THE REGULATION AND FUNCTION OF 1,25-DIHYDROXYVITAMIN D3-
INDUCED GENES IN OSTEOBLASTS
by
AMELIA LOUISE MAPLE SUTTON
Submitted in partial fulfillment of the requirements
for the Degree of Doctor of Philosophy
Advisor: Dr. Paul N. MacDonald
Department of Pharmacology
CASE WESTERN RESERVE UNIVERSITY
August 2005
CASE WESTERN RESERVE UNIVERSITY
SCHOOL OF GRADUATE STUDIES
We hereby approve the thesis/dissertation of
______
candidate for the ______degree *.
(signed)______(chair of the committee)
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(date) ______
*We also certify that written approval has been obtained for any proprietary material contained therein.
I would like to dedicate this dissertation to my amazing mother.
This is the very least I can do to say thank you to the woman who has dedicated her entire life to me.
TABLE OF CONTENTS
Dedication ii
Table of Contents iii
List of Tables iv
List of Figures v
Acknowledgments vii
Abstract xiv
Chapter I Introduction 1
Chapter II The 1,25(OH)2D3-regulated transcription factor MN1 64 stimulates VDR-medicated transcription and inhibits osteoblast proliferation
Chapter III The 1,25(OH)2D3-induced transcription factor 98 CCAAT/enhancer-binding protein-β cooperates with VDR to regulate 24-hydroxylase expression and may be required for osteoblast differentiation and bone mineralization
Chapter IV Targeted overexpression of the 1,25(OH)2D3- 120 regulated gene semaphorin3B in osteoblasts causes increased bone resorption and osteopenia
Chapter V Investigating the function of thrombomodulin, a 160 1,25(OH)2D3-regulated anticoagulant protein, in osteoblasts and in bone
Chapter VI Summary and future directions 183
References 201
iii LIST OF TABLES
Table I-1 Gene Expression Profiling Reveals Decreased Expression 43 of 92 Genes in 1,25(OH)2D3-Treated Osteoblastic Cells
Table I-2 Gene Expression Profiling Reveals Increased Expression of 46 33 Genes in 1,25(OH)2D3-Treated Osteoblastic Cells
iv LIST OF FIGURES
Figure I-1 Metabolism and mineral homeostatic functions of the 48 vitamin D endocrine system
Figure I-2 Domain structure of VDR 50
Figure I-3 The vitamin D receptor undergoes a conformational 52 change upon binding hormone
Figure I-4 Schematic of the conserved domains in SRC family 54 members
Figure I-5 Model of the DRIP complex interacting with liganded VDR 56
Figure I-6 Domain structure of NCoA-62/SKIP 58
Figure I-7 Model of temporal association of coactivators during VDR- 60 mediated transcription
Figure I-8 Long bones are formed by endochondral ossification 62
Figure II-1 1,25(OH)2D3 induces MN1 expression in a time- and dose- 84 dependent manner
Figure II-2 1,25(OH)2D3 induction of MN1 requires de novo RNA 86 synthesis but not protein synthesis
Figure II-3 MN1 augments 1,25(OH)2D3-dependent transcription 88 through the VDR ligand-binding domain
Figure II-4 Effect of MN1 on other nuclear receptors 90
Figure II-5 MN1 synergizes with SRC coactivators 92
Figure II-6 MN1 inhibits proliferation in osteoblastic cells 94
Figure II-7 MN1 decreases S-phase entry in osteoblastic cells 96
Figure III-1 1,25(OH)2D3 increases C/EBPβ expression in primary 112 osteblasts
Figure III-2 C/EBPβ is required for maximal 1,25(OH)2D3-mediated 114 induction of 24OHase expression
v
Figure III-3 Targeted deletion of C/EBPβ disrupts osteoblast 116 mineralization in vitro
Figure III-4 Targeted deletion of both C/EBPβ and VDR causes 118 undermineralized bones in vivo
Figure IV-1 1,25(OH)2D3 induces SEMA3B expression 140
Figure IV-2 SEMA3B is expressed in the long bones of mice 142
Figure IV-3 Establishment of transgenic mice overexpressing SEMA3B 144 in osteoblasts
Figure IV-4 Transgenic mice have reduced body weight and shorter 146 bones
Figure IV-5 Altered cranial morphology in some transgenic mice 148
Figure IV-6 Transgenic mice have decreased bone mineral density 150
Figure IV-7 Transgenic mice have diminished trabecular bone 152
Figure IV-8 Normal osteoblast function but potentially increased bone 154 resorption in transgenic mice
Figure IV-9 Transgenic osteoblasts show increased differentiation and 156 mineralization in vitro
Figure IV-10 Transgenic osteoblasts stimulate increased 158 osteoclastogenesis
Figure V-1 1,25(OH)2D3 induces thrombomodulin expression in a 173 time- and dose-dependent manner
Figure V-2 1,25(OH)2D3 induction of thrombomodulin requires active 175 transcription but does not require protein synthesis
Figure V-3 1,25(OH)2D3 increases protein levels of thrombomodulin 177
Figure V-4 Stable MG-63 cell lines overexpressing thrombomodulin 179 display an increased proliferation rate
Figure V-5 Mice with an osteoblast-selective knockout of 181 thrombomodulin weigh the same as control littermates
vi ACKNOWLEDGEMENTS
There are so many people that have made this dissertation possible that I would need to write an entire other dissertation just to thank them properly. In the next few pages, I hope I can capture just a glimpse of the contributions of so many wonderful people that have touched my life in the last several years.
First, I would like to thank my advisor Dr. Paul MacDonald for guiding me through this process. He gave me the freedom I needed to think and work independently, but always stepped in to offer advice, expertise, or motivation when my work (or
I) was stalled. I will always be deeply indebted to him for all of his encouragement and support while I have been his laboratory. I would also like to thank Dr. Diane Dowd for sharing her technical expertise and continual support both as a member of my thesis committee and in the laboratory. I am very grateful to the other members of my thesis committee, Dr. Hung-Ying Kao, Dr.
Clark Distelhorst, and Dr. John Nilson, for their encouragement, time, and thoughtful scientific discussions throughout this process. I would like to express a special thank you to John for his support throughout my career as an
M.D./Ph.D. student. John’s enthusiasm about science and philosophy of education was one of the things that attracted me to pursuing my Ph.D. in addition to my M.D., and I know that I would not have made it this far without his help.
vii I would like to thank all the past and present members of the MacDonald
laboratory. Dr. Chi Zhang provided much-needed company during many late
nights in the laboratory. I am grateful to Dr. Brenda Altose for her friendship,
kindness, and advice throughout my education. I would like to thank Tara Ellison
for providing endless dietary, intellectual, and emotional nourishment throughout
the past several years. I am indebted to her for her friendship and amazing
culinary prowess. I am grateful to Meika Moore for her excellent technical
assistance in maintaining the mouse breeding and performing flawless
genotyping for me, sometimes on a moment’s notice and always without
complaint. I am very thankful to Xiaoxue Zhang for her hard work in performing
the studies regarding coactivator activity of MN1. I would also like to thank
Fusong Chen for his efforts in continuing the C/EBPβ project.
One of the best aspects of my experience in the Pharmacology department has
been interacting with so many talented, enthusiastic, helpful, entertaining, and supportive graduate students. Whether it has been sharing a protocol or lending
an understanding ear, I would have never been able to complete this dissertation
without them. In particular, I would like to thank Dr. Erin Milliken for
commiserating with me throughout this rather rocky road. I am grateful to both
Erin and Jonathan Mosley for leading the “Science, Non-Science” discussion
group, which has proven to be a very engaging distraction from pipetting. I would
also like to thank Jonathan for his continuous statistical advice and his unique
perspective on just about everything. I am indebted to Mike Davis for his patient
viii assistance with several flow cytometry experiments. Even though these experiments were not fruitful for my project, I learned a great deal from him and really appreciate the time and effort he put into helping me. Through this department and the Medical Scientist Training Program, I have met three lifelong friends that have been an endless source of love and support. First, I would like to thank Dr. Michelle Kahlenberg for her kindness, encouragement, many late nights of ice cream therapy and for forging ahead in medical school so that I may try to follow in her path of excellence. I am indebted to Dr. Nicole Bianco for her support, insightful advice, and hours of comic relief. I would like to express a very special thank you to Dr. Helai Mohammad who has always been there when
I needed her the most, both scientifically and personally, and has provided much entertaining diversions from the laboratory.
I am grateful to all members of the Nilson and Keri laboratories for generously
sharing their technical expertise, equipment, and reagents and for letting me
crash all their parties. I would like to give a special thank you to Dave Peck and
Kristen Lozada, who taught me more than I ever wanted to know about mouse
breeding. I would also like to thank Kristen for her support and friendship, and
taking me out when I needed to get away from the lab. I am indebted to Dr. Ruth
Keri for always being available to discuss science and numerous unrelated
topics, and for always being there with a kind word when I really needed it. Ruth
has also been very open with sharing her laboratory’s reagents and equipment,
and I would not have been able to complete these studies without her generosity.
ix
I am grateful to Drs. Puneet Dhawan and Sylvia Christakos of the Department of
Biochemistry and Molecular Biology at the University of Medicine and Dentistry of
New Jersey for their collaborative efforts on the C/EBPβ project. Likewise, I
would like to thank Dr. Colleen Croniger and her laboratory for performing the
initial breeding and genotyping of the C/EBPβ-null mice and eventually sharing
the strain with us after we established our own animal protocol. I am also
grateful to Colleen for her continuous moral support. I am very thankful for the
collaborative efforts of Drs. Yogendra Kharode and Barry Komm of the Women’s
Heath Research Institute at Wyeth Research in the pQCT and μCT analysis of
the SEMA3B transgenic mice. I would also like to thank Dr. Kimerly Powell and her laboratory of the Department of Biomedical Engineering at the Cleveland
Clinic Foundation for performing the μCT imaging of the VDRKO/C/EBP-βKO and TMflox/flox mice. I am indebted to Drs. Karen Sooy and Marie Demay for sharing their technical expertise in isolating and culturing primary osteoblast
cells. I would also like to thank Dr. Marie Demay for sharing the VDRKO mice with our laboratory. Likewise, I am grateful for the extensive assistance and generous gifts of reagents from Renata Kadlcek and Dr. Edward Greenfield during my forays into osteoclast differentiation studies. Renata was very giving of her time, taking me the through the experiments step-by-step and helping immensely with troubleshooting.
x I am so lucky to have a strangely gigantic family that is full of unconditional love
and unending laughter, which has been much needed over the last several
years. First, I would like to thank my wonderful mother. She raised me as a
single mother throughout most of my formative years, and provided the most
stable loving home I could ever imagine. She taught me that I could be and do whatever I wanted, and has always believed in me even when I did not. She has provided me with an amazing role model for someone who can garner the strength to get through anything that life throws at her. I am incredibly grateful to my aunt, Dr. Barbara Salyer, for her unending emotional, moral, and financial support throughout my education. She has been like a second mother to me, and has encouraged me to keep going even when I didn’t think I could go any further. I would like to thank my late father, Dr. Robert Sutton, for his love and encouragement, and for, both genetically and environmentally, piquing my interest in medicine. I know that he is sharing this special time with me from afar.
I cannot say thank you enough to my sister Mary Ann and brother-in-law John
Church. The labels of “sister” and “brother-in-law” do not adequately describe
my relationship to them, since they have been my second parents, friends, and
the rock I have needed to get through the last 8 years of my life in Cleveland.
Whether it has been bailing me out a financial blunder, providing a hot meal
when I hadn’t seen the inside of my kitchen in weeks, or flying me across the
world for the most memorable vacations, they have enriched my life in the last
several years more than words can describe. I would like to thank my
xi neice/sister, Amy, and her wonderful husband, Romain, for their amazing love
and support, and for living in France so I can come visit them. Amy and I have
grown up together like sisters, and I will always cherish our special relationship. I am grateful to my nephew/brother Tyler and his fiancé Becca for their love and
encouragement. I would like to express my gratitude to my sister Carla and my
sister-in-love Penny for their continuous love and support, and for providing much
needed escapes, replete with animals, from graduate school. I would also like to
thank the rest of the Sutton clan, especially since most of them made the special
trip to share my dissertation defense day with me: Craig, Mary, Molly, Kate, and
Andy, for their love, support, and for hosting Thanksgiving each year; Joel,
Joyce, Erin, Claire for their love and encouragement, and especially Dr. Joel
Sutton for showing me that a Ph.D. is possible for a Sutton; and Marlene, Steve,
Mark, and Jeff for their love and support.
I had the opportunity to interact with many wonderful professors, especially Dr.
William Morgan and Dr. Dean Fraga, during my time at the College of Wooster.
These talented teachers incited a love for science in me. I would like to express my deepest gratitude to Dr. William Morgan, my undergraduate thesis advisor.
His enthusiasm for science and dedication to education was the reason that I decided to pursue science in addition to a medical career. He provided me with my first taste of research and was always extraordinarily patient with while I tried not to break everything in his laboratory. I am incredibly lucky to have worked with such an amazing teacher, scientist, and, most importantly, a truly kind
xii person. I am glad that we have been able to continue our friendship after my graduation, and I know that Bill will always be a fantastic mentor to me.
xiii The Regulation and Function of 1,25-Dihydroxyvitamin D3-Induced Genes in
Osteoblasts
ABSTRACT
By
Amelia Louise Maple Sutton
Vitamin D is essential for calcium and phosphate homeostasis and for protecting
skeletal mineralization. The primary targets of 1,25(OH)2D3, the most bioactive
derivative of vitamin, are the intestine and the kidney where this hormone
stimulates mineral absorption or reabsorption to maintain appropriate serum
concentrations of calcium and phosphate. By tightly regulating systemic mineral ion levels, 1,25(OH)2D3 prevents mobilization of calcium and phosphate stores
from the bone, and, thus indirectly protects bone mineral density. However,
1,25(OH)2D3 also directly influences bone tissue by modulating the activities of
osteoblasts, bone-forming cells, and osteoclasts, bone-resorbing cells.
1,25(OH)2D3 can either inhibit or promote bone mineralization depending on the
physiological or pathophysiological state of the organism. The goal of this
dissertation was to more fully understand the pleiotropic effects of 1,25(OH)2D3 in
the bone by analyzing individual 1,25(OH)2D3-induced target genes. To this end,
we have identified a multitude of 1,25(OH)2D3-regulated genes in osteoblastic
cells through global gene expression profiling. We have chosen four induced
genes, MN1, C/EBPβ, SEMA3B, and thrombomodulin, to further characterize by
xiv exploring the regulation and function of these targets in both isolated osteoblastic
cells and in the more complex bone environment.
First, this dissertation uncovers both a molecular and a cellular function for MN1,
a transcription factor first identified as a gene disrupted in meningiomas, in
augmenting 1,25(OH)2D3-stimulated transcription and in restricting cell growth.
Second, our findings suggest that a cooperative interplay exists between VDR
and the transcription factor C/EBPβ in regulating gene expression and in
modulating bone density. Third, analysis of mice overexpressing SEMA3B, a
secreted cell adhesion molecule, indicates that high levels of this protein causes
severe osteopenia due to enhanced osteoclastogenesis and bone resorption.
Finally, overexpression of thrombomodulin, a transmembrane anticoagulant
protein, accelerates osteoblast proliferation. Osteoblast-selective disruption of
thrombomodulin does not result in overt skeletal abnormalities, but further
analyses are required to fully understand the function of thrombomodulin in bone.
Future studies exploiting the in vitro and in vivo tools developed in this work will
provide a more complete picture of the multiple functions of 1,25(OH)2D3 and its target genes in the bone.
xv CHAPTER I
INTRODUCTION∗
Introduction to the vitamin D endocrine system
The history of vitamin D. Vitamin D was discovered in the 1920’s as the
nutrient that cured rickets (1), a devastating skeletal disease characterized by
undermineralized bones in children (2). Although rickets was recognized as early
as the 15th century, it was relatively rare until the industrial revolution when air
pollution diminished sunlight exposure to the newly urban population (3). This
resulted in an epidemic of rickets in countries such as England and Scotland with
already low sunlight levels. The new concept of dietary compounds, called
“vitamins,” essential for preventing a variety of diseases emerged during the
early part of the 20th century (4). For example, Professor E.V. McCollum
demonstrated that a lipid-soluble substance found in cod liver oil could prevent a
disease characterized by growth retardation and xerophthalmia, a degenerative dry eye condition (1). He termed this substance “vitamin A,” a compound now
known to be critical for proper growth and for maintaining eye health. Building on
the foundation of McCollum’s work, Sir Edward Mellanby performed a series of
experiments with dogs that developed rickets after being housed indoors and fed
exclusively oatmeal. Since these animals were cured with cod liver oil, initially
rickets was erroneously thought to be caused by a vitamin A deficiency. Further
∗ Adapted, in part, from Sutton AL, MacDonald PN 2003 Vitamin D: more than a "bone-a-fide" hormone. Mol Endocrinol 17:777-791 and Dowd DR, Sutton AL, Zhang C, MacDonald PN 2005 Comodulators of VDR-Mediated Gene Expression. In: Feldman D, Pike rel="tag">J.W, Glorieux F.H. (eds) Vitamin D.Academic Press, Amsterdam, 291-304.
1 work by McCollum demonstrated that either heating or bubbling oxygen through
the cod liver oil destroyed the vitamin A activity (supporting growth and eye health), yet retained the ability to cure rickets. Thus, he described this activity as a completely new compound termed “vitamin D” (1). At the same time, several other scientists found that rachitic children could be cured by exposure to ultraviolet light (5). It would not be until decades later when these seemingly
divergent findings would be fully integrated into the complete picture of vitamin D
metabolism.
Synthesis and metabolism of vitamin D compounds. The hormonal or bioactive
form of vitamin D is 1,25(OH)2D3. It is generated from sequential hydroxylations
of vitamin D3, a secosteroid precursor that is obtained from the diet or produced
in the skin upon exposure to ultraviolet light (6, 7) (Fig. 1). The first hydroxylation
of vitamin D3 occurs at the C-25 position and is catalyzed by vitamin D-25-
hydroxylase (25OHase) in the liver to produce 25-hydroxyvitamin D3 (25(OH)D3), the major circulating form of vitamin D in mammals. 25(OH)D3 is the substrate
for a second hydroxylase, the renal 25(OH)D3-1α-hydroxylase (1αOHase),
resulting in the production of the most bioactive metabolite, 1,25(OH)2D3. A classic endocrine feedback system operates to tightly control serum levels of
1,25(OH)2D3 (6, 7). For example, renal 1αOHase activity is stimulated by low
serum calcium and phosphorus levels and by parathyroid hormone. The
expression of 1αOHase is negatively regulated by high levels of 1,25(OH)2D3.
Inactivation, or catabolism, of vitamin D metabolites is initiated by the ubiquitous
2 enzyme 25-hydroxyvitamin D3-24-hydroxylase (24OHase) to generate either
24,25(OH)2D3 or 1,24,25(OH)3D3. The 24-hydroxylated metabolites are further
degraded and eventually excreted as either calcitroic acid or 23-carboxyl
derivatives. This catabolic process is also carefully regulated as 1,25(OH)2D3 stimulates 24-OHase expression to prevent excessive synthesis of the hormone.
Molecular mechanism of action of 1,25(OH)2D3
The vitamin D receptor. The biological effects of 1,25(OH)2D3 are mediated
through the vitamin D receptor (VDR), a member of the nuclear receptor
superfamily of ligand-activated transcription factors (8, 9). Binding of
1,25(OH)2D3 to VDR initiates a cascade of macromolecular interactions ultimately
leading to transcription of select target genes (10). 1,25(OH)2D3 associates with
the VDR and promotes its heterodimerization with retinoid X receptor (RXR), a
common heterodimeric partner for other class II nuclear receptors (11). The
liganded VDR-RXR heterodimer is the functionally active transcription factor in
1,25(OH)2D3-mediated transcription. The heterodimer binds with high affinity to vitamin D response elements (VDREs) in the promoters of target genes. VDREs are characterized by two direct hexameric repeats with an intervening spacer of three nucleotides (DR-3 elements). Thus, 1,25(OH)2D3 target gene selectivity is
conferred, in part, through ligand binding, VDR-RXR heterodimerization, and
high-affinity binding to DR-3 VDREs. Beyond these initial steps, the precise molecular mechanisms involved in target gene activation by VDR are less evident. Recent attention has turned to so-called coactivator proteins that
3 interact directly with VDR and other nuclear receptors in a ligand-dependent manner (12). These coactivators participate in an intricate multiprotein complex together with the basal transcriptional machinery and histone modifiers to stimulate expression of 1,25(OH)2D3-regulated genes (see below).
VDR crystal structure. The vitamin D receptor shares discrete structural and functional domains with other nuclear receptors, but it also exhibits several unique features (6, 10). The hypervariable amino terminal A/B domain of VDR is unusually short and, in contrast to that of most other nuclear receptors, is generally thought to lack potent transactivation domains (Fig. 2). However, as discussed below, there is increasing evidence that the VDR A/B domain helps determine the overall transactivation capacity of the VDR (13). The DNA-binding domain (DBD, or region C) of VDR is similar to other nuclear receptors and is characterized by two zinc-binding modules that direct sequence-specific binding of receptors to DNA (14). The ligand-binding domain (LBD, or region E) is a multifunctional globular domain that mediates selective interactions of the receptor with its cognate hormone (14), with other nuclear receptor partners (15), and with comodulatory or adapter proteins (16). The LBD contains the ligand- dependent activation function-2 (AF-2), which is crucial to ligand-activated transcription. Mutation of the AF-2 renders the nuclear receptor transcriptionally inactive despite retaining the ability to bind ligand (15-17). The DBD and LBD are bridged by the hinge region (domain D), which is thought to confer rotational
4 flexibility between the DBD and LBD and allow for receptor dimerization and interaction with the DNA (18).
While detailed crystal structures for several nuclear receptors have been
available for over a decade (19-23), the structure of the VDR LBD was not solved
until more recently (24). Numerous attempts to crystallize VDR failed, likely due
to the presence of a unique insertion sequence in the ligand-binding domain (see
Fig. 2) that is largely unordered leading to decreased protein solubility. Removal
of this insertion domain allowed for efficient crystallization and structure
determination of the VDR LBD complexed to 1,25(OH)2D3 (24). Although the
lack of this domain may compromise the interpretation of the VDR structure, the
mutant VDR displays normal ligand binding and similar transactivation properties
in vitro (25). Thus, the absence of the insertion sequence does not alter the
conformation significantly so as to compromise VDR function.
