcium-sensing receptor,Hematopoiesis,localization,microenvironment,multiple myeloma,OAI-PMH Harvest,stem cells
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calcium-sensing receptor
localization
microenvironment
multiple myeloma
stem cells