The structure of the VDR LBD is similar to that of other nuclear receptors being
most closely related to that of the retinoic acid receptor-γ (RARγ) LBD (24). The
VDR LBD is organized into thirteen α-helices and three β-sheets, which together
form a hydrophobic ligand-binding pocket. This pocket is larger than that of
RARγ due to variations in the positions of helices H2 and H3n and the H6-H7
loop. Helix 12, containing the ligand-dependent activation function-2 (AF-2), of
ligand-bound VDR is positioned similarly to that of other nuclear receptors (21),
highlighting its central importance in creating a coactivator interaction surface
5 (Fig. 3; see VDR Coactivators). In fact, several residues of H12 directly contact
the ligand, indicating that the ligand conformation may modulate H12
conformation and, therefore, coactivator binding and transcriptional activity.
When additional structures of liganded VDR complexed with various coactivators
are solved, they will likely provide a molecular framework on which to develop
new compounds to modulate the vitamin D endocrine system.
In this regard, numerous synthetic analogs have already been developed that
mimic the advantageous effects of 1,25(OH)2D3 without the hypercalcemic side
effects (see below). Speculation about the mechanisms behind the selective,
pleiotropic effects of 1,25(OH)2D3 analogs centers on the concept that these
analogs induce distinct conformations in VDR compared to that of the natural
ligand, ultimately resulting in analog-selective gene regulation (26, 27). Protease
digestions and coactivator binding studies provide experimental support for this
model (26, 28). However, the VDR ligand-binding cavity is larger than that of
many other nuclear receptors, and the ligand occupies less than half of this volume. Consequently, the VDR ligand-binding pocket can accommodate rather significant structural changes in the ligand including 1,25(OH)2D3 analogs with
bulky side chains (24, 29). Indeed, the crystal structures of VDR complexed with
the MC1288 and KH1060 analogs show that these low calcemic analogs do not
induce different conformations in the VDR compared to the natural ligand (30).
Thus, other mechanisms must be considered to explain the different potencies
and calcemic profiles of the analogs. One potential answer resides in the
6 observation that the VDR-analog complexes are more energetically stable than
the VDR-1,25(OH)2D3 complex (30). The increased half-life of the activated VDR
may result in altered transcriptional activity, which may explain the differences
both in potencies and in target gene selectivity between the natural and synthetic
ligands. Alternatively, the solid-state crystal structure may not reveal subtle
dynamic conformational changes in solubilized VDR evoked by various analogs.
VDR Isoforms. Many nuclear receptors, such as RAR, RXR, and thyroid
receptor (TR), have multiple isoforms that are encoded by separate genes (31).
Unlike these nuclear receptors, only one human VDR genetic locus has been identified (32), and the genomic database does not indicate additional highly-
related sequences. Although the cDNA encoding the human vitamin D receptor
was cloned nearly fifteen years ago (9), only recently have several significant
variations in the VDR gene, transcript, and protein sequences been discovered.
At least fourteen distinct transcripts of human VDR have been identified that
differ in their 5’ ends (33). These transcripts arise from alternative mRNA
splicing and differential promoter usage. Most of these variant transcripts utilize
the same initiator codon producing a VDR that is 427 amino acids in length.
However, two transcripts have upstream in-frame methionines that potentially
generate N-terminal extensions in VDR of 50 or 23 amino acids (33). Low levels
of endogenous VDRB1 protein, the 50 amino acid-extended variant, have been
detected in osteoblast, colon cancer, and kidney cell lines (34). Interestingly,
VDRB1 has reduced transcriptional activity compared to classical VDR. Whether
7 the levels of expression of these isoforms are substantial and whether these
isoforms result in altered biological activity in vivo remains unresolved.
Multiple polymorphic variations also exist in VDR in the human population (35).
The vast majority of these polymorphisms do not result in a structural alteration in
the VDR protein with the exception of the Fok I variant (36). The Fok I
polymorphism is located at the original initiator ATG, which is part of a Fok I
endonuclease site. In some humans, there is an ATG Æ ACG transition at the
+1 position, eliminating the translational initiation site and Fok I recognition
sequence. This transition results in the use of an in-frame methionine as the
initiator codon at the +4 position. Thus, either a 427 (Met-1) or a 424 (Met-4)
amino acid protein is expressed. Numerous epidemiological studies suggest an
association between the shorter form of VDR and increased bone mineral density
in humans (37-41). The molecular mechanism of this association remains
unclear, but there is suggestive evidence that the Met-4 VDR displays enhanced
transcriptional activity due to increased interaction with basal transcription factor
TFIIB (13). While these findings remain controversial (42), they raise the
possibility that the N-terminus possesses some type of structure that influences
transcriptional activity. This, combined with the observations that other N-
terminal extensions of VDR may have reduced transcriptional activity, suggests
that there may be inhibitory domains at the extreme N-terminus of VDR that
decrease its transactivation potential.
8 VDR coactivators. The existence of limiting accessory factors or adapter
proteins in steroid hormone receptor action was proposed in the late 1980’s and
early 1990’s based on the “squelching” phenomenon, in which the ligand-binding
domain of one receptor interferes with ligand-activated transcription mediated by
a second receptor (43). A decade later, these comodulatory proteins were
identified as specific molecules that interact with nuclear receptors and influence
their transactivation potential (44-46). The emergence of coactivators, and their
inhibitory counterparts corepressors, provides new insight into the molecular
mechanism of nuclear receptor-mediated transcription. Upon association with its cognate hormone, the receptor ligand-binding domain (LBD) undergoes a subtle
conformational change (47) (Fig. 3). The critical change occurs in helix 12, the
carboxy-terminal alpha-helix containing the ligand-dependent activation function-
2 (AF-2). In response to ligand binding, helix 12 folds over top of the globular
LBD and caps the ligand-binding cavity (21). This ligand-dependent
conformational shift creates a hydrophobic cleft composed of helices 3, 4, 5, and
12 (48-50). The hydrophophic cleft serves as a docking surface for many nuclear
receptor coactivators by interacting with a complementary hydrophobic domain in
the coactivator containing the consensus LXXLL motif, also referred to as the
nuclear receptor (NR) box (51) (Fig. 3). While these studies provide an elegant
structural model for ligand-activated transcription by nuclear receptors and
LXXLL-containing coactivators, the precise mechanisms governing NR-mediated
transactivation are less clear. The ability of coactivators to interact with
components of the pre-initiation complex, with other transcription factors, and
9 with histone-modifying proteins implies that a complex integration of
transactivator cues occurs at the promoter of nuclear receptor target genes. The
growing number of coactivators identified in the last decade adds yet another
level of complexity to the paradigm of nuclear receptor-mediated transcription.
Steroid receptor coactivator-1 (SRC-1 or NCoA1) is the founding member of the
LXXLL motif-containing SRC family of coactivators (46). This family also
includes transcriptional intermediary factor-2 (TIF2; refs. (52, 53)) and receptor
associated coactivator-3 (RAC3; refs. (54-58). The SRCs interact with VDR and
potentiate its transcriptional activity (16, 59). Each of the SRCs possesses an
autonomous transcriptional activation domain (Fig. 4), as evidenced by their ability to enhance transcription when fused to a heterologous DNA-binding
sequence such as GAL4. SRCs stimulate transcription possibly by recruiting
other transcription factors to the promoter. For example, SRCs interact with
cAMP response element binding protein (CREB)-binding protein (CBP)/p300, a
histone acetyltransferase (HAT) that remodels chromatin structure at the promoter (56, 57, 60). SRC proteins also possess intrinsic HAT activity (61).
CBP/p300 directly associates with nuclear receptors and, together with SRCs,
synergistically stimulates transcription (62, 63). Thus, SRCs directly alter
chromatin structure and recruit other factors that modify histones, potentially
providing more accessible promoter templates on which the transcriptional
machinery assembles and initiates transcription of target genes.
10
A large multiprotein complex called DRIP (vitamin D Receptor Interacting
Proteins) was identified as a coactivator for VDR and other nuclear receptors (64,
65). Many components of this complex were discovered separately as thyroid receptor activating protein (TRAP) and the mammalian Mediator complex (66,
67). The diversity of transactivator interactions with the DRIP/TRAP/Mediator complex clearly suggests a more fundamental role for this complex in stimulus- activated transcriptional processes. In VDR-mediated transcription,
DRIP205/TRAP220 acts as an anchoring subunit of the complex by interacting
directly with VDR/RXR heterodimers through one of two LXXLL motifs (68) (Fig.
5). Biochemical depletion of DRIP in cell-free transcription assays shows that
DRIP is essential for VDR-activated transcription in vitro (64). Since the DRIP
complex does not contain SRC proteins and is not associated with HAT activity
(68), it is likely that DRIP and SRCs potentiate transcriptional activation of VDR
through distinct mechanisms. Chromatin immunoprecipitations (ChIP) studies
indicate that a coactivator exchange occurs in the transcriptional complex on
native nuclear receptor-responsive promoters (69-71). Specifically, SRCs
appear to enter the transcriptional complex first and dissociate followed by
binding of the DRIP multimeric complex (70, 71). DRIP is also known to recruit
the RNA polymerase II holoenzyme to VDR upon ligand binding (72). Although
these data conflict, to some extent, with previous studies that show simultaneous
association of SRCs and DRIP with activated nuclear receptor complexes (69),
they do suggest a temporal model in which SRCs enter the complex first to
11 remodel the chromatin, followed by DRIP complex entry and subsequent recruitment of RNA polymerase II (Fig. 7).
In addition to DRIP and SRC coactivators, several other proteins that potentiate
VDR-mediated transcription have been described. One example is NCoA-
62/SKIP, which is a coactivator unrelated to DRIPs, SRCs, and other LXXLL- containing coactivators (73). It interacts with VDR and other nuclear receptors and augments their transcriptional activity. Bx42, the D. melanogaster orthologue of NCoA-62/SKIP, is also implicated in transcriptional processes activated by the insect steroid ecdysone (74). NCoA-62/SKIP was identified independently as ski-interacting protein (SKIP), placing it as part of the TGF-β- dependent Smad transcriptional complex (75). It is also implicated in a number of other transcriptional pathways (76). NCoA-62/SKIP lacks LXXLL motifs and selectively associates with the VDR-RXR heterodimer through the LBD, but through a domain that is distinct from the H3-H5/H12 interactions surface (77)
(Fig. 6). Consequently, NCoA-62/SKIP and SRCs can bind VDR simultaneously to form a ternary complex in vitro (77). NCoA-62/SKIP and SRCs synergistically enhance VDR-stimulated transcription, suggesting a potential interplay between different coactivator classes for maximal activity (77). Although the detailed mechanisms underlying the coactivator activity of NCoA-62/SKIP are unclear, recent evidence suggests that this coactivator may couple VDR-mediated transcription and mRNA splicing. ChIP studies indicate that NCoA-62/SKIP associates with VDR-responsive promoters after SRC recruitment, suggesting
12 that NCoA-62/SKIP functions at a later step in transcription (78). NCoA-62/SKIP
has been identified in subcomplexes of the spliceosome (79-83), indicating a
distinct function in mRNA splicing. Expression of a dominant negative form of
NCoA-62/SKIP suppresses both 1,25(OH)2D3-mediated transcriptional activation
of (77) and appropriate splicing of a VDR-responsive reporter gene (78). Thus,
NCoA-62/SKIP may function as a coupling factor to bridge transcriptional
processes with proper splicing of nascent transcripts. A similar coupling activity
has been observed with other nuclear receptor coactivators (84) such as PGC-1
(85), the DEAD-box RNA helicase p72 (86), and CoAA (87).
Nuclear receptor activity is also modulated by corepressors, the negative
transcriptional counterparts to coactivators (88). Many nuclear receptors actively
repress gene transcription when the receptor is unliganded or bound to
antagonists. This repressive activity is especially prominent in the RXR-
hetereodimeric partners TR and RAR. Unliganded TR- and RAR-RXR
heterodimers interact with corepressors such as nuclear receptor corepressor (N-
CoR) and silencing mediator of retinoid and thyroid hormone receptor (SMRT)
(89). N-CoR and SMRT are components of mulitprotein histone decetylase
complexes, enzymes that oppose the actions of HATs and render the chromatin
environment inaccessible to the core transcriptional machinery. Although VDR
heterodimerizes with RXR, unlike TR and RAR, VDR interacts only weakly with
N-CoR and SMRT (90). However, VDR does interact with two distinct co-
13 repressors, hairless (see below) and alien, both of which repress VDR
transcriptional activity (91, 92).
Since over 30 nuclear receptor coactivators have been identified, recent efforts
have focused on characterizing the unique functions of these proteins and how they cooperate to promote ligand-activated transcription. Biochemical analyses and ground-breaking ChIP studies in intact cells have provided insight into the
complexities of VDR-dependent transcriptional processes. Based on current
data, it appears that each class of coactivator serves a distinct function at defined
time points in transcriptional stimulation (Fig. 7). First, 1,25(OH)2D3 associates
with VDR and promotes heterodimerzation with RXR and binding to VDREs in
the promoters of target genes. The active heterodimer interacts with SRCs,
promoting reorganization of the local chromatin. SRCs dissociate, allowing entry of the DRIP complex, recruitment of the core transcriptional machinery, and initiation of mRNA synthesis. Next, NCoA62/SKIP appears in the complex and promotes splicing of the nascent transcript. This working model provides a reasonable glimpse into the intricacies of VDR-mediated transcription, it is likely that more coactivators and accessory proteins will be discovered and woven into a more complete integrated picture of nuclear receptor-dependent activation.
Non-genomic actions of 1,25(OH)2D3 According to the classical paradigm of
nuclear receptor action, ligand-activated nuclear receptors recruit the basal
transcriptional machinery and other activator complexes to the promoters of
14 target genes to induce transcription. Since these responses require transcription and translation of target genes, they are typically delayed by at least 30 minutes.
However, more rapid (within seconds to minutes) effects in response to steroid hormones are also apparent. The rapid nature of these effects and their relative insensitivity to transcriptional and translational inhibitors, such as actinomycin D and cycloheximide, precludes the possibility that the traditional “genomic” model is operating. Recent attention to these rapid, “non-genomic” hormone effects has spawned renewed interest in this long-standing area of membrane-initiated signaling in the steroid hormone field (93, 94).
Over two decades ago, 1,25(OH)2D3 was shown to evoke transcellular movement of calcium across chick enterocytes within several minutes (95). This phenomenon is theorized to be adaptively beneficial for a hypocalcemic animal in that rapid absorption of calcium occurs without a delayed response involving transcription and translation of calcium-binding proteins or calcium transporters
(96). In addition to the enterocyte, the osteoblast is a target for 1,25(OH)2D3- induced rapid calcium mobilization from internal stores, a process that involves a membrane-initiated signaling cascade including phospholipase C (PLC) activation and inositol triphosphate (IP3) formation (97). This process also occurs
in skeletal muscle cells, where 1,25(OH)2D3 induces calcium release from the
sarcoplasmic reticulum (98), potentially through MAP kinase activation (99).
These are just a few of the many examples of in vitro systems in which these
non-genomic actions of 1,25(OH)2D3 have been studied.
15
Although the rapid effects of 1,25(OH)2D3 and numerous other steroids are well documented, there has been great debate regarding the identity of the putative membrane receptor or receptors that trigger these non-genomic effects. While some studies suggest that the classical VDR mediates the rapid effects, there are other reports indicating that a distinct protein encoded by a separate gene elicits the non-genomic signaling. For example, osteoblast cultures derived from two strains of VDR-mutant mice are unable to initiate a rapid calcium flux in response to 1,25(OH)2D3 (100, 101), strongly suggesting that the traditional VDR mediates at least some of the non-genomic effects. However, a recent report identified distinct gene responsible for the non-genomic responses (102). This gene has been called membrane-associated rapid response steroid binding
(MARRS) protein, and sequence analysis revealed that it is identical to a thioredoxin-like protein called glucose responsive protein 58 kD (GRP58) or endoplasmic reticulum protein 57/60 kDa (ERp57 or ERp60). Treatment of cells with either an inhibitory antibody or with ribozymes targeted against MARRS indicated that this protein, and not the classical VDR, mediated rapid phosphate uptake and stimulation of protein kinase C activity (102). Clearly more detailed analyses in VDRKO cells and in MARRS-depleted cells are required to determine which gene product regulates some or all of these rapid non-genomic effects of
1,25(OH)2D3.
16 Physiological functions of the vitamin D endocrine system in non-classical target tissues
Overview. Much of our understanding of the physiology of the vitamin D
endocrine system has stemmed from classic dietary manipulations and from the
analysis of inherited disorders in humans (103, 104), The recent development of murine genetic models in which key genes in this endocrine system have been systematically eliminated highlight the essential role of 1,25(OH)2D3 in
maintaining mineral homeostasis as well as reveal more subtle actions of this
hormone in other physiological processes. In the current section, I will discuss
some of the effects of 1,25(OH)2D3 and VDR in “non-classical” processes, such
as hair follicle cycling, blood pressure regulation, mammary gland development,
and tumorigenesis. The following section (Effects of the vitamin D endocrine
system on bone development and maintenance) provides a more detailed
discussion of the actions of the vitamin D endocrine system in mineral
homoeostasis and in the skeleton in the context of other factors that control bone
development and remodeling.
As discussed below, the vitamin D receptor knockout mouse provides a murine
model of the human disorder rickets, characterized by undermineralized bones
due to hypocalcemia (105, 106). These mice also exhibit alopecia, the absence
of functional hair follicles, a phenotype that is not corrected by normalization of
serum calcium levels (107). While the VDRKO keratinocytes proliferate and
differentiate normally, they fail to properly initiate hair re-growth following
17 depilation (108, 109). Due to the lack of feedback regulation of the anabolic 1α-
hydroxylase enzyme and the catabolic 24-hydroxylase, the VDRKO animals have
abnormally high levels of 1,25(OH)2D3. Thus, one potential cause of alopecia in
VDRKO animals is 1,25(OH)2D3 toxicity. To address this possibility, VDRKO
mice were raised and bred for five generations in an ultraviolet light-free
environment and on a diet lacking vitamin D derivatives (109). Despite having
undetectable levels of 1,25(OH)2D3, fifth-generation vitamin D-deficient VDRKO
mice still have alopecia. Thus, 1,25(OH)2D3 toxicity does not cause alopecia in
VDRKO mice. Since wild-type littermates of VDRKO mice raised under the same
vitamin D-deficient conditions do not display alopecia, Sakai and colleages
proposed that VDR may regulate hair follicle cycling in a ligand-independent
fashion (109). Further support for this hypothesis comes from the observation
that mice with a targeted deletion in 25(OH)D3-1α-hydroxylase (see below), the
biosynthetic enzyme that produces 1,25(OH)2D3, do not display alopecia (110).
Expression of a keratinocyte-selective transgene that encodes a mutant VDR incapable of binding 1,25(OH)2D3 completely restores hair growth in VDRKO
mice, while expression of VDR with a mutation in the coactivator-binding domain
partially rescues the alopecia phenotype (111). This study indicates that the
ligand-binding activity is dispensable for VDR to properly regulate hair follicle
cycling. The alopecia observed in VDRKO mice is, in fact, a phenocopy of mice
with a targeted deletion in the transcription factor hairless (112). Hairless
interacts with VDR and potentially acts a co-repressor for VDR (91). Thus, VDR
may control hair follicle cycling by interacting with hairless and repressing select
18 genes. While these studies introduce a novel and potentially significant concept
in VDR biology, identifiying target genes and establishing molecular mechanisms
that govern the function of unliganded VDR in keratinocytes are important goals
for future research.
Several clinical studies have proposed that 1,25(OH)2D3 may be beneficial to the
cardiovascular system by decreasing blood pressure (113, 114). Consistent with
these observations in humans, VDRKO mice have increased renin expression
resulting in higher levels of angiotensin II, increased water intake, electrolyte
disturbances, elevated blood pressure, and cardiac hypertrophy (115).
Furthermore, high levels of renin and angiotensin II persist despite normalization
of mineral ion levels with the rescue diet. This response appears to be at the
transcriptional level as 1,25(OH)2D3 suppresses renin promoter activity. This
study suggests that VDR negatively regulates the expression of renin, allowing
for decreased angiotensin production and lower blood pressure. The relevance
of this study to human hypertension is not entirely clear since there are no
reports of hypertensive HVDRR patients. Regardless, these are provocative
observations in the VDRKO model that may stimulate a more extensive
examination of 1,25(OH)2D3 and its synthetic analogs as potential therapies for some forms of hypertension.
Mammary gland development is also emerging as a biological process that is
impacted by 1,25(OH)2D3. The mammary glands of VDRKO mice demonstrate a
19 hyperproliferative phenotype as evidenced by increased numbers of terminal end buds and enhanced ductal branching compared to wild-type littermates (116).
VDRKO mammary glands also show accelerated ductal development and increased proliferation in response to exogeneously administered estrogen and progesterone (116). After cessation of lactation, VDRKO glands demonstrate delayed regression due to impaired apoptosis (117). While dietary calcium supplementation normalizes estrogen levels in VDRKO mice, the abnormal mammary phenotype is retained (116). These data indicate a significant developmental role for VDR in the mammary gland potentially in restricting ductal growth and promoting apoptosis. Recently, VDRKO mice were bred with an established murine model of mammary cancer, which is caused by transgenic expression of the oncogene neu under the control of the mouse mammary tumor virus (MMTV) promoter(118). MMTV-neu mice heterozygous for VDR developed increased numbers of mammary tumors with decreased latency. Combined with the observations that 1,25(OH)2D3 and its analogs inhibit the growth and induce the differentiation of breast cancer cell lines (119, 120), the mammary phenotype in the VDRKO model indicates that 1,25(OH)2D3 and its derivatives may be useful therapies for breast cancer. In addition to inhibiting mammary tumor development, VDR also protects against skin cancer in mice. Treatment of
VDRKO mice with the chemical carcinogen 7,12-dimethylbenzanthracene
(DMBA) induces rapid formation of multiple skin tumors (121). The VDRKO skin exhibits a hyperproliferative phenotype (121), which may account for the propensity to develop tumors. In fact, a similar effect of the vitamin D endocrine
20 system has been observed in human skin. Calcipotriol, a synthetic analog of
1,25(OH)2D3, is a first-line treatment for the hyperproliferative skin disorder psoriasis (122). These studies highlight the therapeutic potential of the vitamin D endocrine system in preventing and treating various forms of cancer.
Effects of the vitamin D endocrine system on bone development and maintenance
Overview. Vitamin D is essential for regulating mineral ion homeostasis and protecting skeletal integrity. 1,25(OH)2D3 affects the bone both indirectly, through promoting dietary calcium absorption, and directly, by modulating osteoblast and osteoclast activity. Recent studies have uncovered numerous complexities in the actions of 1,25(OH)2D3 and VDR. Before discussing the specific functions of the vitamin D endocrine system in the skeleton, I will first provide a general overview of bone development and remodeling.
Bone development. Skeletal organogenesis is one of the last developmental events to be completed in mammals. While bone development begins as early as the fifth week of gestation in humans, the mature skeleton is not fully formed until puberty (123). The primordial skeleton arises from three embryonic lineages. Neural crest cells differentiate to form the majority of the bones of the skull and face, mesodermal cells from somites develop into the axial skeleton, and lateral plate mesodermal cells create the bones in the limbs (124).
These undifferentiated precursor cells condense at the sites that will eventually become skeletal elements. Precursor cells form bone through two discrete
21 pathways: intramembraneous ossification and endochondral ossification (125).
Mesenchymal condensations directly differentiate into osteoblasts during intramembraneous ossification to form the clavicles, mandibles, and skull plates.
Endochondral ossification forms the rest of the future skeleton and proceeds through a cartilage intermediate that is eventually replaced by mature bone.
In endochondral ossification, pluriopotent mesenchymal cells nucleate and begin to differentiation into chondrocytes, cartilage-forming cells (126) (Fig. 8). These immature chondrocytes secrete type II collagen into the extracellular matrix
(ECM). The innermost chondrocytes further differentiate into hypertrophic chondrocytes, secrete an ECM rich in type X collagen, and begin to mineralize the ECM. Some of the cells remain undifferentiated around the periphery of the primitive cartilage template and form the perichondrium. These periphery cells differentiate directly into osteoblasts and mineralize the perichondrium to create a bone collar, the precursor of the future dense cortical bone (127). After the initial cartilage is calcified, the hypertrophic chondrocytes undergo apoptosis while blood vessels invade the cartilage anlage. This blood vessel invasion creates a conduit for the migration of chondroclasts, osteoclasts, osteoblasts, and hematopoetic stem cells into the center of the tissue. The chondroclasts and osteoclasts destroy most of the cartilage, leaving behind remnants that form scaffolds for osteoblasts to begin depositing and mineralizing collagen type I- containing ECM (128). This bone forms in a lace-like pattern that will develop
22 into the future trabecular bone, which is interwoven with the hematopoetic marrow tissue (Fig. 8).
Ossification proceeds radially until most of the cartilage template is replaced by mature bone. However, some chondrocytes remain on the two epiphyseal ends of the ossification center and continue to proliferate and differentiate, forming the growth plates of each long bone (129). The growth plates allow for elongation of the bone tissue until an organism reaches full stature at puberty. Growth plate chondrocytes are arranged in columns according to their differentiation stage
(Fig. 8). The resting chondrocytes reside at the epiphyseal end of the growth plate and, when signaled to, will develop a more flattened appearance as they begin to divide and move into the proliferative zone. Once proliferation ceases, chondrocytes begin differentiating and proceed to the pre-hypertrophic and hypertrophic zone, where the cells are large and cuboidal. Hypertrophic chondrocytes represent the terminal stage of differentiation and will express and mineralize an ECM primarily composed of collagen type X (127). At puberty, when longitudinal growth ceases, the cartilaginous growth plates are completely replaced by mature bone.
Factors controlling bone development. Like all developmental processes, a complex network of transcription factors, signaling molecules, and hormones directs bone development. Members of the Sox family of transcription factors, which contain a high-mobility-group (HMG)-box DNA-binding domain (DBD)
23 similar to the DBD of the sex determining factor SRY, are essential for normal chondrocyte differentiation (130). Gene targeting strategies in mice have revealed that Sox9 is absolutely required for any cartilage formation (131) and likely acts early in chondrocyte differentiation to direct initial mesenchymal condensations (132). Sox9 is also mutated in the human genetic disease campomelic dysplasia, characterized by hypoplasia of all endochondrally-formed bones (133, 134), further highlighting the importance of this gene in cartilage development. Two related transcription factors, Sox5 and Sox6, function at a slightly later step in chondrocyte differentiation. In mice with disruptions in both
Sox5 and Sox6, mesenchymal cells coalesce normally but do not differentiate appropriately into proliferating chondrocytes and form severely abnormal cartilage (135). Since deletion of either Sox5 or Sox6 alone results in only minor cartilage abnormalities and the genes share a high degree of sequence identity, these two transcription factors are likely functionally redundant (135).
Core binding factor α1 (Cbfa1) or runx2, a member of the Runt-domain family of transcription factors, is a master regulator of osteoblast differentiation. Mutations in just one copy of runx2 in humans cause cleidocranial dysplasia (CD), a disease characterized by short stature, hypoplasia or aplasia of the clavicle, a wide pubic symphysis, delayed closure of the cranial fontanels and sutures, and other skeletal abnormalities (136, 137). Homozygous runx2-deficient mice form no mature osteoblasts or bone, and heterozygous mutant mice display bone abnormalities similar to CD in humans (138, 139). The bone defect in runx2
24 mutant mice is, in part, due to an early block in osteoblast differentiation since only osteoblast progenitor cells are detected in the mutant mice (138).
Furthermore, differentiation of hypertrophic chondrocytes is impaired in some bone precursors in ruxn2-deficient mice (140, 141). Transgenic expression of runx2 selectively in chondrocytes in runx2 mutant mice partially rescues the phenotype resulting in a skeleton of mineralized cartilage that still lacks intramembraneously-formed bones (142). These results indicate that runx2 is essential for bone formation by stimulating differentiation of both osteoblasts and chondrocytes from mesenchymal precursors. Osterix is another transcription factor critical for osteoblast differentiation and bone formation (143). Inactivation of osterix results in a skeleton composed of only mineralized collagen due to an arrest of osteoblast differentiation after mesenchymal condensation. Since osterix-null mesenchymal precursor cells maintain runx2 expression, osterix is thought to function downstream of runx2 in the osteoblast differentiation pathway.
In addition to transcription factors, several soluble factors are critical in bone development. For example, tight control of fibroblast growth factor (FGF) signaling is essential to proper bone formation. FGFs signal through fibroblast growth factor rectors (FGFRs), tyrosine kinase receptors that, upon ligand binding, homodimerize and initiate autophosphorylation. This autophosphorylation activates the kinase activity, allowing for phosphorylation of downstream targets and stimulation of a complex signaling cascade (144).
FGFRs were recognized as important players in skeletal development when
25 mutations in FGFR3 were identified as the cause of achondroplasia and other skeletal dysplasias (145). These mutations cause constitutive activation of
FGFR3, resulting in shortened malformed limbs. Mice with an analogous mutation in FGFR3 display a phenotype similar to human achondroplasia (146,
147). Accordingly, mice with targeted deletions in FGFR3 have longer bones
(148, 149). Analysis of these genetic mouse models and further in vitro experiments show that FGF signaling inhibits chondrocyte proliferation and promotes chondrocyte differentiation and hypertrophy (145).
Bone morphogenic proteins (BMPs) are members of the TGF-β superfamily of secreted growth factors (150). As their name implies, BMPs regulate bone development, but they also modulate organogenesis and homeostasis in a variety of other tissues. Like other TGF-β-related ligands, BMPs bind transmembrane serine threonine kinase receptors resulting in phosphorylation of
Smad proteins (151). Smads transmit the signal to the nucleus by modulating expression of target genes. BMPs were first discovered as bone-derived proteins capable of stimulating ectopic bone formation (152). Since then, accumulating evidence in mice indicates that BMP signaling regulates skeletal patterning, induces chondrocyte differentiation, promotes osteoblast proliferation, and controls bone mass acquisition (126).
Two additional secreted factors, Indian hedgehog (Ihh) and parathyroid hormone related peptide (PTHrP), participate in a regulatory circuit that tightly controls
26 bone development (128). During bone development, Ihh is expressed by
prehypertrophic chondrocytes and stimulates perichondrial cells to produce
PTHrP. PTHrP feeds back on the chondrocytes to inhibit terminal differentiation
(153). Ihh also directly promotes prehypertrophic chondrocyte proliferation (154,
155). Thus this signaling pathway is important in preserving appropriate
chondrocyte proliferation and preventing premature chondrocyte differentiation.
Ihh is also critical for osteoblast differentiation since Ihh-null mice completely lack
osteoblasts in endochondrally-formed bones likely due to defects in runx2
expression (155).
The Ihh pathway regulates bone development through yet another mechanism by
stimulating Wnt signaling in osteoblasts (156). Wnts are secreted proteins that
bind to and activate receptors of the frizzled family and co-receptors of the
LRP5/6 family (157). The activated receptors initiate a signaling cascade
resulting in stablilization of the β-catenin protein. β-catenin enters the nucleus
where it cooperates with the tcf/lef family of transcription factors to regulate
expression of target genes. Recent studies have shown that Wnt signaling
modulates osteoblast differentiation and bone mass acquisition in both humans
and mice. Disruptive mutations in the LRP5 co-receptor gene cause the genetic
disease osteoporosis psueodoglioma syndrome (158), characterized by
undermineralized bones, blindness, and cognitive impairment (159). Conversely,
an activating mutation in LRP5 results in a high bone mass phenotype (160,
161). These observations have been recapitulated in genetically-engineered
27 mice with either targeted deletions in LRP5 (158) or with transgenic expression of
an activating mutant of LRP5 (162). Thus, wnt signaling appears to play a crucial
role in regulating bone mineralization. The importance of Wnt signaling in
osteoblast differentiation is further underscored by studies in conditional
knockout mice. When β-catenin is selectively disrupted in mesenchymal
precursor cells, osteoblast differentiation is completely blocked and mineralized
bone does not form (156, 163).
Vascular endothelial growth factor (VEGF) has a well-defined role in directing
vascular development and in regulating angiogenesis in both normal and
neoplastic tissues (164), yet the importance of this secreted growth factor in bone
development has just recently been recognized. Alternative splicing of the VEGF
mRNA results in several isoforms of the protein: VEGF120, VEGF164, and
VEGF188. VEGF molecules signal through two classes of receptors: the VEGF protein tyrosine kinase receptors (VEGFRs) and neuropilins (164).
Characterization of mutant mice that only express the VEGF120 isoform, which only binds the VEGFRs but not neuropilins, revealed a surprising defect in bone development (165). These mutant mice display a dramatic decrease in blood vessel invasion into the ossification centers of developing long bones and, thus, they undergo significant delay in bone mineralization. Administration of a neutralizing soluble VEGF receptor to mice has a similar effect in bone formation
(166) and inhibits fracture repair (167). These studies demonstrate that vascular
28 invasion of the cartilage template is a critical, VEGF-dependent step during bone
development.
Bone remodeling. Even once the mature skeleton is formed, the bone tissue undergoes continuous renewal, or “remodeling,” throughout an organism’s lifetime to preserve the structural integrity of the tissue and to maintain systemic mineral homeostasis. This remodeling requires a delicate balance between the activities of osteoblasts, the bone forming cells, and osteoclasts, the bone resorbing cells. Any imbalance in the opposing activities of osteoblasts and osteoclasts can result in a variety of metabolic bone diseases, the most common of which is osteoporosis. In osteoporosis, bone resorption exceeds bone formation. The progressive destruction of skeletal microarchitecture in osteoporosis causes decreased bone density and an increased risk for fractures
(168). An imbalance in the other direction, when bone formation exceeds bone resorption, results in the rare disease osteopetrosis. Osteopetrosis is characterized by abnormally dense bone that is brittle and occludes the marrow cavity, impairing proper hematopoiesis (169). Therefore, bone formation and bone destruction are two tightly coupled processes that remain in equilibrium through an intimate communication between osteoblasts and osteoclasts in the skeleton.
Adult bones are composed of two types of tissue: cortical and trabecular bone
(Fig. 8). The cortical bone, derived from the primitive perichondral bone collar, is
29 the dense mineralized tissue that surrounds the marrow cavity and constitutes
about 80% of the skeletal mass (170). The trabecular bone is a complex scaffold-like tissue that intercalates with the hematopoetic marrow and provides
the majority of structural support in the skeleton. Bone remodeling occurs
primarily on the surfaces of the tissue. Since trabecular bone has a much
greater surface-to-volume ratio than cortical bone, the majority of remodeling occurs in trabecular bone. Thus, disorders characterized by abnormalities in bone turnover tend to affect trabecular bone disproportionately as compared to cortical bone (170).
As discussed previously, mesenchymal stem cells differentiate into osteoblasts,
which secrete and mineralize ECM proteins. Once osteoblasts are surrounded by calcified bone, they terminally differentiate into osteocytes and become trapped within the matrix where they continue to synthesize mineralized bone tissue. Osteoblasts and osteocytes primarily secrete type I collagen, a triple- helical protein consisting of two α(1)I and one α(2)I chains (171). These collagen molecules are extensively post-translationally modified and form multiple intra- and intermolecular cross-links to provide mechanical stability. Although the bone
ECM is composed of 85-90% type I collagen, it also contains several other proteins, including proteoglygans, hyaluronan, glycoproteins, and γ-carboxylic acid (gla)-containing proteins. Osteocalcin, one of the gla-containing proteins, is the most abundant of the non-collagenous proteins in the bone and is expressed only in osteoblasts and osteocytes. Although the precise role of osteocalcin in
30 bone is unclear, osteocalcin-deficient mice have an increase in bone mass indicating that this protein likely limits excess mineralization (172).
After secreting these matrix proteins, osteoblasts and osteocytes begin depositing minerals to form mature bone. Bone is primarily composed of hydroxyapaptite (Ca10(PO4)6(OH)2) (173). Osteoblasts initiate mineralization by releasing membrane-bound extracellular vesicles. These vesicles promote crystal formation by concentrating calcium and phosphate ions in a protected environment. Several ECM phosphoproteins, such as osteopontin and bone sialoprotein, regulate this initial nucleation event (174, 175). Alkaline phosphatase, a membrane-bound enzyme highly expressed by osteoblasts, removes phosphate groups from these proteins, thus increasing the local phosphate concentration and stimulating mineral deposition (173). In fact, humans and mice with mutations in alkaline phosphatase are unable to properly mineralize bone (176, 177). Minerals continue to grow larger in all three dimensions to form a hardened lattice that provides mechanical stability to the bone tissue (173).
While osteoblasts and osteocytes create bone during initial skeletal development and during the remodeling process, osteoclasts are the specialized cells responsible for destroying bone tissue (169, 178, 179). Early studies indicating that bone marrow transplants could correct osteopetrosis in mice and in humans revealed the hematopoietic origin of osteoclasts (180, 181). Osteoclasts are now
31 known to differentiate from the monocyte/macrophage lineage to form giant
multinucleated cells that remain in close communication with osteoblasts to
properly renew bone tissue.
The earliest event in osteoclast differentiation is expression of the Ets domain transcription factor PU.1. Mice deficient in PU.1 are severely osteopetrotic since they completely lack osteoclasts and their precursors monocytes (182). PU.1 can interact with the microophthalamia-associated transcription factor (MITF), which contains basic helix-loop-helix and leucine-zipper motifs, to regulate
several genes essential for osteoclastogenesis (183). Although mutations in
MITF cause osteopetrosis in mice (184), this transcription factor is thought to act
later than PU.1 in differentiation since osteoclasts are present but appear
abnormal in MITF mutant mice (185). As bone marrow transplants correct the osteopetrosis in mice with mutations in PU.1 or MITF (182, 186), these transcription factors are thought to act intrinsically in osteoclast development.
In addition to these intrinsic genes, several extrinsic osteoblast-derived factors
are essential for proper osteoclast development. Analysis of the osteopetrotic
(op/op) mouse revealed the pivotal role that the cytokine macrophage colony
stimulating factor (M-CSF) plays in osteoclast formation (187). Young op/op mice have no osteoclasts and the phenotype cannot be rescued by bone marrow transplantation (187). Further studies revealed that osteoblasts are the source of
M-CSF (188). M-CSF signals through the receptor tyrosine kinase c-fms to
32 promote chemotaxis, proliferation, and survival of osteoclast precursors and
mature osteoclasts (189, 190).
Another extrinsic factor essential for osteoclastogenesis is receptor activator of
nuclear factor-κB ligand (RANKL; also called osteoprotegerin ligand) (191-193).
RANKL is a member of the tumor necrosis factor family of cytokines and is
expressed by osteoblasts. There are three forms of RANKL: an extracellular
membrane-bound protein, a shorter soluble peptide formed by cleavage of the membrane bound form, and a secreted factor (194). In the bone, the membrane-
bound form is thought to be the dominant signaling molecule (191), implying that
close contact between osteoblasts and osteoclasts is required for
communication. RANKL stimulates osteoclast differentiation from immature
precursor cells (195-198), promotes osteoclast survival and activation (197, 199-
201), and induces osteoclast adhesion to the bone surface (202). RANKL
signals through its receptor, receptor activator of nuclear factor-κB (RANK), a
transmembrane protein expressed on osteoclast precursors (192, 203-205).
RANKL can also associate with a soluble decoy receptor, osteoprotegerin, which prevents RANKL from binding to RANK and inhibits osteoclast differentiation
(206). Upon RANKL binding, RANK relays the signal by associating with tumor necrosis factor receptor-associated factor 6 (TRAF6) and the tyrosine kinase c- src. These effectors signal through numerous pathways, including the mitogen- activated protein kinase (MAPK) cascade and calcium fluxes, eventually converging on transcription factors including NFκB, activator protein-1 (AP1), and
33 NFATc1 (207). NFATc1 (also called NFAT2) is a member of the nuclear factor of
activated T cells (NFAT) family of transcription factors originally identified as
important immunomodulatory proteins (208). This transcription factor is
considered a master regulator of osteoclast differentiation since cells with a
targeted deletion in NFATc1 cannot form osteoclasts and ectopic expression of
NFATc1 drives osteoclast differentiation in the absence of RANKL stimulation
(209).
As osteoclasts differentiate from hematopoietically-derived precursors, they fuse
into multinucleated giant calls and attach to the bone tissue by forming
podosomes, foot-like structures containing filamentous actin and integrins (210,
211). Thus, the osteoclast multikaryon polarizes and creates two distinct
membrane surfaces: the basolateral membrane and the bone-facing membrane
where resorption occurs. The resorptive surface develops a “ruffled membrane”
where the cell secretes acidifying vesicles that lower the pH in the local
microenvironment (212, 213). At a low pH, the inorganic hydroxyapatite of bone dissolves into free calcium and phosphate ions and water (214). The osteoclast also secretes proteases, such as cathepsin K and matrix metalloproteinases, to degrade the organic components of bone (215, 216). The resulting debris is packaged into vesicles and taken up by the osteoclast cell body. The enzyme tartrate-resistant acid phosphatase (TRAP) is contained within the vesicles and produces reactive oxygen species that participate in the further destruction of collagen and other bone proteins (217). Eventually, the degradation products are
34 vesicularly transported through the cytoplasm and released through the opposite basolateral membrane (218, 219). Once local bone degradation is complete, osteoclasts vacate the resorptive surface and osteoblasts appear and begin synthesizing new bone, as described above, to complete the cycle of bone remodeling (170).
The impact of the vitamin D endocrine system on bone. The effects of
1,25(OH)2D3 and VDR on the skeleton are multifaceted, involving both alterations in systemic mineral ion homeostasis and changes in local osteoblast and osteoclast activity. A combination of both in vitro and in vivo studies in rodent models and in humans have underscored the fundamental role the vitamin
D endocrine system plays in maintaining serum calcium and phosphate concentrations and the more complex direct actions of this hormone system in bone tissue.
Mice with targeted deletions of key genes in the vitamin D endocrine system provide important tools to study the in vivo functions of vitamin D signaling pathways. Two groups independently created mouse strains with targeted deletions in the VDR gene by disrupting either exon 2 (105) or exon 3 (106). Not surprisingly, the VDR knockout (VDRKO) mice display all of the features of the human disease hereditary vitamin D-resistant rickets (HVDRR), a rare genetic disorder caused by mutations in the VDR gene (103, 104). The VDRKO mice are viable and develop normally until the weaning period. However, shortly after
35 weaning VDR null mice exhibit alopecia, as discussed above, and growth retardation accompanied by progressive hypocalcemia, hypophosphatemia, and compensatory hyperparathryoidism. VDRKO mice display diminished intestinal expression of several 1,25(OH)2D3-regulated genes putatively involved in calcium
transport, including calbindin D9K, calcium transport protein-1, and epithelial
calcium channel (220-222). In addition to the intestinal defect in calcium
absorption, mice that express a mutant VDR that lacks the DNA binding domain
have decreased renal reabsorption of calcium (100). These studies reinforce the concept that both the intestine and the kidney are essential VDR target organs in maintaining calcium homeostasis.
These metabolic imbalances in VDRKO mice result in severe skeletal defects,
including decreased bone mineral density, thinned bone cortex, and widened
undermineralized growth plates. However, when VDRKO mice are fed a rescue
diet rich in calcium and phosphorus to normalize serum calcium and PTH levels,
the mice develop without bone abnormalities (107). Further analyses indicated
that the skeletons of these mice on the rescue diet appear grossly, histologically,
and biometrically normal (223). These findings suggest that the bone defect in
VDR null mice is secondary to the malabsorption of calcium in the intestine and
is not due to the lack of a direct effect of 1,25(OH)2D3 on the bone. A phenotype
similar to the VDRKO mice was observed in animals with a targeted disruption of
1α-hydroxylase (1αOHase), the enzyme responsible for the regulated synthesis of the active hormone (110, 224). 1αOHase knockout (1αOHaseKO) mice also
36 display hypocalcemia, hyperparathyroidism, growth retardation, and
osteomalacia consistent with rickets but, as discussed previously, do not have alopecia. Administration of exogenous 1,25(OH)2D3 (225) or the high calcium
rescue diet (226) has been reported to correct the bone abnormalities.
However, a recent study examining VDRKO, 1αOHaseKO, and
VDRKO/1αOHaseKO mice with various dietary manipulations has somewhat contradicted these earlier findings (227). Even fed the high-calcium rescue diet, all three knockout animals still exhibited significantly decreased trabecular bone volume, reduced numbers of osteoblasts, and a diminished osteoblastic activity as measured by the mineral apposition rate. Thus, although serum calcium and parathyroid hormone levels were normal, the bones remained osteopenic. The differences in these studies could be attributed to the duration of the rescue diet as mice were fed the diet for approximately 100 days in the Panda et al. study
(227), while they were only fed the diet for 39 or 54 days in the previous
1αOHaseKO (226) and VDRKO studies (107), respectively. Further, the mice utilized in the Panda et al. study were of a mixed genetic background, with contributions from the BALB/c and C57BL/6J strains (227), while the mice in the previous rescue diet studies were in C57BL/6J exclusively (107, 226). Thus, genetic background or subtle diet differences or both may modify the actions of the vitamin D endocrine system in regulating bone mass and osteoblastic function. In further support for a direct function of 1,25(OH)2D3 and VDR in bone,
transgenic mice overexpressing VDR in osteoblasts display increased trabecular
37 bone volume, increased bone strength, and decreased bone resorption (228).
These transgenic bones are more mineralized as VDR-overexpressing
osteoblasts deposit increased levels of calcium in the ECM (229). Such data
indicate that the vitamin D endocrine system has a direct anabolic effect on bone.
In contrast, studies with isolated VDRKO bones and osteoblasts suggest that
VDR actually inhibits mineralization. To avoid the effects of VDR on systemic mineral ion levels, bones from VDRKO mice were transplanted into wild-type mice and evaluated for changes in mineral density (230). Surprisingly, VDRKO bones in wild-type hosts had an approximately 48% increase in bone density compared to wild-type bones transplanted into wild-type hosts. Similarly, primary osteoblasts derived from VDRKO mice appear to differentiate and mineralize at an accelerated rate in vitro (231). Thus, these two studies suggest that VDR functions to limit bone mineral deposition. The opposing findings in these various studies will require more detailed in vivo and in vitro analyses to more fully understand how 1,25(OH)2D3 and VDR function locally in the bone.
Prior to development of these knockout models, several in vitro studies indicated that 1,25(OH)2D3 has a biphasic effect on osteoblast differentiation (232, 233).
When appropriately stimulated, primary osteoblasts in culture proceed through a
regulated program of differentiation eventually resulting in mineralized bone
nodules. If 1,25(OH)2D3 is added prior to cell confluence, the hormone inhibits
maturation of osteoblasts. However, when more mature osteoblasts are treated
38 with 1,25(OH)2D3, the cells differentiate at an accelerated rate (234). These
data suggest that 1,25(OH)2D3 may inhibit early osteoprogenitor differentiation while promoting late-stage maturation of osteoblasts and osteocytes. In fact,
1,25(OH)2D3 induces synthesis and secretion of the bone matrix proteins
osteocalcin and osteopontin (235, 236), molecules associated with a more
mature stage of osteoblast differentiation. 1,25(OH)2D3 regulates these two genes transcriptionally by binding to VDREs in their promoters (237, 238).
1,25(OH)2D3 also increases osteoblastic alkaline phosphatase activity (239),
which, as previously discussed, promotes mineralization of the extracellular
matrix.
Some of the conflicting findings in the actions of the vitamin D endocrine system
on bone may be explained by the confounding effects of 1,25(OH)2D3 on osteoclasts. In conjunction with PTH, 1,25(OH)2D3 stimulates bone mineral
mobilization (240) by promoting osteoclastogenesis (241). Co-culture systems
have demonstrated that cell-cell contact between osteoblasts and osteoclast
precursors is essential for 1,25(OH)2D3-stimulated osteoclastogenesis (242). In
fact, 1,25(OH)2D3 promotes surface expression of RANKL on osteoblasts (243)
by stimulating mRNA levels of RANKL (244). Although RANKL is thought to be
the major factor that mediates 1,25(OH)2D3-induced osteoclastogenesis,
1,25(OH)2D3 also decreases expression of osteoprotegerin, the soluble decoy
receptor for RANKL (245). These bone-resorbing effects of 1,25(OH)2D3 relate back to the prominent role of this hormone in regulating mineral ion homeostasis.
39 It is thought that under hypocalcemic conditions, 1,25(OH)2D3 acts in concert with
PTH to mobilize calcium from skeletal stores through enhancing expression of
factors such as RANKL (246). In contrast, when serum calcium levels are
adequate, 1,25(OH)2D3 may promote appropriate mineralization of bone (233).
Thus, the vitamin D endocrine system exerts pleiotropic effects on mineral ion
homeostasis and osteoblast and osteoclast activity to fine-tune the level of bone
density under a variety of physiological and pathophysiolgical states
Although neither 1,25(OH)2D3 or related compounds are currently approved for
treating osteoporosis in the United States, these compounds are widely used to prevent and treat osteoporosis throughout the world (247, 248). Furthermore, vitamin D3 is currently recommended as a dietary supplement in addition to any pharmacological treatment for all patients with decreased bone mass or osteoporosis (168). In fact, a recent meta-analysis demonstrated that high doses of vitamin D3 supplementation prevents hip fractures in the elderly (249).
However, these higher doses of vitamin D3 or 1,25(OH)2D3 required for maximal
improvement in bone density can cause toxic levels of hypercalcemia.
Consequently, more than 800 synthetic 1,25(OH)2D3 analogues have been
developed in attempt to preserve the favorable activities of 1,25(OH)2D3 while avoiding the side effects (250). One novel analogue, Ro-26-9228, protects against osteopenia in ovariectomized rats, but it does not increase serum calcium except at very high doses. These observations are potentially explained by the tissue-selective action of Ro-26-9228, which stimulates osteocalcin and
40 osteopontin expression in osteoblasts but does not affect calbindin D9K or plasma
membrane calcium pump expression in the intestine (251). Shevde et al. (252)
found that another analogue, 2-methylene-19-nor-(20S)-1α,25(OH)2D3 (2MD), potently stimulates bone formation in vitro and markedly improves bone mass in ovariectomized rats without dramatically increasing serum calcium. Such studies support the concept that more selective 1,25(OH)2D3 analogues will be useful
therapies for osteoporosis by enhancing bone mineral density without causing
toxic hypercalcemia.
Novel 1,25(OH)2D3-regulated genes in osteoblasts
As I have discussed, the effects of the 1,25(OH)2D3 on the bone are complex,
involving systemic changes in mineral ion concentrations, modulation of
osteoblast differentiation and mineralization, and stimulation of bone resorption.
However, despite these multi-faceted actions, relatively few 1,25(OH)2D3 target genes had been described in osteoblasts and other tissues. The purpose of the current project was to identify novel 1,25(OH)2D3-induced genes in osteoblastic
cells and to characterize their regulation and function in osteoblasts and in the
bone. By discovering global changes in gene expression, I had hoped to better
understand the multifaceted direct actions of 1,25(OH)2D3 as well as uncovering
novel functions for the target genes. To these ends, MG-63 human osteoblastic
cells were treated with 10 nM 1,25(OH)2D3 for 6 hours, and changes in mRNA
levels were measured using Affymetrix HU95A microarrays. Of more than
12,000 genes represented on the gene chip, 1,25(OH)2D3 treatment decreased
41 92 and increased 33 genes by more than 2.0-fold (Tables I-1 and I-2). We
further characterized three of the up-regulated genes, MN1 (Chapter II),
semaphorin3B (Chapter IV), and thrombomodulin (Chapter V), identified in our
microarray study and one additional gene, C/EBPβ (Chapter III), found through a separate gene expression profiling analysis performed by our collaborators (253).
42 Table I-1 Gene Expression Profiling Reveals Decreased Expression of 92
Genes in 1,25(OH)2D3-Treated Osteoblastic Cells.
Probe Set # % of Unigene # Description Control 510_g_at ~50∗ Hs.75862 SMAD, mothers against DPP homolog 4 (Smad4) 40441_g_at ~50 Hs.530412 PAI-1 mRNA-binding protein (PAI-RBP1) 1986_at ~50 Hs.513609 retinoblastoma-like 2 (p130) 36309_at ~50 Hs.41565 growth differentiation factor 8 (GDF 8) 34328_s_at ~50 Hs.3068 SWI/SNF related, matrix associated, actin dependent regulator of chromatin, subfamily a, member 3 (SMARCA3) 604_at ~50 Hs.194143 breast cancer 1, early onset (BRCA1) 1920_s_at ~50 Hs.79101 cyclin G1 33715_r_at ~50 Hs.191356 general transcription factor IIH, polypeptide 2, 44kDa 41089_at 50 Hs.343522 ATPase, Ca++ transporting, plasma membrane 4 (ATP2B4) 784_g_at 50 Hs.533440 WW domain containing E3 ubiquitin protein ligase 1 (WWP1) 1929_at 50 Hs.369675 angiopoietin 1 (ANGPT1) 1585_at ~48 Hs.118681 v-erb-b2 erythroblastic leukemia viral oncogene homolog 3 (ERBB3) 1423_at 48 Hs.462035 ubiquitin-conjugating enzyme E2G 1 (UBE2G1) 36922_at 48 Hs.226390 ribonucleotide reductase M2 polypeptide (RRM2) 36565_at 48 Hs.458365 zinc finger protein 183 (RING finger, C3HC4 type; ZNF183) 39695_at 48 Hs.527653 decay accelerating factor for complement (CD55, Cromer blood group system; DAF) 41123_s_at ~48 Hs.190977 ectonucleotide pyrophosphatase/phosphodiesterase 2 (autotoxin; ENPP2) 39627_at ~48 Hs.506309 early endosome antigen 1, 162kD (EEA1) 38100_at 48 Hs.529591 translocation protein 1 (TLOC1) 41788_i_at 48 Hs.52526 TSC22-related inducible leucine zipper 4a (TILZ4a) 1605_g_at 45 Hs.479853 EPH receptor A5 (EPHA5) 353_at 45 Hs.7370 phosphatidylinositol transfer protein, beta (PITPNB) 39100_at 45 Hs.124611 Sparc/osteonectin, cwcv and kazal-like domains proteoglycan (testican) (SPOCK) 31616_r_at 45 Hs.502328 CD44 antigen (homing function and Indian blood group system) 40621_at 45 Hs.406074 PRKC, apoptosis, WT1, regulator (PAWR) 1994_at 45 Hs.425104 activating transcription factor 2 (ATF2) 1719_at 45 Hs.280987 mutS homolog 3 (MSH3) 35458_r_at 45 Hs.17614 ATP-binding cassette, sub-family B (MDR/TAP), member 10 (ABCB10) 1648_at 45 Hs.120658 oncostatin M receptor (OSMR) 35209_at 44 Hs.28020 EPM2A (laforin) interacting protein 1 (EPM2AIP1) 33126_at 44 Hs.297304 glycosyltransferase 8 domain containing 1 (GLT8D1) 1628_at 44 Hs.220529 carcinoembryonic antigen-related cell adhesion molecule
∗ The tilda (~) indicates that the fold change is an approximation due to a low signal-to-noise ratio in one of the samples.
43 5 (CEACAM5) 33565_at 44 Hs.406687 thyroid stimulating hormone, beta (TSHB) 40431_at 42 Hs.16349 KIAA0431 31849_at 42 Hs.368282 KIAA0564 31665_s_at 42 Hs.378808 Eukaryotic translation initiation factor (eIF) 2A (eIF2A) 39366_at 42 Hs.303090 protein phosphatase 1, regulatory (inhibitor) subunit 3C (PPP1R3C) 32683_at 40 Hs.11355 thymopoietin (TMPO) 33556_at ~40 Hs.234961 Huntingtin interacting protein E (HYPE) 39806_at 40 Hs.292316 ASF1 anti-silencing function 1 homolog A (ASF1A) 38902_r_at ~40 Hs.425104 activating transcription factor 2 (ATF2) 33864_at 38 Hs.292265 zinc finger, MYND domain containing 11 (ZMYND11) 37552_at ~38 Hs.208544 potassium channel, subfamily K, member 1 (KCNK1) 40633_at 38 Hs.531029 KIAA1008 975_at 38 Hs.172052 polo-like kinase 4 (PLK4) 40447_at 37 Hs.268581 lipin 2 34910_s_at 37 Hs.203965 putative homeodomain transcription factor 2 (PHTF2) 40121_at 37 Hs.50308 huntingtin interacting protein 2 (HIP2) 38500_at 37 Hs.508765 transmembrane emp24 protein transport domain containing 7 (TMED7) 40430_at 36 Hs.405770 Transcribed locus, moderately similar to XP_372039.2 similar to hypothetical protein (L1H 3 region) 1159_at 36 Hs.536926 interleukin 7 (IL7) 37528_at 36 Hs.332422 aspartate beta-hydroxylase (ASPH) 35033_at ~36 Hs.519884 glucosaminyl (N-acetyl) transferase 2, I-branching enzyme (GCNT2) 460_at ~36 Hs.431043 PBX/knotted 1 homeobox 1 (PKNOX1) 40178_at ~34 Hs.143604 zinc finger and BTB domain containing 33 (ZBTB33) 40310_at ~34 Hs.519033 toll-like receptor 2 (TLR2) 1328_at ~34 Hs.523930 TNF receptor-associated factor 5 (TRAF5) 33502_at ~34 Hs.32769 mRNA full length insert cDNA clone EUROIMAGE 362430 41739_s_at 34 Hs.490203 caldesmon 1 (CALD1) 509_at 32 Hs.75862 SMAD, mothers against DPP homolog 4 (Smad4) 34662_at 32 Hs.476972 MYC induced nuclear antigen (MINA) 38677_at 32 Hs.352341 stress 70 protein chaperone, microsome-associated, 60kDa (STCH) 40993_r_at 31 Hs.413835 sin3-associated polypeptide, 30kDa (SAP30) 36708_at ~30 Hs.334873 carboxypeptidase M (CPM) 37143_s_at 29 Hs.88139 phosphoribosylformylglycinamidine synthase (FGAR amidotransferase; PFAS) 1139_at ~29 Hs.515018 guanine nucleotide binding protein (G protein), alpha 13 (GNA13) 34248_at ~29 Hs.445857 protease, serine, 12 (neurotrypsin, motopsin; PRSS12) 41317_at 28 Hs.508480 RAP2A, member of RAS oncogene family 34732_at 27 Hs.327736 kinesin family member 5B (KIF5B) 34909_at ~27 Hs.203965 putative homeodomain transcription factor 2 (PHTF2) 40579_at 26 Hs.443507 HIV-1 Rev binding protein (HRB) 31729_at ~26 Hs.502508 TAF13 RNA polymerase II, TATA box binding protein (TBP)-associated factor, 18kDa (TAF13) 1036_at ~26 Hs.311958 interleukin 15 (IL15) 943_at ~23 Hs.149261 runt-related transcription factor 1 (acute myeloid leukemia 1; aml1 oncogene; RUNX1) 1560_g_at ~22 Hs.518530 p21 (CDKN1A)-activated kinase 2 (PAK2) 35397_at ~22 Hs.504550 RAD51 associated protein 1 RAD51AP1)
44 33306_at ~22 Hs.514802 zinc finger protein 24 (KOX 17; ZNF24)/zinc finger protein 396 (ZNF396) 37486_f_at 22 Hs.356135 Meis1, myeloid ecotropic viral integration site 1 homolog 4 (MEIS4) 1931_at ~22 Hs.508423 ATP-binding cassette, sub-family C (CFTR/MRP), member 4 (ABCC4) 41290_at 21 Hs.503878 neural cell adhesion molecule 1 (NCAM1) 1990_g_at ~20 Hs.34012 breast cancer 2, early onset (BRCA2) 35494_at ~19 Hs.388400 Pantothenate kinase 3 (PANK3) 35731_at ~18 Hs.440955 integrin, alpha 4 (antigen CD49D, alpha 4 subunit of VLA- 4 receptor; ITGA4) 32862_at ~18 Hs.369188 RAS p21 protein activator 3 (RASA3) 32405_at 16 Hs.476882 thioesterase, adipose associated (THEA) 1606_at ~13 Hs.371218 EPH receptor A4 31682_s_at ~13 Hs.443681 chondroitin sulfate proteoglycan 2 (versican) 1101_at ~12 Hs.372840 amyloid beta (A4) precursor protein-binding, family B, member 1 (APBB1) 39531_at 11 Hs.335079 microtubule-associated protein 1B (MAP1B) 1812_s_at ~11 Hs.132966 Met proto-oncogene (hepatocyte growth factor receptor; MET) 39364_s_at ~11 Hs.303090 protein phosphatase 1, regulatory (inhibitor) subunit 3C (PPP1R3C) 1993_s_at ~11 Hs.194143 breast cancer 1, early onset (BRCA1)
45 Table I-2 Gene Expression Profiling Reveals Increased Expression of 33
Genes in 1,25(OH)2D3-Treated
Osteoblastic Cells.
Probe Set # Fold Unigene # Description Increase 34759_at 2 Hs.519469 Solute carrier family 30 (zinc transporter), member 1 (SLC30A1) 38505_at 2 Hs.449415 Eukaryotic translation initiation factor 2C, 2 (EIF2C2) 40401_at 2 Hs.473133 docking protein 5 (DOK5) 1052_s_at 2.2 Hs.440829 CCAAT/enhancer binding protein (C/EBP), delta (CEBPD) 37680_at 2.2 Hs.371240 A kinase (PRKA) anchor protein (gravin) 12 (AKAP12) 41083_at 2.2 Hs.99093 hypothetical protein MGC51082 1949_at ~2.4∗ Hs.369675 Angiopoietin 1 (ANGPT1) 41369_at ~2.4 Hs.501135 pancreatic lipase (PNLIP) 38195_at ~2.4 Hs.159918 PHD finger protein 14 (PHF14) 850_r_at ~2.5 Hs.471508 insulin receptor substrate 1 (IRS1) 34017_s_at 2.5 Hs.391464 ATP-binding cassette, sub-family C (CFTR/MRP), member 1 (ABCC1) 40448_at 2.5 Hs.534052 zinc finger protein 36, C3H type (ZFP36) 1005_at 2.5 Hs.171695 dual specificity phosphatase 1 (DUSP1) 575_s_at ~2.7 Hs.692 tumor-associated calcium signal transducer 1 (TACSTD1) 34150_at ~2.7 Hs.126195 UDP-N-acetylglucosamine:a-1,3-D-mannoside beta-1,4- N-acetylglucosaminyltransferase IV (HGNT-IV-H) 995_g_at 2.7 Hs.49774 protein tyrosine phosphatase, receptor type, M (PTPRM) 38585_at ~2.9 Hs.302145 hemoglobin, gamma G (HBG2) 31780_f_at 2.9 Hs.278457 Killer cell immunoglobulin-like receptor, three domains, long cytoplasmic tail, 1 (KIR3DL1) 32107_at 3.1 Hs.473894 chromosome 21 open reading frame 25 37393_at 3.3 Hs.250666 hairy and enhancer of split 1 (HES1) 37878_at ~3.4 Hs.25791 PARK2 co-regulated (PACRG) 1397_at ~3.7 Hs.502872 mitogen-activated protein kinase kinase kinase 11 (MAP3K11) 37090_at ~3.9 Hs.98848 a disintegrin and metalloproteinase domain 3a (cyritestin 1; ADAM3A) 34297_at ~4.5 Hs.406094 G protein-coupled receptor 37 (endothelin receptor type B-like; GPR37) 32143_at 5.6 Hs.253247 odd-skipped related 2 (OSR2) 33970_at ~5.9 Hs.73021 melanoma antigen family B, 1 (MAGEB1) 37279_at 6.4 Hs.345139 gene overexpressed in skeletal muscle (GEM) GTPase+ 37283_at 8.1 Hs.268515 meningioma-1 (MN1) 32025_at ~8.3 Hs.501080 transcription factor 7-like 2 (TCF7L2; hTCF-4) 33118_at 10 Hs.82222 semaphorin3B (SEMA3B) 33803_at ~14.8 Hs.2030 thrombomodulin
∗ The tilda (~) indicates that the fold change is an approximation due to a low signal-to-noise ratio in one of the samples. + Genes in italics have been confirmed by Northern blot analysis (see corresponding chapter or data not shown).
46 36253_at ~20.7 Hs.530479 bone gamma-carboxyglutamate (gla) protein (osteocalcin; BGLAP)# 38022_s_at ~23.5 Hs.434248 plectin 660_at ~39.9 Hs.89663 1,25-dihydroxyvitamin D3 24-hydroxylase (24OHase)
# Genes in bold are well-characterized 1,25(OH)2D3-induced genes.
47 Figure I-1
Metabolism and mineral homeostatic functions of the vitamin D endocrine
system.
Bioactive 1,25(OH)2D3 is generated by sequential hydroxylations of its precursor
vitamin D3 in the liver and the kidney. 1,25(OH)2D3 operates in a negative
feedback loop by inducing expression of the catabolic enzyme 24OHase and by inhibiting expression of the anabolic enzyme 1αOHase. In response to low serum calcium, PTH is produced and stimulates 1αOHase expression in the
kidney, promotes calcium mobilization from the bone, and increases calcium
reabsorption from the kidney. 1,25(OH)2D3, in turn, induces calcium absorption in the intestine and calcium release from the skeleton.
48 UV light Heat
Diet Vitamin D3 + PTH
25-OHase 25(OH)D3 1αOHase KIDNEY - TARGET ORGANS + (including 24OHase 1,25(OH) D kidney) 2 3
24,25(OH)2D3 1,24,25(OH)3D3
excreted metabolites
Serum Calcium and Phosphate
49 Figure I-2
Domain structure of VDR.
Functional domains of VDR include: A/B, amino-terminal region; the DBD, DNA-
binding domain showing two zinc finger modules (Zn); and the LBD, ligand- binding domain including the long insertion domain and helix 12 encompassing the activation function-2 (AF-2).
50 LBD
A/B DBD insertion
domain AF-2 hinge 1 24 89 115 157 220 409 427
51 Figure I-3
The vitamin D receptor undergoes a conformational change upon binding hormone.
1,25(OH)2D3 (ligand) associates with VDR and induces a conformational change in helix 12 of the AF-2 domain. Helix 12 folds over to trap the ligand in the binding pocket. This change also creates a hydrophobic cleft or surface on VDR comprised of helices H3, H4, H5, and H12. LXXLL motifs in coactivator proteins use this cleft for docking to VDR.
52
H4
H5
H3 H12
+ ligand
H4
H5 H3
LXXL
H12
53 Figure I-4
Schematic of the conserved domains in SRC family members.
The three SRC proteins are similar in several conserved domains. bHLH, basic helix-loop-helix domain; PAS, region characteristic of the Per/Arnts/Sim family of transcription factors; NR boxes, nuclear receptor interaction domains containing the LXXLL motifs; CBP interaction, region that interacts with the CREB-binding protein; Q rich, region rich in glutamine residues; HAT, histone acetyltransferase domains in SRC-1 and SRC-3.
54 NR CBP bHLH PAS boxes interaction Q rich HAT SRC-1
SRC-2
SRC-3
55 Figure I-5
Model of the DRIP complex interacting with the liganded VDR.
DRIP205/TRAP220 (205) uses an LXXLL NR box motif for docking the complex to VDR.
56 10 240 36 17 cdk8 95 130 230 33 97 150 205 78 34 70 100 D VDR
57 Figure I-6
Domain structure of NCoA-62/SKIP.
LPXP, a strictly conserved motif of unknown function, SNW, region containing a central conserved serine, asparagines, and tryptophan as well as the receptor interaction domain (RID); TAD-1, transcriptional activation domain; NLS, nuclear localization sequence.
58 RID 274 342 LPXP NLS
SNW TAD-1
1 174 339 432 530 536
NCoA62/SKIP
59 Figure I-7
Model of temporal association of coactivators during VDR-mediated transcription.
The liganded (D) VDR-RXR heterodimer associates with VDREs in the promoters of target genes. First, SRCs and p300/CBP are recruited to the promoter and acetylate (Ac) histones, resulting in a more open chromatin template. Next, SRCs and p300/CBP disassociate allowing for binding of the
DRIP complex and entry of the core transcriptional machinery. Finally, NCoA-
62/SKIP associate with the complex and tethers the splicing machinery to the activated promoter, thereby allowing for immediate splicing of the nascent transcript.
60 LXXLL D SRCs
RXR VDRVDR p300
VDRE Ac
p300
SRCs
NCoA62 /SKIP LXXLL D DRIP 205 RXR VDR RNA Pol II VDRE D TBP
Ac Ac AcAc Ac Ac Ac Ac
spliceosome NCoA62 /SKIP LXXLLDRIP D 205 RXR VDR
VDRE TBP RNA Pol II
Ac Ac AcAc Ac Ac Ac Ac
61
Figure I-8
Long bones are formed by endochondral ossification.
A, Anatomy of a mature long bone. B, Schematic of endochonral ossification.
Undifferentiated mesenchymal cells coalesce at the future site of the bone. The innermost cells differentiate into hypertrophic chondrocytes and deposit mineralized cartilage. The cells in the periphery differentiate directly into osteoblasts and form a bone collar, which will eventually form dense cortical bone. Vascular invasion recruits osteoclasts, chondroclasts, osteoblasts, and hematopoietic stem cells. While the osteoclasts and chondroclasts degrade the cartilage tissue, osteoblasts replace the cartilage with mineralized trabecular bone. Hematopoietic stem cells populate the space between trabeculae to form bone marrow. Some chondrocytes remain on the epiphyseal ends of the ossification center and organize into growth plates, allowing for elongation of the bone tissue as the organism grows. RC, resting chondrocytes, PC, proliferating chondrocytes, and HC, hypertrophic chondrocytes.
62 A metaphysis cortical epiphysis trabecular bone bone
marrow diaphysis
B
hypertrophic migrating osteoblasts, chondroclasts, chondrocytes osteoclasts, and hematopoetic cells RC
PC
HC
mesenchymal cartilage vascular bone condensation template invasion replacement
63 CHAPTER II
THE 1,25(OH)2D3-REGULATED TRANSCRIPTION FACTOR MN1 STIMULATES VDR-MEDIATED TRANSCRIPTION AND INHIBITS OSTEOBLASTIC CELL PROLIFERATION∗
INTRODUCTION
The vitamin D endocrine system is crucial for maintaining calcium and phosphate
homeostasis and protecting skeletal integrity (254, 255). 1,25(OH)2D3 is the bioactive hormonal form of vitamin D. This hormone functions through the vitamin D receptor (VDR), a member of the nuclear hormone receptor family, to regulate transcription of target genes. 1,25(OH)2D3 induces expression of
various calcium binding and transport proteins in the intestine to stimulate active
calcium uptake, thus preserving normocalcemia and, indirectly, maintaining bone
mineralization. However, 1,25(OH)2D3 also acts directly on osteoblasts, the
resident bone-forming cells of the skeleton, to inhibit proliferation, modulate
differentiation, and regulate mineralization of the extracellular matrix (256).
Upon binding 1,25(OH)2D3, VDR heterodimerizes with retinoid X receptor (RXR)
and associates with vitamin D response elements (VDREs) in the promoter
regions of target genes (257). The active VDR/RXR heterodimer forms multiple
protein complexes with general transcription factors and coactivator proteins,
factors that interact with nuclear receptors and enhance their transcriptional
activity. A variety of coactivators for VDR have been described, including the
∗ Adapted from Sutton AL, Zhang X, Ellison TI, MacDonald PN 2005 The 1,25(OH)2D3-regulated transcription factor MN1 stimulates VDR-mediated transcription and inhibits osteoblastic cell proliferation. Mol Endocrinol, in press.
64 steroid receptor coactivator (SRC) proteins, the DRIP (VDR-interacting proteins)
complex, and NCoA-62/ski-interacting protein (SKIP) (258). The SRC family
represents the first nuclear receptor coactivators identified and includes SRC-1
(46), SRC-2 (GRIP1/TIF2; (52, 53)), and SRC-3 (RAC3/ACTR/AIB1;(54-56)).
SRCs are thought to stimulate nuclear receptor activity by recruiting histone
acetyl-transferases (HATs), such as cAMP response element binding protein
(CREB)-binding protein (CBP)/p300, to the promoter regions of target genes (56,
57, 60). SRCs also possess intrinsic HAT activity (61) and, in concert with
CBP/p300, modify histones thereby allowing the transcriptional machinery better
access to regulatory regions to initiate transcription. DRIP is a multimeric
complex that includes more than 10 proteins and is nearly identical to the
transcriptional coactivators thyroid receptor activating proteins (TRAP) and the
mammalian Mediator complex (64, 65). Since DRIP interacts with the RNA
polymerase II holoenzyme (72), it may serve as a bridge between liganded
VDR/RXR and the basal transcriptional machinery. Finally, NCoA-62 or ski-
interacting protein (SKIP) represents a distinct VDR coactivator (73) that may
couple transcriptional activation to mRNA splicing (78). SRCs and DRIP205, the
anchoring subunit of the DRIP complex, interact with liganded VDR through
LXXLL motifs, hydrophobic domains that interact with a complementary hydrophobic cleft in the nuclear receptor ligand-binding domain (LBD) (48, 51).
While these LXXLL motifs are essential for SRCs and DRIP205 to interact with
VDR, other coactivators interact with the receptor through different mechanisms.
For example, NCoA-62/SKIP associates with VDR through a central domain that
65 does not contain LXXLL sequences (77). All three classes of coactivators,
acting through distinct mechanisms, are necessary for 1,25(OH)2D3-mediated
transcription (64, 77). The complexity of VDR-activated transcriptional processes
is highlighted by the ever-expanding number of protein factors that are involved
in this mechanism.
The cellular consequences of 1,25(OH)2D3-mediated transcriptional activation
include inhibition of proliferation and stimulation of differentiation in many tissues
(259, 260). In fact, a number of synthetic 1,25(OH)2D3 analogs have been
developed to treat cancer and other hyper-proliferative diseases. How
1,25(OH)2D3 accomplishes this growth inhibition is not fully understood, but likely
involves alterations in the expression of numerous growth factors, cell-cycle
related proteins, transcription factors, and other genes (259). Two well-
established target genes of 1,25(OH)2D3 are the cell cycle-dependent kinase
WAF1/cip1 kip1 (cdk) inhibitors p21 (261) and p27 (262). 1,25(OH)2D3 stimulates
expression of these genes resulting in cell cycle accumulation at the G1/S checkpoint (263). GADD45α, another cell cycle regulator and DNA-damage repair protein, is also induced by 1,25(OH)2D3 (264, 265) and causes growth
arrest at G2/M (265). Further identification of potential 1,25(OH)2D3 target genes
involved in cell proliferation and differentiation has accelerated recently with the
advent of microarray technology (266), but the functional implications of these
genes remain largely unexplored.
66 Here, we identify and characterize MN1 (meningioma-1) as a novel target of
1,25(OH)2D3 in osteoblasts. MN1 was first identified as a gene disrupted in some meningiomas as part of a balanced translocation t(4;22) (267) and as part of a fusion with the ETS domain transcription factor TEL in some myeloid leukemias
(268). Sequence analysis reveals that MN1 exhibits several attributes indicative of a transcription factor, including an N-terminal nuclear localization signal, proline-rich sequences, and two polyglutamine stretches (267). MN1 localizes to the nucleus and activates transcription of a Moloney sarcoma virus long terminal repeat (MSV-LTR) reporter gene (269). Subsequent studies showed that MN1 associates with a retinoic acid-responsive element (RARE) in the MSV-LTR and that MN1 stimulates RAR-dependent transcription of the MSV-LTR, a previously unrecognized RAR-responsive promoter (270). These studies support a role for
MN1 in transcriptional regulation. However, beyond these initial reports, there are currently no studies addressing the role of MN1 in other nuclear receptor pathways, the factors that control expression of MN1, or the potential function of
MN1 in cellular events such as proliferation or differentiation.
In the current study, we demonstrate that 1,25(OH)2D3 strongly increased MN1 mRNA levels in osteoblastic cells. We provide data supporting a coactivator role for MN1 in VDR/1,25(OH)2D3-mediated transcription through a mechanism that involves the VDR LBD and SRC coactivators. Finally, expression of MN1 in osteoblastic cells potently inhibited cellular proliferation by reducing the proportion of cells entering S-phase of the cell cycle, uncovering an important
67 role for MN1 in regulating osteoblastic cell growth. This study is the first
demonstration that MN1 stimulates VDR transcriptional activity and inhibits
osteoblastic cell proliferation.
MATERIALS AND METHODS
Cell culture. MG-63 human osteosarcoma cells were maintained in
modified essential media supplemented with 10% fetal bovine serum. COS-7
green monkey kidney cells were maintained in Dulbecco’s modified essential
media supplemented with 10% bovine calf serum. For experiments using
1,25(OH)2D3 compounds, cells were grown in media supplemented with
charcoal-stripped bovine calf serum for three days prior to the experiment.
Northern blot analysis. mRNA was isolated from MG-63 cells with the
FastTrack system (Invitrogen, Carlsbad, CA) according to the manufacturer’s
instructions. mRNA was separated on a formaldehyde/agarose gel and
transferred to a Duralon membrane (Stratagene, La Jolla, CA) by capillary action.
α32P-labeled probes were synthesized using the Prime-A-Gene kit (Promega,
Madison, WI) according to the manufacturer’s instructions and hybridized to the
blots using standard methods. After exposure to X-ray film, blots were
quantitated by densitometry.
Plasmids. The reporter plasmids (VDRE)4-TATA-luciferase (luc), (VDRE)4-TK-
luc, (GRE)2-TK-luc, (RARE)4-TK-luc, and (GAL4)5-TK-luc were generated by
68 subcloning the promoter regions from (VDRE)4-TATA-GH (77), (VDRE)4-TK-GH
(271), (GRE)2-TK-GH (272), (RARE)4-TK-GH (73), and (GAL4)5-TK-GH (273), respectively, into pGL3 (Promega). The rat 24-hydroxylase promoter-luciferase reporter plasmid (r24OHase-luc) was constructed by cloning the PCR-amplified promoter sequence (-369 to +113) from ROS 17/2.8 cells into pGL3. pSG5-VDR
(271), pSG5-GAL4-VDR (16), pSG5-RARα (73), pSG5-GR (274), pCR-SRC-1
(275), and pSG5-SRC-2/GRIP1 (276) have been previously described. Plasmids encoding GAL4 DBD fusions with all other nuclear receptor LBDs were constructed by PCR amplification of the LBDs from either a human fetal brain or human placental cDNA library (BD Biosciences Clontech, Palo Alto, CA) and ligating them in frame with the GAL4 DBD. CMVTAG2B-MN1 was constructed by subcloning the MN1 cDNA from pSCTOP-MN1 (269) in frame with the FLAG epitope tag in the CMVTAG2B vector (Stratagene). CMVTAG2A-RORβ was created by PCR amplification of the full-length RORβ coding sequence from a human fetal brain library (BD Biosciences Clontech) and ligating it in frame with
FLAG. The dual expression vector pCMS-EGFP-MN1 drives expression of
EGFP and MN1 from two separate promoters on a single plasmid. This plasmid was constructed by subcloning FLAG-tagged MN1 from CMVTAG2B-MN1 into pCMS-EGFP (BD Biosciences Clontech). pCMS-EGFP-SRC1 was created by subcloning full-length human SRC1 from pCR-SRC1 (275) into pCMS-EGFP.
Transient transfection assays. COS-7 cells were seeded at a density of
70,000 per well of six-well plates and transfected by the standard calcium
69 phosphate precipitation method as previously described (271). CMV-renilla
(Promega) was co-transfected with other plasmids to normalize for transfection
efficiency. Plasmids encoding MN1 or other coactivators and their respective
vector controls were transfected in molar balance in all experiments. Cells were
dosed with the indicated amounts of ligand for 24 hours. Cell lysates were
isolated after 24 hours of ligand stimulation, and the luciferase and renilla
reporter gene activities were determined using a commercial dual luciferase
assay kit (Promega) according to the manufacturer’s recommendations. An LMax
microplate luminometer (Molecular Devices, Sunnyvale, CA) was used to
measure luciferase activity. Data presented the mean of duplicate or triplicate
wells -/+ standard deviation of a representative experiment that has been
repeated at least three times.
MG-63 cell growth assay. MG-63 cells were plated at a density of 8 X 104 cells/well of a six well dish in media containing charcoal-stripped serum. Cells were treated with either ethanol vehicle control or 10 nM 1,25(OH)2D3 for up to
five days. Viable cells were quantified by counting trypan blue-excluding cells
with a hemocytometer. Data presented represent the mean of triplicate wells -/+
standard deviation of a representative experiment performed twice.
MG-63 stable pool growth assay. MG-63 cells were plated the day prior to
transfection at a density of 1.4 X 106 cells/150 mm plate. Cells were transfected
with CMVTAG2B, CMVTAG2B-MN1, or CMVTAG2A-RORβ using Fugene
70 (Roche Applied Science, Indianapolis, IN). After 48 hours, cells were trypsinized
and 250,000 cells/well were plated in G418-containing media to select for
transfected cells. After 7 days, resulting colonies were stained with 0.25% crystal
violet in 50% methanol. After extensive washing in PBS, colonies were air-dried
for at least one hour and photographed. Dye was extracted in 100 μl of 0.1 M
sodium citrate in 50% ethanol and quantified spectrophotometrically at 540 nm
using a Versimax microplate reader (Molecular Devices). Data presented
represent the mean of triplicate wells -/+ standard deviation of a representative experiment performed three times.
BrdU incorporation. 22,000 MG-63 cells were seeded per chamber of a
four-well Lab-Tek chamber slides (Nalge Nunc International, Rochester, NY)
coated with poly-L-lysine and bovine calf serum to promote cell attachment.
Cells were transfected with pCMS-EGFP, pCMS-EGFP-MN1, or pCMS-EGFP-
SRC1 with Fugene (Roche Applied Science) and allowed to express for 24 hours. Cells were pulse-labeled with 10 μM 5-bromo-2'-deoxyuridine (BrdU) for one hour, fixed in 4% paraformadehyde, and permeabilized in PBS containing
3% BSA and 0.3% Triton X-100. BrdU incorporation sites were exposed by treating cells with 100 U/ml of bovine pancreatic DNase I (Amersham
Biosciences, Piscataway, NJ). Cells were immunostained with an anti-BrdU mouse monoclonal antibody (BD Biosciences) followed by a goat anti-mouse secondary antibody conjugated to Alexa Fluorophore 594 (Molecular Probes,
Eugene, OR). Nuclei were counterstained with 4',6-diamidino-2-phenylindole
71 dihydrochloride (DAPI; Roche Applied Science). 100-150 EGFP-positive cells
were counted and scored for BrdU positivity in each sample. Data represent the
mean percentage of BrdU-positive cells of three independent experiments -/+
standard deviation.
RESULTS
Characterization of MN1 as a 1,25(OH)2D3 target gene in osteoblastic cells.
Microarray analysis was used as an initial screen to identify 1,25(OH)2D3- regulated genes in MG-63 cells, a human osteoblastic cell line. In this analysis,
MN1 was induced approximately 8-fold in response to a 6 hour treatment with 10 nM 1,25(OH)2D3 (see Introduction). To validate the microarray results in MG-63
cells, Northern blot analysis was performed. As shown in Figure 1, 1,25(OH)2D3 increased MN1 mRNA levels in a time- and dose-dependent manner. This increase was apparent as early as 3 hours and was maximal at 12 hours (Fig.
1A). As little as 0.1 nM 1,25(OH)2D3 stimulated MN1 expression, and mRNA
levels continued to rise up to 100 nM 1,25(OH)2D3 (Fig. 1B). This effect was
specific for 1,25(OH)2D3, since cholicalciferol (vitamin D3), an inactive precursor
molecule of 1,25(OH)2D3, and 24,25(OH)2D3, a metabolite of 1,25(OH)2D3, had little effect on MN1 mRNA levels (Fig. 1A and data not shown). RNA synthesis was required for this response since inhibition of transcription by actinomycin D abolished the 1,25(OH)2D3-mediated induction of MN1 (Fig. 2A). Further, when
de novo protein synthesis was inhibited with cycloheximide, 1,25(OH)2D3-
mediated induction of MN1 mRNA was preserved (Fig. 2B). However, the basal
72 expression of MN1 was enhanced upon cycloheximide treatment, diminishing the
extent of MN1 induction by 1,25(OH)2D3. These data also suggest that synthesis
of other proteins is required for maximal induction of MN1 mRNA. Taken
together, these data imply that 1,25(OH)2D3 increases steady-state levels of MN1
mRNA by a process that, in part, involves active transcription and does not require on-going protein synthesis.
MN1 augments VDR-mediated transcription. Given recent evidence indicating
that MN1 stimulates RAR-mediated transcription of MSV-LTR (270), we tested
whether MN1 modulates VDR-mediated transcription. COS-7 cells were
transfected with a VDR expression vector and a 1,25(OH)2D3-responsive reporter
gene composed of 4 copies of a vitamin D response element (VDRE) and a
minimal TATA promoter fused upstream of firefly luciferase. Transfection of an
expression plasmid encoding MN1 augmented 1,25(OH)2D3-mediated expression
of the reporter gene approximately 6-7 fold (Fig. 3A). MN1 also stimulated
1,25(OH)2D3-mediated induction of a reporter gene driven by the 24-hydroxylase
promoter, a native 1,25(OH)2D3-responsive regulatory sequence (data not
shown). The effect of MN1 was both VDR- and 1,25(OH)2D3-dependent since
MN1 expression had minimal effects on basal reporter gene activity in the absence of VDR (Fig. 3B) and in the absence of 1,25(OH)2D3 (Fig. 3A). MN1
also enhanced the ligand-dependent transcriptional activity of a fusion protein
composed of the GAL4 DNA-binding domain (DBD) and the ligand-binding
domain (LBD) of VDR (Fig. 3C). Since the LBD of VDR was sufficient for the
73 increase in transcriptional activation, it is likely that MN1 acts through the VDR
LBD to stimulate VDR-mediated transcription.
MN1 selectively stimulates other nuclear receptors. To test the putative role of
MN1 in other nuclear receptor pathways, MN1 was cotransfected along with
plasmids encoding the GAL4 DBD fused to the LBDs of two other nuclear
receptors that function as RXR heterodimers. The activity of these fusion
proteins was measured by activation of a GAL4-responsive reporter gene. As
shown in Figure 4A, MN1 selectively stimulated the VDR LBD. In addition to
ligand-activated nuclear receptors, we also tested whether MN1 impacts the
transcriptional activity of the ROR orphan nuclear receptors. MN1 modestly
stimulated RORγ activity but did not enhance RORα or RORβ activity (Fig. 4B).
Although MN1 failed to activate the LBD of RARα (Fig. 4A), it did augment the
transcriptional activity of the full-length RARα on a synthetic RARE-driven
reporter gene (Fig. 4C). In contrast, MN1 inhibited the transcriptional activity of
the glucocorticoid receptor (GR). Collectively, these data indicate that MN1
selectively stimulates the transcriptional activity of VDR, RAR, and potentially
RORγ. Furthermore, it is likely that the mechanisms underlying MN1 transactivation are distinct among these nuclear receptors since only the VDR
LBD is required for activation by MN1 whereas the full-length RARα is necessary for MN1-mediated stimulation.
74 MN1 synergizes with SRCs. The SRC proteins are well-characterized
coactivators of VDR and other nuclear receptors (277). To test whether MN1 synergizes with SRCs to stimulate VDR-mediated transcription, COS-7 cells were transfected with VDR, a VDRE-driven reporter gene, and MN1, either alone or in combination with SRC-1 or SRC-2 (GRIP1/TIF2). SRC-1 and SRC-2 both stimulated the VDR-mediated transcription by approximately 3- and 4-fold, respectively (Figs. 5A and 5B). MN1 alone similarly stimulated VDR activity by about 2-3-fold (Figs. 5A and 5B). However, MN1 and SRC-1 together
augmented VDR-mediated transcription by 20-fold, while MN1 and SRC-2 stimulated VDR transactivation by 16-fold. These data suggest that MN1 cooperates with the SRCs to synergistically stimulate VDR-mediated
transcription.
MN1 inhibits osteoblastic cell proliferation. In agreement with previous
studies (278, 279), 1,25(OH)2D3 inhibited the proliferation of MG-63 osteoblastic
cells in a time-dependent manner (Fig. 6A). MN1 was first described as a gene
disrupted in some meningiomas and leukemias (267, 268, 280), suggesting that
it functions as a tumor suppressor. Given these data, we hypothesized that MN1
may be one of the 1,25(OH)2D3-regulated genes responsible for its growth
inhibition. Since we were unable to isolate stable clones that maintain elevated
MN1 expression, we chose to use a short-term stable pool approach to study the
effect of MN1 expression on cell growth (281). MG-63 cells were transfected
with empty vector, MN1, or RORβ as an irrelevant control plasmid. Stable
75 transfectants were selected and grown in G418-containing media for one week, and resulting colonies were stained with crystal violet. Cells transfected with
MN1 formed much smaller colonies than those transfected with empty vector
(Fig. 6B). Spectrophotometric quantification of the crystal violet staining as a measure of total cell number revealed that forced expression of MN1 resulted in a greater than 90% decrease in cell growth (Fig. 6C). In contrast, transfection of
RORβ had no effect on colony size or total cell number (Figs. 6B and 6C), indicating that the effect observed with MN1 expression was unlikely to be caused by non-specific toxicity due to overexpression of a nuclear protein.
Transfection efficiency, as measured by luciferase activity of a co-transfected reporter gene plasmid, was similar among all three groups (data not shown).
These data indicate that MN1 dramatically inhibits the growth of osteoblastic cells.
MN1 decreases S-phase entry in osteoblastic cells. To determine if the MN1- mediated decrease in cell proliferation is due to alterations in cell cycle progression, MG-63 cells transiently expressing MN1 were analyzed for BrdU incorporation as a measure of S-phase entry (Fig. 7A and 7B). To easily identify transfected cells, MN1 was expressed using a dual expression vector that drives transcription of EGFP and MN1 from two separate promoters on a single plasmid. Whereas 38% of MG-63 cells transfected with the EGFP vector alone were BrdU-positive, only 13% of cells transfected with MN1 were BrdU-positive
(Fig. 7B). The growth inhibitory effect was specific for MN1 since expression of
76 another VDR coactivator, SRC-1, did not affect cell proliferation (Fig. 7A and 7B).
These data indicate that MN1 expression inhibits osteoblastic cell proliferation by
slowing entry into the S-phase of the cell cycle.
DISCUSSION
The VDR and 1,25(OH)2D3 control a variety of biological process in a diverse
array of tissues. One of the most important effects of 1,25(OH)2D3 is to stimulate
active calcium transport across the intestinal epithelium to maintain
normocalcemia and preserve bone mineralization (254, 255, 257). In addition to
these systemic effects on mineral ion homeostasis, 1,25(OH)2D3 also acts
directly on osteoblasts to control proliferation, differentiation, and mineralization.
Beyond a handful of target genes, such as p21WAF1/cip1, p27kip1, and GADD45,
little is known about the mechanisms through which 1,25(OH)2D3 inhibits proliferation of target cells. Microarray analysis has led to an expansion of our knowledge of 1,25(OH)2D3-regulated genes in a number of cell types (198, 266,
282-287). The precise molecular and cellular functions of these novel target
genes are just now being uncovered. Functional characterization of
1,25(OH)2D3-regulated genes is essential to understand the mechanisms
underlying the pleiotropic effects, including growth regulation, of 1,25(OH)2D3. In this report, we show that MN1 is a novel 1,25(OH)2D3-induced gene in
osteoblastic cells and that MN1 augments VDR-mediated transcription and
potently inhibits osteoblastic cell growth.
77 MN1 was revealed as a 1,25(OH)2D3-induced gene in MG-63 osteoblastic cells in our microarray studies. Another recent microarray study also showed that MN1 mRNA expression was increased by the synthetic 1,25(OH)2D3 analog EB1089 in squamous cell carcinoma cells (266). Although the regulation of MN1 expression and its functional roles were not addressed in this global expression profiling study, together our two studies clearly point to MN1 as a 1,25(OH)2D3-induced
gene in multiple 1,25(OH)2D3 target cells. In agreement with our microarray
results, Northern blot analysis confirmed that 1,25(OH)2D3 increased MN1 mRNA levels in a time- and dose-dependent manner in osteoblastic cells (Fig. 1).
1,25(OH)2D3 likely induces MN1 expression through a transcriptional mechanism
since inhibition of transcription with actinomycin D completely abolished the
increase in MN1 mRNA observed with 1,25(OH)2D3 (Fig. 2A). Further, it appears
that protein synthesis is not necessary, although may enhance, 1,25(OH)2D3- mediated induction of MN1 (Fig. 2B). These data suggest that VDR directly regulates MN1 expression by binding to and activating the promoter region of
MN1. Although the putative promoter of MN1 does not contain any canonical
VDREs, the sequence contains many nuclear receptor response element half- sites that may support binding of the VDR/RXR heterodimer. Further studies, including promoter analysis, are required to dissect the molecular details of the
1,25(OH)2D3-mediated induction of MN1 expression.
MN1 is a nuclear protein with transcriptional activity (269) and its transcriptional
targets and mechanisms of actions are currently being uncovered (270). The
78 present study shows that MN1 stimulates VDR-dependent transcriptional activation. This stimulation occurs through the LBD of VDR since MN1 increased the transcriptional activity of a GAL4 DBD-VDR LBD chimeric protein (Fig. 3C).
We did not detect a similar activation of GAL4 DBD-RAR LBD (Fig. 4A), but did observe an increase in the transcriptional activation of full-length RARα by MN1 using a canonical RARE-driven reporter gene (Fig. 4C). This corroborates and extends a previous study that demonstrated that MN1, cooperating with RAR, activates the MSV-LTR, a previously unrecognized RAR-responsive promoter
(270). It is not surprising that MN1 augments the activity of both VDR and RAR since these two nuclear receptors are structurally related and both utilize RXR as their heterodimeric partner (11). However, MN1 likely enhances the transcriptional activity of these two nuclear receptors through distinct mechanisms. Whereas MN1 stimulates VDR through its LBD, RARα activation does not occur exclusively through the LBD and potentially requires an intact amino-terminal region (the activation function-1) and/or DBD of the receptor.
Further, these data demonstrate that MN1 is not a general nuclear receptor coactivator; rather, it selectively stimulates VDR, RAR, and, to a lesser extent,
RORγ activity, inhibits GR activity, and does not affect the transactivation potential of the other nuclear receptors analyzed in this study. While the exact mechanism underlying the transcriptional stimulation of nuclear receptors is not known, the current study suggests that MN1 cooperates with both SRC-1 and
SRC-2 in a synergistic manner (Fig. 5). Coupled with the previous report that showed that MN1 synergizes with SRC-3 in RAR-dependent transcription (270),
79 these data provide further evidence of a functional link between MN1 and the
SRC family of coactivators.
Although it is clear from our studies and others (269, 270) that MN1 is a
transcription factor that stimulates nuclear receptor-dependent transactivation,
the cellular consequences of this activity previously had not been explored.
Gene expression profiling has revealed that transforming growth factor-β (TGF-
β), which is growth inhibitory in epithelial cells, induces expression of MN1 in breast cancer cells (288), untransformed breast epithelial cells, keratinocytes, and lung epithelial cells (289). Likewise, the current study and a recent microarray analysis (266) demonstrate that MN1 expression is induced by
1,25(OH)2D3 and its synthetic analog, which inhibit proliferation in a wide array of
tissues. These data, coupled with the clinical findings that MN1 is disrupted in
some meningiomas and leukemias (267, 268, 280), point to a role for MN1 in
modulating cell proliferation. However, until now, this concept has not been
directly tested. In the current study, we present the first experimental evidence
that MN1 negatively regulates cell growth. Stable expression of MN1 in MG-63
osteoblasts results in a greater than 90% reduction in cell growth (Fig. 6B and
6C). This anti-proliferative effect is likely due to decreased entry into S-phase as
MN1-transfected cells display a reduction in BrdU incorporation (Fig. 7).
Together these data indicate that MN1 inhibits osteoblastic cell proliferation by
slowing cell cycle progression. It is likely that MN1 regulates cell growth in a
variety of other tissues since it is mutated in tumors, such a meningiomas and
80 leukemia, and is induced by the growth inhibitory factors TGF-β and EB1089 in
numerous cell types. Evaluation of the potential anti-proliferative activity of MN1
in these other tissues will be important in understanding whether MN1 acts in a
cell type selective manner or as a more global regulator of cell growth.
The mechanism of this growth suppression by MN1 is not clear, but, given the
transcriptional activity of MN1, likely involves regulating expression of
downstream targets involved in cell cycle progression and proliferation.
Although most studies suggest that 1,25(OH)2D3/VDR directly regulates the
expression of cell cycle regulatory genes such as p21WAF1/cip1 (261) and p27kip1
(262), 1,25(OH)2D3-mediated induction of GADD45α expression is partially
blocked by cycloheximide (264), suggesting that synthesis of other transcription
factors are involved in this response. It is possible that 1,25(OH)2D3 increases
MN1 levels and that VDR and MN1 cooperate to regulate the expression of genes such as GADD45α that inhibit cell proliferation. However, since MN1 expression inhibits cell growth in the absence of 1,25(OH)2D3 (Figs. 6 and &7), it
is unlikely that the growth inhibitory effects of MN1 are completely dependent on liganded VDR. Further, given that SRC-1 expression did not affect cell proliferation (Fig. 7), it is unlikely that MN1 suppresses cell growth by acting as a general VDR coactivator. MN1 may modulate cell proliferation through a number of transcriptional mechanisms given that it is induced by more than one growth regulatory signal. Further studies aimed at understanding whether MN1
81 modulates the expression or activity of cell cycle regulators are required to
delineate the molecular details underlying the growth inhibitory effects of MN1.
In summary, we have demonstrated that MN1 is a novel transcriptional target of
1,25(OH)2D3 in osteoblastic cells. 1,25(OH)2D3 appears to directly regulate the
expression of MN1 mRNA through a transcriptional mechanism. Our studies
uncover both a molecular and cellular function of MN1 in regulating VDR- and
RAR-mediated transcription by synergizing with SRC proteins and in potently
inhibiting osteoblastic cell proliferation by decreasing S-phase entry. Thus, VDR
and MN1 may be involved in a positive feed-back circuit in which 1,25(OH)2D3 stimulates expression of MN1, and MN1, in turn, augments VDR-mediated transcription.
Note added in proof
While this manuscript was in press, a mouse model with a targeted deletion in
MN1 was described (290). MN1-deficient mice die in the perinatal period presumably due to a cleft palate defect preventing suckling. A closer examination of the skeleton of MN1 knockout embryos revealed dramatic defects in numerous cranial bones formed by intramembraneous ossification but normal development of the appendicular and axial bones. Some cranial bones did not form at all, and several were either hypoplastic or delayed in development and ossification. Although the mechanisms underlying these defects have not been examined, this study provides in vivo evidence for the essential role of MN1 in
82 intramembranous ossification and skull development. Coupled with the findings in the current study that indicate that MN1 regulates osteoblastic cell proliferation, the phenotype of the MN1 knockout mice suggests that this protein controls osteoblast differentiation or mineralization and provide the groundwork for further investigation into the molecular details of MN1 functions in osteoblasts in vivo and in vitro.
ACKNOWLEDGEMENTS
We gratefully acknowledge Dr. Ellen Zwartkoff for providing pSCTOP-MN1, Dr.
Bert O’Malley for providing pCR-SRC-1, and Dr. Michael Stallcup for providing pSG5-SRC-2/GRIP1. We would also like to thank Dr. Milan Uskokovic for supplying 1,25(OH)2D3.
83 Figure II-1
1,25(OH)2D3 induces MN1 expression in a time- and dose-dependent
manner.
A, MG-63 cells were treated for the indicated times with 10 nM 1,25(OH)2D3 or 10 nM cholecalciferol (chol). mRNA was analyzed by Northern blots for MN1 and β- actin. B, MG-63 cells were treated with ethanol vehicle control (-) or 0.1 to 100 nM 1,25(OH)2D3 for 6 hours. mRNA was analyzed by Northern blots for MN1 and
β-actin. Fold increases in MN1 mRNA levels are expressed relative to β-actin
mRNA levels.
84 A 10 nM 10 nM B 1,25(OH)2D3 chol time (hr): 0 3 6 12 24 24 nM 1,25(OH)2D3: - 0.1 1 10 100
MN1 MN1
β-actin β-actin
fold fold increase: 1.0 1.8 2.0 4.8 4.2 1.2 increase: 1.0 1.9 2.7 4.1 4.4
85 Figure II-2
1,25(OH)2D3 induction of MN1 requires de novo RNA synthesis but not protein synthesis.
A, MG-63 cells were pre-treated with methanol vehicle control (- ACTD) or 1 ug/ml actinomycin D (+ ACTD) for 1 hour. Cells were then treated with ethanol control (Et) or 10 nM 1,25(OH)2D3 (1,25) for 6 hours. mRNA was analyzed by
Northern blots for MN1 and β-actin. B, MG-63 cells were pre-treated with ethanol control (-CHX) or 10 ug/ml cycloheximide (+ CHX) for 1 hour. Cells were then treated with ethanol control (Et) or 10 nM 1,25(OH)2D3 (1,25) for 6 hours. mRNA was analyzed by Northern blots for MN1 and β-actin. Fold increases in MN1 mRNA levels are expressed relative to β-actin mRNA levels. A “–“ indicates that
MN1 mRNA levels are undetectable in actinomycin D-treated samples.
86 AB -ACTD + ACTD - CHX + CHX
Et1,25 Et 1,25 Et1,25 Et 1,25
MN1 MN1
β-actin β-actin fold fold increase: 1.0 5.0 - - increase: 1.0 3.6 2.6 5.7
87 Figure II-3
MN1 augments 1,25(OH)2D3-dependent transcription through the VDR
ligand-binding domain.
A, COS-7 cells were transiently transfected with (VDRE)4-TATA-luc, SG5-VDR,
CMV-renilla luciferase, and either CMVTAG2B (vector) or CMVTAG2B-MN1.
Cells were treated with either ethanol vehicle control or 10 nM 1,25(OH)2D3 for
24 hours. B, COS-7 cells with transiently transfected with (VDRE)4-TATA-luc,
CMV-renilla, SG5 (vector) or SG5-VDR, and CMVTAG2B (vector) or
CMVTAG2B-MN1. Cells were treated with either ethanol vehicle control or 100 nm 1,25(OH)2D3 for 24 hours. C, COS-7 cells were transiently transfected with
(GAL4)5-TK-luc, CMV-renilla, GAL4 or GAL4-VDR LBD, and CMVTAG2B
(vector) or CMVTAG2B-MN1. Cells were treated with either ethanol vehicle
control or indicated doses of 1,25(OH)2D3 for 24 hours.
88 A 1.2 vehicle 1.0 1,25(OH)2D3 0.8
0.6
0.4
0.2
Normalized Activity Luciferase 0.0 vector MN1 B 12 vehicle 10 1,25(OH)2D3 8
6
4
2 Normalized Luciferase Activity 0 vector MN1 vector MN1 vector VDR
C 5.0 4.5 vehicle 4.0 10 nM 1,25(OH)2D3 3.5 100 nM 1,25(OH)2D3 3.0 2.5 2.0 1.5 1.0 Normalized Luciferase Activity 0.5 0.0 vector MN1 vector MN1 GAL4 GAL4-VDR LBD
89 Figure II-4
Effect of MN1 on other nuclear receptors.
A, COS-7 cells were transiently transfected with (GAL4)5-TK-luc, CMV-renilla,
GAL4 fusions of the ligand binding domain of the indicated nuclear receptor, and either CMVTAG2B (vector) or CMVTAG2B-MN1. Cells were treated with either ethanol vehicle control or 10 nM 1,25(OH)2D3 (VDR), 10 nM thyroid hormone
(TRβ), or 100 nM all-trans-retinoic acid (RARα). B, COS-7 cells were transiently
transfected with (GAL4)5-TK-luc, CMV-renilla, GAL4 fusions of the ligand-binding
domain of the indicated receptor, and either CMVTAG2B (vector) or
CMVTAG2B-MN1. C, COS-7 cells were transiently transfected (VDRE)4-TK-luc,
(GRE)2-TK-luc, or (RARE)4-TK-luc, CMV-renilla, SG5-VDR, -GR, or -RARα, and
either CMVTAG2B (vector) or CMVTAG2B-MN1. Cells were treated with either
ethanol vehicle control or 10 nM 1,25(OH)2D3 (VDR), 1 μM dexamethasone (GR),
or 100 nM all-trans-retinoic acid (RARα) for 24 hours.
90 A 60
50
40
30
20
10 Normalized luciferase activity
0 ligand: -+-+-+-+-+-+ vector MN1 vector MN1 vector MN1 VDR TRβ RARα
B 10 9 vector 8 MN1 7 6 5 4 3 2 Normalized luciferase activity 1 0 RORα RORβ RORγ
C 50 45 vehicle 40 ligand 35 30 25 20 15 10 Normalized luciferase activity 5 0 vector MN1 vector MN1 vector MN1 VDRE GRE RARE
91 Figure II-5
MN1 synergizes with SRC coactivators.
A and B, COS-7 cells with transiently transfected with (VDRE)4-TATA-luc, CMV- renilla, SG5-VDR, and either vector control or the indicated coactivators. Cells were treated with either ethanol vehicle control or 100 nM 1,25(OH)2D3 for 24
hours.
92 A
70 vehicle 60 100 nM 1,25(OH)2D3 50
40
30
20 Normalized luciferase activity 10
0 vector MN1 SRC-1 MN1 + SRC-1
B
45 vehicle 40 100 nM 1,25(OH) D 35 2 3
30
25
20
15
10 Normalized luciferase activity 5
0 vector MN1 SRC-2 MN1 + SRC-2
93 Figure II-6
MN1 inhibits proliferation in osteoblastic cells.
A, MG-63 cells were cultured in the presence of either ethanol vehicle control or
10 nM 1,25(OH)2D3 and counted at the indicated times. B, MG-63 cells were
transfected with either CMVTAG2B (vector), CMVTAG2B-MN1, or CMVTAG2A-
RORβ. Stable transfectants were selected by culture in the presence of G418 for
7 days. Cells were visualized by staining with crystal violet. C, Total cell number was quantified by solubilizing the crystal violet dye in 0.1 M sodium citrate in 50% ethanol, and absorbance was measured at 540 nm.
94 A 40
35 vehicle ) 4 30 1,25 25 20 15
cell count (X10 cell count 10 5 0 0 24487296 time in culture (hr) B
vector
MN1
RORβ
C 1 0.9 0.8 0.7 0.6 0.5 0.4 0.3 Average Absorbance 0.2 0.1 0 vector MN1 RORβ
95 Figure II-7
MN1 decreases S-phase entry in osteoblastic cells.
A, MG-63 cells were transiently transfected with either pCMS-EGFP, pCMS-
EGFP-MN1, or pCMS-EGFP-SRC-1 and allowed to express for 24 hours. Cells were pulse labeled with BrdU for one hour, fixed, and immunostained to visualize
BrdU incorporation. Nuclei were counterstained with DAPI. Representative fields are shown. EGFP-positive BrdU-positive cells are indicated by open arrows, and EGFP-positive BrdU-negative cells are indicated by closed arrows.
B, Quantification of percentage of EGFP-expressing cells that were BrdU- positive.
96 A EGFP BrdU DAPI
vector
MN1
SRC-1
B 45 40 35 30 25 20 15
% BrdU-positive cells 10 5 0 Vector MN1 SRC-1
97 CHAPTER III
THE 1,25(OH)2D3-INDUCED TRANSCRIPTION FACTOR CCAAT/ENHANCER- BINDING PROTEIN-β COOPERATES WITH VDR TO REGULATE 24- HYDROXYLASE EXPRESSION AND MAY BE REQUIRED FOR OSTEOBLAST DIFFERENTIATION AND BONE MINERALIZATION∗
INTRODUCTION
Vitamin D is essential for mineral ion homeostasis and for proper bone
development and maintenance. The most bioactive form of vitamin D, 1,25- dihydroxyvitamin D3 (1,25(OH)2D3), is generated by two sequential
hydroxylations of vitamin D3, or cholicalciferol. Vitamin D3 is first hydroxylated by
25-hydroxylase in the liver and then by 1α-hydroxylase in the kidney and other
tissues. 1,25(OH)2D3 functions through the vitamin D receptor (VDR), a member
of the nuclear hormone receptor family. Upon ligand binding, VDR
heterodimerizes with the retinoid X receptor (RXR) and interacts vitamin D
response elements (VDREs), specific DNA motifs in the promoters of
1,25(OH)2D3-regulated genes. The active heterodimer recruits the basal
transcriptional machinery to these promoters to stimulate transcriptional
activation of target genes. Accessory factors, called coactivators, facilitate
transcriptional activation mediated by VDR and other nuclear receptors.
Numerous VDR coactivators have been described, including the steroid receptor
coactivator (SRC) proteins, the DRIP (VDR-interacting proteins) complex, and
∗ Adapted, in part, from Dhawan P, Peng X, Sutton AL, MacDonald PN, Croniger CM, Trautwein C, Centrella M, McCarthy TL, Christakos S 2005 Functional cooperation between CCAAT/enhancer-binding proteins and the vitamin D receptor in regulation of 25-hydroxyvitamin D3 24-hydroxylase. Mol Cell Biol 25:472-487.
98 NCoA-62/ski-interacting protein (SKIP) (255, 258). These coactivators promote
transcription of target genes by modifying the local chromatin environment (61),
recruiting the RNA polymerase II holoenzyme (72), and through coupling
transcriptional activation to mRNA splicing (78), among other mechanisms. The
ever-expanding number of protein factors that are involved in VDR-mediated
transcription highlights the molecular complexities in regulated target gene
expression.
One of the most potently induced 1,25(OH)2D3 target gene encodes the catabolic
enzyme 24-hydroxylase (24(OH)ase) (291). 24(OH)ase catalyzes the
hydroxylation of both 1,25(OH)2D3 and its precursor, 25(OH)D3, to yield
metabolites that are further degraded and eventually excreted (292). Thus,
1,25(OH)2D3 operates in a negative feed-back loop to prevent excess synthesis
of this hormone. 1,25(OH)2D3 increases 24(OH)ase expression by binding to two
tandemly arranged vitamin D response elements (VDREs) in the 24(OH)ase
promoter (293). Other proteins, such as Ets-1, also contribute to modulating
24(OH)ase promoter activity by binding to adjacent DNA elements and
cooperating with VDR to stimulate transcription (294, 295).
In the current study, we characterize the function of CCAAT/enhancer-binding
protein-β (C/EBPβ) in regulating 1,25(OH)2D3-mediated 24(OH)ase expression
and in osteoblast mineralization in vitro and in vivo. C/EBPβ is a member of the
C/EBP family of basic-leucine zipper (bZIP) transcription factors (296). C/EBPs
99 interact with CCAAT box motifs present in several gene promoters through the
basic amino acid-rich DNA-binding domain. The “leucine zipper” module
mediates homo- and heterodimerization among C/EBP family members.
Additionally, all but one (C/EBPγ) of these transcription factors contain an N-
terminal activation domain that associates with the basal transcriptional
machinery. Alternative transcriptional initiation (297) or proteolytic processing
(298, 299) can generate three protein isoforms of C/EBPβ: liver activating
protein* (LAP*; 38 kD), liver activating protein (LAP; 35 kD), and liver inhibitory
protein (LIP; 20 kD). LAP* and LAP contain both the transcriptional activation
domain and the DNA-binding bZIP domain and, therefore, are fully functional
transcription factors. LIP contains only the bZIP domain and often acts as a
dominant-negative inhibitor of C/EBPs by forming inactive heterodimers with
other family members.
Mice with a targeted deletion in C/EBPβ have provided insight into the in vivo
functions of this transcription factor (300, 301). These murine models indicate that C/EBPβ is essential for normal macrophage differentiation (300, 301), liver regeneration (302), mammary gland development (303, 304), glucose and lipid metabolism (305, 306), and female fertility (307). In the bone, C/EBPβ knockout
mice display disorganized growth plates and increased chondrocyte apoptosis
(308), suggesting an important role for C/EBPβ in skeletal development and
homeostasis. However, it is unknown if the osteoblast lineage is affected in
these mice.
100
In the current study, we characterized C/EBPβ as a 1,25(OH)2D3-regulated gene in primary osteoblasts that was critical for maximal VDR-mediated induction of
24(OH)ase expression. Additionally, C/EBPβ-deficient osteoblasts failed to mineralize in vitro, uncovering a novel function for this transcription factor in modulating osteoblast differentiation. Preliminary studies indicated that although neither adult C/EBPβ knockout nor VDRKO knockout mice had obvious bone defects, a double C/EBPβ/VDR knockout mouse had dramatically reduced trabecular and cortical bone thickness and abnormal bone structure. Together, these data suggest that C/EBPβ cooperates with VDR to regulate 24(OH)ase expression and potentially bone mineralization in vivo, and that C/EBPβ is essential for osteoblast mineralization in vitro.
MATERIALS AND METHODS
Animals. All experiments involving mice were approved by the Institutional
Animal Care and Use Committee (IACUC) at Case Western Reserve University.
Mice with a targeted deletion in the C/EBPβ locus have been described previously (300). Fetal mice used to generate C/EBPβ WT and KO primary osteoblasts were a mixture of 129/Sv/Ev and C57Bl/6. Adult VDR/CEBPβ double knockout (VDRKO/CEBPβKO) mice were generated by crossing VDR heterozygous mice (kindly provided by Dr. Marie Demay, Endocrine Research
Unit, Massachusetts General Hospital; (106)) to CEBPβ heterozygous mice.
Both single knockout strains of mice were previously backcrossed into C57Bl/6
101 for more than 10 generations. All adult mice were fed a high-calcium high-
lactose calcium diet to maintain normocalcemia in VDRKO mice (223).
Primary osteoblast isolation and culture. 18.5 dpc mice were killed by
decapitation. Animals were genotyped by PCR. Individual calvaria were
dissected and the adherent tissue was removed. Primary osteoblasts were
liberated from the bone by serial collagenase digestions (309). Cultures were
established in α-MEM (Invitrogen, Calsbad, CA) supplemented with 15% fetal
bovine serum and combined based on genotype.
Nuclear extracts and western blot analysis (performed by Dr. Puneet Dhawan).
Primary osteoblast cells were lysed in hypotonic buffer containing 10 mM HEPES
(pH 7.4), 1.5 mM MgCl2, 10 mM KCl, 0.5 mM dithiothreitol, phosphatase
inhibitors (1 mM sodium orthovanadate, 10 mM sodium fluoride), protease
inhibitors (0.5 mM phenylmethylsulfonyl fluoride, 1 mg of pepstatin A per ml, 2 mg
of leupeptin per ml, 2 mg of aprotinin per ml), and 1% Triton X-100. Nuclei were pelleted at 3,500 x g for 5 min, and cytoplasmic supernatants were separated.
Nuclei were resuspended in hypertonic buffer containing 0.42 M NaCl, 0.2 mM
EDTA, 25% glycerol, and the phosphatase and protease inhibitors indicated above. Soluble nuclear proteins were released by 60 min of incubation at 4°C,
and insoluble material was separated by centrifugation at 12,000 x g for 5 min.
Fifty micrograms of protein was separated by SDS-polyacrylamide gel electrophoresis (PAGE) and analyzed by Western blot analysis using standard
102 methods with an anti-C/EBPβ antibody (Santa Cruz Biotechnology, Santa Cruz,
California).
Transient transfections and CAT assays. Cells were grown in α-MEM with 15% fetal bovine serum and transfected by Lipofectamine PLUS reagent (Invitrogen) with a rat24OHase promoter-CAT reporter plasmid containing the –1367 to +74
(–1367/+74) promoter fragment, which encompasses both VDREs at –258/–244
and –151/–137 (293). Cells were treated for 48 hours with the indicated dosages
of 1,25(OH)2D3. CAT activity was determined essentially as described (310) and
normalized to the protein concentration of the cell extract.
RNA extraction and RT-PCR analysis. Total RNA was extracted from primary
osteoblasts after 24 hours of ligand stimulation using RNA-Bee (Tel-Test, Inc.,
Friendswood, TX) according the manufacturer’s instructions. RNA was treated
with DNase I to remove contaminating genomic DNA (Promega, Madison, WI). 1 ug of total RNA was subjected to RT-PCR (performed by Dr. Puneet Dhawan) using Superscript one-step RT-PCR system with Platinum Taq DNA polymerase
(Invitrogen). Primers used were 24(OH)ase (5'-GCCGAGCCTGCTGGAA3' and
5'-CCCCATAAAATCAGCCAAGAC-3') and β-actin (5'-
CCTGTGGATCTGACAGCTGAA-3' and 5'-TCCCAAATCGGTTGGAGATA-3').
PCR cycle numbers used were as follows: with 24(OH)ase, 32 cycles, and with
actin, 30 cycles. The cycles were chosen so that the amplification was conducted
in the linear range of amplification efficiency. The resulting PCR products were
103 subjected to electrophoresis on a 1% agarose gel containing ethidium bromide,
and bands were visualized under UV light. Gel data were recorded using the
Gene Genius bioImaging system (Syngene, Frederick, Md.), and relative
densities of the bands were determined using Gene Tool software (Syngene).
Data were normalized for the expression of β-actin within the sample.
Osteoblast differentiation and Alizarin Red S staining. Cells were seeded
at a density of 50,000 cells per well of a six-well dish and grown in α-MEM
supplemented with 10% fetal bovine serum, 50 μg/ml L-ascorbic acid, and 10 mM β-glycerophosphate. Media was replenished twice per week during the 31 day differentiation period. Cells were fixed in 10% phosphate-buffered formalin for one hour. Mineralized nodules were visualized by staining with 2% alizarin red S in 1% ethanol for 10 minutes and destaining in distilled water.
Ex-vivo μCT imaging. Adult mice between 5-9 weeks of age were killed by
asphyxiation with CO2. Tibiae were dissected, fixed in 10% neutral-buffered
formalin overnight at room temperature, and then transferred 70% ethanol for storage at 4° C. Micro-computerized tomography (μCT) was performed by the
laboratory of Dr. Kimerly Powell from the Department of Biomedical Engineering
at the Lerner Research Institute of the Cleveland Clinic Foundation using a custom designed imaging system. Images with a spatial resolution of 18 μm were obtained by collecting one-hundred and eighty 512 X 512 12-bit projection
104 radiographs at 1° intervals around one-half of the tibiae. Final images were
processed and reconstructed using custom in-house software.
RESULTS
1,25(OH)2D3 increases C/EBPβ expression in osteoblasts. C/EBPβ was initially identified as a 1,25(OH)2D3-induced gene by microarray analysis. C/EBPβ mRNA was increased approximately 4-fold in kidneys from mice treated with
1,25(OH)2D3 (data not shown). Since osteoblasts are also important targets of
1,25(OH)2D3, we determined if C/EBPβ was similarly regulated by 1,25(OH)2D3 in
primary murine osteoblasts. Northern blot and western blot analyses showed
that 1,25(OH)2D3 also induced C/EBPβ mRNA (data not shown) and protein levels in primary osteoblasts (Fig. 1).
C/EBPβ is critical for maximal 1,25(OH)2D3-mediated induction of 24(OH)ase
expression. Sequence analysis of the 5’ regulatory region of r24(OH)ase, a classical 1,25(OH)2D3 target gene, revealed two putative C/EBPβ-binding
elements (Fig. 2A). Subsequent studies showed that C/EBPβ enhanced the
induction of the r24(OH)ase promoter by 1,25(OH)2D3, and that both sites were
required for this enhancement in COS-7 cells (data not shown). To determine if
C/EBPβ is essential for 1,25(OH)2D3-mediated increases in 24(OH)ase
expression, we utilized osteoblasts derived from C/EBPβ-null fetal mice and their wild-type littermates. The r24(OH)ase promoter was transfected into these osteoblasts and stimulated with 1,25(OH)2D3. As shown in Fig. 2B, 1,25(OH)2D3-
105 induced promoter activity was reduced by nearly 10-fold in C/EBPβ-deficient osteoblasts as compared to wild-type osteoblasts. Likewise, stimulation of endogenous 24(OH)ase mRNA levels by 1,25(OH)2D3 was impaired in the absence of C/EBPβ (Fig. 2C). Together, these data show that C/EBPβ is essential for maximal induction of 24(OH)ase expression in osteoblasts.
C/EBPβ is required for osteoblast mineralization. Given that C/EBPβ regulates cellular differentiation in a variety of other tissues (296), we tested whether it was involved in osteoblast differentiation. Under the appropriate culture conditions, primary osteoblasts will differentiate and mineralize the extracellular matix, forming bone nodules visible following alizarin red S staining.
Whereas wild-type osteoblasts formed numerous large bone nodules, C/EBPβ knockout cells formed very few nodules (Fig. 3A). Furthermore, C/EBPβ- deficient cells were larger and more cuboidal as compared to the normal fibroblast-like appearance of wild-type osteoblasts (Fig. 3B), suggesting an alteration in the differentiation pathway of cells lacking C/EBPβ. These data indicate that C/EBPβ is crucial for normal osteoblast differentiation and mineralization in vitro.
Targeted deletion of both C/EBPβ and VDR results in severe bone abnormalities.
Since C/EBPβ-deficient osteoblasts cannot appropriately mineralize in vitro,
C/EBPβ knockout mice were analyzed for bone mineralization defects. However, preliminary μCT imaging indicated that the C/EBPβ knockout tibia had no
106 obvious abnormalities (Fig. 4). Given that C/EBPβ is induced by 1,25(OH)2D3 and cooperates with VDR to regulate gene expression, we determined if these two proteins cooperate in regulating bone mineralization in vivo. Consistent with previous studies (223), the single VDR knockout bone had no apparent defects
(Fig. 4). Although each single knockout bone did not display abnormal mineralization, the double C/EBPβ/VDR-deficient tibia had very thin cortical and trabecular bone (Fig. 4). In fact, the bone was so fragile that the growth plate virtually disintegrated during dissection. Further, the metaphysis of the tibia was widened and disorganized. Thus, while deletion of either C/EBPβ or VDR alone does not cause obvious bone defects, disruption of both genes causes a drastic mineralization defect that affects both cortical and trabecular bone. These observations suggest that C/EBPβ and VDR compensate for one another in regulating bone mineralization and may signal through convergent pathways during bone development or maintenance.
DISCUSSION
An important target of 1,25(OH)2D3 is 24(OH)ase, which operates in a negative
feedback loop to negatively regulate synthesis of this hormone (292). Although
VDR directly interacts with two response elements in the 24(OH)ase promoter
(293), the present study indicates that additional mechanisms are involved in
1,25(OH)2D3-mediated induction of 24(OH)ase. Our data show that C/EBPβ is a novel target of 1,25(OH)2D3 in osteoblasts and targeted deletion of this
transcription factor impaired 1,25(OH)2D3-mediated induction of 24(OH)ase.
107 Thus, 1,25(OH)2D3 regulates 24(OH)ase both directly by binding to VDREs in the
promoter and indirectly by stimulating expression of C/EBPβ, which binds to and
activates distinct response elements in the promoter. Furthermore, we demonstrated that C/EBPβ was critical for osteoblast differentiation and mineralization in vitro since osteoblasts derived C/EBPβ-deficient mice failed to
form bone nodules in culture. Finally, μCT imaging indicated that while disruption
of either C/EBPβ or VDR alone did not cause bone abnormalities, deletion of
both genes caused profound mineralization defects in both cortical and trabecular bone. These data suggest that C/EBPβ and VDR not only
cooperatively control gene expression, but may also cooperate to regulate bone
mineralization. This study uncovers a novel interaction between VDR and
C/EBPβ signaling pathways in regulating gene expression and bone
mineralization.
Studies utilizing C/EBPβ-null osteoblasts indicated that C/EBPβ is essential for
maximal 1,25(OH)2D3-mediated stimulation of 24(OH)ase transcription (Fig. 2).
Previous reports have indicated that C/EBPs regulate numerous other genes in
osteoblasts. For example, C/EBPδ, in conjunction with prostaglandin signaling,
promotes transcription of insulin-like growth factor (311, 312). C/EBPβ
synergizes with runx2, the master osteoblast transcription factor, to enhance
expression of the bone-specific matrix protein osteocalcin (313). Thus, evidence
is emerging for an important role for C/EBPβ in regulating osteoblastic gene
expression. Extending the use of the C/EBPβ-null osteoblasts to studying other
108 genes, such as insulin-like growth factor and osteocalcin, will further confirm the physiological relevance of C/EBPβ in their regulation.
In the present study, microarray analysis identified C/EBPβ as a 1,25(OH)2D3 target gene in the kidney, and subsequent experiments showed that 1,25(OH)2D3 also regulates C/EPBβ expression in murine osteoblasts (Fig. 1). Consistent with these findings, previous studies have shown that 1,25(OH)2D3 induces C/EBPβ
expression in rat osteoblasts (313). A recent report indicates that 1,25(OH)2D3 also induces C/EBPβ expression in myeloid cells (314), suggesting that this regulation is not tissue-specific. Either siRNA-mediated silencing of C/EBPβ or transfection of decoy C/EBPβ binding sequence oligonucleotides diminished
1,25(OH)2D3-induced differentiation of myeloid cells (314), indicating that this
transcription factor is an important mediator of the 1,25(OH)2D3 differentiative
signal. Likewise, the current study suggests that C/EBPβ is important in
osteoblastic cell differentiation (Fig. 3). However, the role of C/EBPβ in
osteoblast differentiation and bone mineralization remains a subject of some
controversy. Overexpression of LAP, the intermediate-length form of C/EBPβ
that still includes the transactivation domain, in the murine MC3T3 osteoblastic
cell line inhibits osteogenic differentiation and mineralization (315). Likewise,
overexpression of a dominant negative inhibitor of C/EBPβ enhances osteoblast
differentiation in the multipotent ST-2 cell line (316). However, it is important to
note that the dominant negative inhibitor used this study interferes with the
function of all C/EBP family members, indicating that one or more of the C/EBPs
109 could be inhibiting osteoblast differentiation in this model. A third report indicates that overexpression of LIP, which is thought to act as a dominant negative inhibitor of full-length C/EBPβ, in C3H10T1/2 multipotent mesenchymal cells induces osteogenic differentiation (317). Curiously, the same study shows that overexpression full-length C/EBPβ also promotes osteoblastic differentiation in both C3H10T1/2 multipotent mesenchymal cells and in primary osteoblasts.
Given such conflicting data, it is not clear from these three in vitro studies whether C/EBPβ plays a positive or negative role in osteogenic differentiation and mineralization. However, a recent study reported a transgenic mouse that overexpresses LIP in osteoblasts (318). Bones from these transgenic animals are osteopenic, and derived osteoblastic cells are impaired in their differentiative capacity. These findings corroborate our data since genetic deletion of C/EBPβ has similar effects in cultured osteoblasts (Fig. 3). The disparities among these studies may be explained by differences in cell culture lines or levels of forced expression. Thus, analysis of in vivo models in which C/EBPβ is disrupted may prove more useful in uncovering the true function of C/EBPβ in osteoblasts and in bone.
Our findings that dramatic bone defects are present in a VDRKO/C/EBPβKO mouse but absent in either of the single knockout animals (Fig. 4) suggests that a compensatory relationship exists between 1,25(OH)2D3 and C/EBPβ. Given that these two proteins cooperate to stimulate 24(OH)ase transcription (Fig. 2), it is possible that they modulate expression of other genes that are important in
110 maintaining bone mineral density. Identification of such genes will provide even
further insight into how these two transcription factors functionally interact in the
skeleton and potentially other tissues.
In summary, the current study has demonstrated that C/EBPβ is a 1,25(OH)2D3 target gene that cooperates with VDR to stimulate expression of 24(OH)ase.
Furthermore, C/EBPβ-null osteoblasts were unable to appropriately differentiate and mineralize the extracellular matrix in vitro. Analysis of VDRKO, C/EBPβKO, and double KO bones showed that although neither single knockout strain exhibited notable deformities, a double C/EBPβ/VDR knockout mouse had markedly reduced and abnormal trabecular and cortical bone. These findings suggest a functional interplay between VDR and C/EBPβ in regulating target gene expression and in promoting appropriate bone mineralization.
111 Figure III-1
1,25(OH)2D3 increases C/EBPβ expression in primary osteoblasts.
Nuclear extracts from primary murine osteoblast cells treated with either vehicle
–8 (Et) or with 10 M 1,25(OH)2D3 (+D) for 24 or 48 hours were subjected to western blot analysis using an anti-C/EBPβ antibody.
112 D
Et 24 hr 48 hr
C/EBPβ
113 Figure III-2
C/EBPβ is required for maximal 1,25(OH)2D3-mediated induction of
24OHase epression.
A, Schematic of r24(OH)ase promoter showing both the VDREs and the potential
C/EBPβ binding sites (C/EBPβ BE). B, 1,25(OH)2D3-mediated stimulation of
r24(OH)ase promoter is impaired in C/EBPβ-deficient cells. Primary osteoblastic
cells from C/EBPβ-/- mice and control littermates (WT) were transfected with the
r24(OH)ase promoter and treated with either vehicle control (basal) or the
indicated doses of 1,25(OH)2D3. CAT assays were performed using cell lysates.
C, Diminished 1,25(OH)2D3-mediated induction of endogenous 24(OH)ase
mRNA in C/EBPβ-/- osteoblasts. Primary osteoblastic cells from C/EBPβKO mice and control littermates (WT) were treated with either vehicle control (-D/basal) or
-8 10 M 1,25(OH)2D3. RNA was extracted and subjected to RT-PCR for either
24(OH)ase or β-actin.
114 A
C/EBPβ BE C/EBPβ BE VDRE VDRE CAT -964/-955 -395/-388 -258/-244 -151/-137
B
C
115 Figure III-3
Targeted deletion of C/EBP-β disrupts osteoblast mineralization in vitro.
A, C/EBPβKO osteoblasts fail to mineralize in vitro. Primary osteoblasts were isolated from C/EBPβKO or WT littermate fetal mice and stimulated to differentiate by supplementing the culture medium with 50 ug/ml ascorbic acid and 10 mM β-glycerophosphate for 31 days. Mineralized nodules were visualized with alizarin red S staining. Shown are duplicate wells from a representative experiment that was repeated with similar results. B, C/EBPβKO osteoblasts have an altered cellular morphology. Primary osteoblasts were isolated and cultured as in A. Photomicrographs of cells are shown from a representative experiment that was repeated with similar results.
116 A
WT C/EBPβ KO B
WT C/EBPβ KO
117 Figure III-4
Targeted deletion of both C/EBPβ and VDR causes undermineralized bones
in vivo.
Tibiae of 5-9 week-old mice were dissected, fixed, and analyzed by μCT imaging.
Shown are longitudinal sections. Note that the double knockout bone was so fragile that the growth plate fractured during dissection.
118 VDR WT VDR WT VDR KO VDR KO CEBPβ WT CEBPβ KO CEBPβ WT CEBPβ KO
119 CHAPTER IV
TARGETED OVEREXPRESSION OF THE 1,25(OH)2D3-REGULATED GENE SEMAPHORIN3B IN OSTEOBLASTS CAUSES INCREASED BONE RESORPTION AND OSTEOPENIA∗
INTRODUCTION
1,25(OH)2D3 is the bioactive, or hormonal, form of vitamin D. This hormone
functions through the vitamin D receptor (VDR) to regulate the transcription of
target genes in a number of tissues (255, 257). The 1,25(OH)2D3/VDR endocrine
system functions in diverse biological processes, such as hair follicle cycling,
mammary gland development, and immune cell function (255). However, the
most profound action of 1,25(OH)2D3 is to protect skeletal integrity since
deficiencies in either the hormone or the receptor result in undermineralized
bones (104, 254).
Acting in concert with parathyroid hormone, 1,25(OH)2D3 preserves bone
mineralization primarily by maintaining calcium and phosphate homeostasis.
1,25(OH)2D3 controls serum levels of these minerals by stimulating calcium and
phosphate uptake by the intestine, by increasing reabsorption of calcium and phosphate in the kidney, and by liberating calcium and phosphate from skeletal stores (254). When dietary sources of calcium are inadequate, 1,25(OH)2D3 promotes osteoclastogenesis and bone resorption, in part, by stimulating osteoblasts to synthesis and secrete receptor activator of NF-κB ligand (RANKL)
∗ Adapted, in part, from Sutton AL, Kharode YP, Komm BS, MacDonald PN 2005 Targeted overexpression of the 1,25(OH)2D3-regulated gene Semaphorin3B causes increased bone resorption and osteopenia, manuscript in preparation.
120 (243), a molecule essential for osteoclast formation and function (191, 193).
Under conditions of normocalcemia, the 1,25(OH)2D3/VDR endocrine system
also modulates osteoblast differentiation and mineralization (232, 251, 252, 256).
Thus, 1,25(OH)2D3 functions both systemically to regulate serum concentrations
of calcium and phosphate and locally to fine-tune of the balance between bone
formation and bone resorption. However, with the exception of RANKL and bone
matrix proteins (236, 243, 319, 320), the target genes that mediate these effects
of 1,25(OH)2D3 in osteoblasts remain largely unknown.
In this study, we characterize the regulation and the function of semaphorin3B
(SEMA3B), a novel 1,25(OH)2D3 target gene in osteoblasts. Semaphorins are a
diverse group of signaling molecules that were originally identified as axonal guidance proteins, but subsequently have been found to regulate cell migration, cell growth, differentiation, and angiogenesis in a variety of tissues (321).
SEMA3 molecules are secreted proteins that signal through neuropilin receptors
(322, 323) and plexin co-receptors (324, 325). The SEMA3B gene was first identified based on its position in the chromosomal region 3p21.3, a frequent site of loss-of-heterozygosity in lung, kidney, ovarian and testicular cancers (326).
Re-expression of SEMA3B in either lung or ovarian cancer cells diminishes their
proliferative and tumorigenic potential (327, 328), indicating that SEMA3B is probably a tumor suppressor. SEMA3B expression has been detected in odentoblasts, the dental counterpart to skeletal osteoblasts (329) and in osteoblasts in vitro (330). Furthermore, neuropilin-1 expression has been
121 detected in osteoclasts and in osteoblasts in vitro and in vivo and appears to be
down-regulated as osteoblasts differentiate into more mature osteocytes (331).
Targeted deletion of a related class 3 semaphorin, sema3A, causes patterning
defects leading to rib duplications (332). Beyond these initial reports, very little is
known regarding the regulation of SEMA3B and the function of this signaling
system in osteoblasts and osteoclasts.
The present study characterizes SEMA3B as a novel 1,25(OH)2D3-regulated
gene in osteoblastic cells. SEMA3B expression was detected in cells of the
osteoblast lineage as well as a select population of chondrocytes in vivo. To
probe the function of SEMA3B in bone, transgenic mice were created that
express SEMA3B under the control of the osteoblast-selective 2.3 kb promoter of
the mouse pro-α 1 (I) collagen gene. Male transgenic mice that highly express
the transgene had lower body weights, shorter tibiae, and displayed a profound
defect in trabecular and cortical bone mineralization at four weeks of age. While
osteoblast function appeared normal SEMA3B transgenic mice in vivo, osteoclast
number and bone resorption were increased. In vitro studies indicated that
transgenic osteoblasts actually differentiated and mineralized at an accelerated rate, but supported increased osteoclastogeneis. Thus, this study identifies
SEMA3B as a novel regulator of bone mass that may function by stimulating
osteoclastogenesis and osteoclast activity.
122 MATERIALS AND METHODS
Cell Culture. MG-63 human osteoblastic cells were maintained in modified
essential media (MEM) supplemented with 10% fetal bovine serum (FBS). For
experiments using 1,25(OH)2D3 compounds, MG-63 cells were grown in media
supplemented with charcoal-stripped calf serum for three days prior to the experiment.
RNA extraction and Northern blot analysis. mRNA was isolated from MG-63
cells with the FastTrack system (Invitrogen, Carlsbad, CA) according to the
manufacturer’s instructions. For bone RNA isolation, humeri were ground to a
fine powder using a mortar and pestle over liquid nitrogen. Total RNA was
extracted from the powder using Trizol (Invitrogen) according to the manufacturer’s instructions. RNA was separated on a formaldehyde/agarose gel and transferred to a Duralon membrane (Stratagene, La Jolla, CA) by capillary action. [α32P]-labeled probes were synthesized using the Prime-A-Gene kit
(Promega, Madison, WI) according to the manufacturer’s instructions and
hybridized to the blots using standard methods.
Antibodies and immunohistochemistry. Affinity purified rabbit polyclonal anti-
SEMA3B antibodies were generated against a synthetic peptide corresponding to
the extreme C-terminus (327) (Bio-synthesis Incorporated, Lewisville, TX), which
is 100% conserved between the human and mouse proteins. Tibiae from 8-week
male C57BL/6J mice were fixed in 10% neutral buffered formalin overnight at
123 room temperature, decalcified in formic acid for 4 days at room temperature,
embedded in paraffin, and sectioned. Following blocking in goat serum, sections were incubated with a 1:200 dilution of the anti-SEMA3B antibody. The rabbit
IgG Vectastain ABC kit and the DAB peroxidase substrate kit (Vector
Laboratories, Burlingame, CA) were used for detection of the primary antibody
according to the manufacturer’s recommendations.
Animals and transgenic mouse construction. All animal studies were approved by the in the Institutional Animal Care and Use Committee at Case Western
Reserve University (CWRU). Mice were housed in pathogen-free microisolator
units with a 12-hour light, 12-hour dark cycle and given food and water ad libitum.
The osteoblast-targeted transgene was constructed by replacing the lacZ cassette in the 2300lacZ plasmid (333) with the human SEMA3B coding sequence from pcDNA3-SEMA3B (326) to drive expression of SEMA3B from the
2.3-kb osteoblast-specific promoter region of the mouse pro-α 1 (I) collagen
gene. The CWRU Transgenic Core Facility created transgenic mice by standard methods. Transgenic mice were identified by PCR amplification of tail genomic
DNA using transgene-specific primers. All transgenic animals and non- transgenic littermates used for analyses were greater than 95% C57BL/6J. All adult mice were killed by asphyxiation with CO2.
Imaging analysis. Tibiae from non-transgenic and transgenic littermates were
dissected, fixed in formalin overnight, and transferred to 70% ethanol.
124 Volumetric bone mineral density (mg/cm3) measurements of the tibiae were performed using an XCT Research peripheral quantitative computer tomography
(pCQT) densitometer (Stratec Medizintechnik, Pforzheim, Germany) as previously described (162, 334). Trabecular bone volume fraction and microarchitecture were evaluated on the proximal tibia 12 mm distal to the end of growth plate using a μCT 40 (Scanco Medical, Southeastern, PA). One hundred
12 μm slices of each bone were analyzed.
Undemineralized histology and histomorphometry. Mice were injected intraperitoneally with calcein (10 mg/kg) at 5 days prior to killing and with tetracycline (25 mg/kg) at 1 day prior to killing at 31 days of age (335). Tibiae were dissected, fixed in formalin overnight under a vacuum, and sequentially dehydrated in 70% ethanol and 95% ethanol to preserve the fluorescent labels.
Longitudinal undemineralized 5 µm sections were cut from methyl methacrylate
(MMA) plastic embedded blocks of frontal sections of each tibia. Sections were stained with Goldner’s Trichrome stain for the static measurements, and unstained sections were used to visualize the fluorescent labels and perform the dynamic measurements. Standard bone histomorphometry was performed as described (336) using Bioquant Image Analysis software (R & M Biometrics,
Nashville, TN).
Primary osteoblast isolation and differentiation. 1-3 day old newborn mice were killed by decapitation. Animals were genotyped by PCR. Individual
125 calvaria were dissected and the adherent tissue was removed. Primary
osteoblasts were liberated from the bone by serial collagenase digestions (309).
Cultures were established in alpha-MEM (Invitrogen, Calsbad, CA)
supplemented with 15% fetal bovine serum and combined based on genotype.
For differentiation studies, cells were seeded at a density of 50,000 cells per well
of a six-well dish and grown in alpha-MEM supplemented with 10% fetal bovine
serum, 50 μg/ml L-ascorbic acid, and 10 mM β-glycerophosphate. Media was
replenished twice per week during the 25 day differentiation period. Alkaline
phosphatase (ALP) activity was determined by using a colorimetric kit that
measures the conversion of p-nitrophenyl phosphate to p-nitrophenol according
to the manufacturer’s instructions (Sigma Aldrich, St. Louis, MO). ALP activity
was normalized to protein concentration of the lysate as measured by the
bicinchoninic acid (BCA) assay (Pierce Biotechnology, Inc., Rockford, IL).
Mineralized nodules were visualized by staining fixed cells with 2% alizarin red S
in 1% ethanol for 10 minutes and destaining in distilled water.
Osteoclast differentiation and TRAP staining. Primary osteoblasts were
differentiated for 15 days as described above prior to overlaying osteoclast precursor cells. Osteoclast precursor cells were derived from the spleens of 7-10
week old female C57BL/6J mice. Cells were plated overnight in phenol red-free
α-MEM supplemented with 10% heat-inactivated FBS and 10 ng/ml macrophage
colony-stimulating factor (R & D Systems, Inc., Minneapolis, MN). Non-adherent
cells were added to the osteoblast cultures at a density of 500,000 cells per cm2
126 in phenol red-free α-MEM containing 10% heat-inactivated FBS, 100 nM
dexamethasone, and 10 nM 1,25(OH)2D3. The cocultures were grown for an
additional 8 days, and media was replenished twice. Cells were stained for
tartrate-resistant phosphatase (TRAP) expression using the leukocyte acid
phosphatase kit (Sigma Aldrich) according to the manufacturer’s instructions except that the tartrate concentration was increased to 50 mM as described
(337). TRAP-positive cells with > 3 nuclei were counted in triplicate wells.
RESULTS
SEMA3B is a 1,25(OH)2D3-regulated gene in osteoblastic cells. We utilized
microarray analysis as an initial screen to identify 1,25(OH)2D3-regulated genes
in MG-63 osteoblastic cells. Following a six-hour treatment with 10 nM
1,25(OH)2D3, the SEMA3B transcript was induced by approximately 10-fold (data
not shown). Northern blot analysis confirmed that 1,25(OH)2D3 increased
SEMA3B mRNA levels in a time- and dose-dependent manner (Fig. 1A and 1B).
This increase occurred as early as 3 hours, and mRNA levels continued to rise
up to 24 hours (Fig. 1A). Similarly, as little as 0.1 nM 1,25(OH)2D3 induced
SEMA3B, and transcript levels continued to increase up to 100 nM 1,25(OH)2D3
(Fig. 1B). This increase in SEMA3B mRNA was specific for 1,25(OH)2D3 since
neither cholecalciferol, an inactive 1,25(OH)2D3 precursor molecule, nor
24,25(OH)2D3, a vitamin D metabolite, altered SEMA3B mRNA levels (Fig. 1B
and data not shown).
127 To investigate the mechanism through which 1,25(OH)2D3 increases SEMA3B
mRNA levels, we assessed the ability of 1,25(OH)2D3 to induce SEMA3B
expression in the absence of active transcription by inhibiting transcription with
actinomycin D. As shown in Figure 1C, 1,25(OH)2D3 failed to increase SEMA3B
mRNA levels when transcription was blocked with actinomycin D. These data
suggest that 1,25(OH)2D3 regulates SEMA3B expression through a
transcriptional mechanism. Further, 1,25(OH)2D3-mediated induction of SEMA3B
required de novo synthesis of other protein factors since inhibition of protein
synthesis by cycloheximide nearly abolished the response (Fig. 1D).
Collectively, these data suggest that 1,25(OH)2D3 increases SEMA3B mRNA
levels through an active transcriptional process that requires expression of additional proteins.
SEMA3B is expressed in the osteoblasts in vivo. To determine if SEMA3B
is expressed in osteoblasts in vivo, immunohistochemistry was performed on
bone sections using an anti-SEMA3B antibody. As shown in Fig 2, SEMA3B
staining was detected in several cell types in the growth plate. SEMA3B was
selectively expressed in the calcifying and proliferating chondrocytes, while it was
not expressed in the hypertrophic chondrocytes. Both osteoblasts and
osteocytes expressed SEMA3B throughout the growth plate (Fig. 2B).
Additionally, strong staining was observed in the bone marrow, indicating that
SEMA3B is also expressed in hematopoietic stem cells (Fig. 2A).
128 SEMA3B transgenic mice are smaller and have shorter tibiae. To understand
the function of SEMA3B in bone, transgenic mice were created in which
SEMA3B expression is driven by the osteoblast-selective mouse 2.3 kb pro-α 1
(I) collagen promoter (Fig. 3A). Two lines of transgenic mice were established.
Line 2 showed robust expression of SEMA3B, but line 1 had very low expression
of the transgene only detectable by RT-PCR (Fig. 3B and data not shown).
While transgenic mice from line 1 were comparable in size to their non-
transgenic littermates (data not shown), male transgenic mice from line 2 were
smaller, displaying to a 34% reduction in body weight (Fig. 4A & B). Female
transgenic mice had a similar decrease in body weight (data not shown).
Furthermore, line 2 transgenic mice had shorter tibiae than their non-transgenic
littermates (Fig. 4C).
Some transgenic mice have altered cranial morphology. A subset of
transgenic mice from line 2 also displays abnormal skull morphology with a doming of the cranial vault (Fig. 5). This anomaly is associated with hydrocephalus, ataxia, and other neurological disturbances in some severely affected mice. Other transgenic mice, although smaller, have normally shaped skulls (Fig. 5B, left). Although the exact penetrance of this phenotype has not been determined, it has been observed in both males and females of various ages.
129 SEMA3B transgenic mice have reduced bone mineral density and altered
trabecular structure. Tibiae from male transgenic mice and their non-
transgenic littermates were analyzed by pQCT to measure bone mineral density.
Transgenic mice from line 1, which had very low transgene expression, had no
differences in bone density (data not shown). However, transgenic tibiae from
the high-expressing line 2 displayed reduced trabecular, cortical, and total bone
mineral density (Fig. 6). The most dramatic decrease was observed in the
transgenic trabecular density with a 68% reduction compared to non-trangenic
littermates.
Since the trabecular bone density of the line 2 transgenic mice was most affected
in the densitometry analysis, we next examined the microarchitecture of the
trabecular bone using high-resolution µCT (Fig. 7). Consistent with the pQCT
analysis, μCT measurements indicated that transgenic bones had decreased
trabecular bone volume (BV/TV) with an increase in the exposed surface of
trabecular bone (BS/BV; Fig. 7B). Transgenic bones had significantly fewer and
thinner trabeculae as compared to non-transgenic littermates (Fig. 7B). Further,
there was increased spacing between trabeculae and a dramatic reduction in the connectivity density in the trabecular network of the transgenic bones (Fig. 7B).
Together these results indicate that four-week transgenic animals in line 2 have reduced trabecular bone and decreased bone density.
130 SEMA3B transgenic mice have increased osteoclasts and bone resorption.
To explore the mechanism underlying the trabecular bone defect in the transgenic mice, histomorphometric analysis was performed to examine osteoblastic and osteoclastic activity (Fig. 8). Transgenic bones displayed an approximately 33% reduction in the number of osteoblasts when compared to the non-transgenic littermates (Fig. 8A). However, dynamic histomorphometric measurements of calcein- and tetracycline-labeled bone indicated that transgenic bones displayed an increase in mineral apposition rate (MAR), although this change was not statistically significant, and no change in the bone formation rate
(BFR; Fig. 8A). Thus, although there were fewer osteoblasts in the transgenic bones, it appears that osteoblastic activity and bone formation in these animals was intact. In contrast, there was a trend toward increased osteoclasts by about
2.8 fold in the transgenic bones (Fig. 8B). This increase in osteoclast number was reflected in a nearly four-fold enhancement of bone resorption, although this increase was not statistically significant (Fig. 8B). Collectively, these data suggest that the defect in bone mineralization was not caused by impaired osteoblastic activity but rather by increased osteoclastogenesis and bone resorption in SEMA3B transgenic mice.
Transgenic osteoblasts display increased differentiation and mineralization in vitro. To confirm the histomorphometric findings that osteoblastic activity was intact in SEMA3B transgenic mice, we examined transgenic osteoblast differentiation in vitro (Fig. 9). Upon stimulation with ascorbic acid and β-
131 glycerophosphate, transgenic osteoblasts expressed higher levels of alkaline phosphatase, a marker of osteoblast differentiation, as compared to osteoblasts derived from non-transgenic littermates (Fig. 9A). This increased alkaline phosphatase activity was reflected in increased numbers and size of bone nodules (Fig. 9B and data not shown). These data indicate that overexpression of SEMA3B does not impair, but rather promotes, osteoblastic differentiation in vitro. Importantly, similar results were observed in osteoblasts derived from the both the high- and low-expressing transgenic lines, indicating that even lower levels of transgene expression stimulate osteoblast differentiation.
Transgenic osteoblasts stimulate increased osteoclastogenesis in vitro. Since histomorphometric analysis suggested that osteoclastogenesis was increased in transgenic mice, we compared the ability of transgenic and non-transgenic osteoblasts to support in vitro osteoclast formation from spleen cell precursors.
Primary osteoblasts derived from transgenic and non-transgenic mice were differentiated for 15 days and then co-cultured with osteoclast precursor cells in the presence of dexamethasone and 1,25(OH)2D3. As shown in Figure 10, more osteoclasts formed in the co-cultures with transgenic osteoblasts as compared to non-transgenic osteoblasts. These in vitro data support the in vivo observation that overexpression of SEMA3B in osteoblasts stimulates increased osteoclastogenesis.
132 DISCUSSION
The primary role of the vitamin D endocrine system is to tightly control serum
concentrations of mineral ions by driving absorption calcium and phosphate from
the intestine. However, when dietary sources of calcium are lacking,
1,25(OH)2D3 stimulates bone resorption by increasing osteoclastogenesis and
activity (254). The effects of 1,25(OH)2D3 on osteoclasts are thought to be
mediated by signaling through osteoblasts. Specifically, 1,25(OH)2D3 stimulates
osteoblasts to express RANKL (243), a cytokine that is essential for
osteoclastogenesis and activity (191, 193). However, beyond increasing RANKL
signaling, the precise mechanisms governing osteoblast-osteoclast
communication modulated by 1,25(OH)2D3 are not well understood. In the
present study, we show that SEMA3B is a novel 1,25(OH)2D3 target gene and
provide evidence suggesting that SEMA3B modulates bone density in vivo.
SEMA3B was initially identified in our laboratory as a 1,25(OH)2D3-induced gene
in osteoblastic cells through a preliminary microarray screen (data not shown).
Northern blot analysis showed that SEMA3B mRNA levels were strongly induced
by 1,25(OH)2D3 in a time- and dose-dependent manner (Fig. 1). Another group also reported SEMA3B as a 1,25(OH)2D3 target in squamous cell carcinoma cells
through a large-scale microarray analysis (266), indicating that SEMA3B may be
regulated by 1,25(OH)2D3 in numerous tissues. Our data indicate that the induction of SEMA3B likely involves transcriptional stimulation since transcript levels were not increased by 1,25(OH)2D3 when transcription was blocked with
133 actinomycin D (Fig. 1C). Further, synthesis of additional protein factors were
required for this 1,25(OH)2D3-mediated response as cycloheximide blocked the induction of SEMA3B by 1,25(OH)2D3 (Fig. 1D). It is not clear whether the effect
is wholly mediated by another 1,25(OH)2D3-induced transcription factor(s) or
whether this factor(s) cooperates with VDR to stimulate SEMA3B expression.
While additional studies, including promoter analysis, are required to precisely
delineate the mechanisms underlying the regulation of SEMA3B expression,
these findings clearly show that SEMA3B is regulated by 1,25(OH)2D3 in osteoblastic cells through a mechanism that requires both active transcription and de novo protein synthesis.
Previous studies have demonstrated that SEMA3B is expressed in osteoblasts in vitro (330) and that its receptor, neuropilin, is expressed in osteoblasts and osteoclasts in vivo (331). Additionally, a related secreted semaphorin, SEMA3A, has been detected by RT-PCR in human osteoclasts (338). The current report shows that SEMA3B is expressed in osteoblasts, osteocytes, and a select population of chondrocytes in vivo (Fig. 2). Insight into the function of semaphorins the skeleton is limited to one report in which inactivation of a related protein, SEMA3A, in mice leads to wide-spread patterning abnormalities, including rib duplications (332). However, bone density was not assessed in this study. These data suggest that secreted semaphorin signaling may be involved in early skeletal patterning and development. The mechanisms underlying these bone defects in the SEMA3A knock-out mouse and the role of other secreted
134 semaphorins in regulating bone development and maintenance has not been
investigated. To determine the effect of modulating SEMA3B expression in osteoblasts in vivo, we generated a transgenic mouse model that expresses
SEMA3B under the control of an osteoblast-selective promoter.
Two lines of transgenic mice were established. Line 1 had relatively low levels of
expression, only detectable by RT-PCR (data not shown), whereas line 2
displayed high levels of expression (Fig. 3B). Line 2 had a dramatic phenotype,
displaying decreased body mass and tibial length (Fig. 4). These data indicate
that high levels of SEMA3B expression restrict longitudinal bone growth. Some
transgenic mice from line 2 also displayed malformed skulls (Fig. 5). The cranial
vault appeared domed in several transgenic mice, and hydrocephalus developed
in severely affected mice. These hydrocephalic mice were ataxic, presumably
due to neurological damage from the abnormal skull shape. This domed skull
morphology is similar to mutant mice that have craniosynostosis, a premature
closure of the cranial sutures (339, 340). Although a more detailed analysis of
the skull bone morphology is required to elucidate the underlying defects, these
observations suggest that overexpression of SEMA3B alters intramembraneous
bone formation and skull bone development.
pQCT analysis of the tibiae from four-week male transgenic mice in line 2
showed a dramatic reduction in bone mineral density (Fig. 6). The most
dramatic decrease was observed in the transgenic trabecular bone, which
135 exhibited a 68% reduction in density as compared with non-transgenic littermates. Consistent with the pQCT data, μCT analysis revealed that the
trabecular microarchitecture was dramatically disrupted in transgenic mice (Fig.
7). SEMA3B transgenic tibiae displayed an overall reduction in the trabecular
bone volume (BV/TV), an increase in the amount of exposed trabecular surface
(BS/BV), decreased numbers of thinned trabeculae, increased spacing between trabeculae, and dramatically diminished connectivity density within the trabecular lattice (Fig. 7B). These data strongly support a role for osteoblast-derived
SEMA3B in negatively regulating trabecular bone mass and density.
To investigate the mechanisms underlying the osteopenic effects of SEMA3B
overexpression, histomorphometric analysis and in vitro culture systems were
used to evaluate the numbers and function of osteoblasts and osteoclasts.
Histomorphometry revealed that transgenic bones had significantly decreased
osteoblast numbers, but, surprisingly displayed normal, if not somewhat
increased, osteoblastic activity as reflected by the mineral apposition rate (Fig.
8). Thus, even though there are less total osteoblasts in the transgenic bones, it
is possible that each osteoblast is more active to result in a modest increase in
the overall mineral apposition rate. This hypothesis is supported by the in vitro
osteoblast analysis, which indicated that transgenic osteoblasts differentiate and mineralize to a greater extent than non-transgenic osteoblasts (Fig. 9). These findings suggest that SEMA3B overexpression drives osteoblast differentiation.
Furthermore, histomorphometric analysis indicated a strong trend toward
136 increased number of osteoclasts and an elevated amount of bone surface
undergoing resorption (Fig. 8). Consistent with these findings, in vitro co-culture
studies showed that transgenic osteoblasts stimulated increased
osteoclastogenesis as compared to non-transgenic osteoblasts (Fig. 10). Taken
together, these data suggest that the osteopenia observed in SEMA3B
transgenic mice was not due to impaired osteoblast function but rather a result of
increased osteoclastogenesis and osteoclast activity. These findings are
somewhat similar to the the phenotype observed in osteopenic mice carrying an
osteoblast-selective transgene driving expression of runx2 (341), a transcription
factor essential for osteoblast differentiation (138, 139). Although runx2-
overexpressing mice display normal mineral apposition rates, they have
increased osteoclastogenesis and bone turnover leading to low bone mass (341).
The current study suggests that osteoblast-derived SEMA3B may stimulate
osteoclast formation and activity. This stimulation may be direct since
osteoclasts express neuropilin-1, one of the receptors for secreted semaphorins
(331). In fact, recent studies have shown that rac1, a signaling effector of semaphorin signaling (342), is required for osteoclast differentiation and bone resorption (343, 344). Thus, SEMA3B may bind neuropilin expressed on the surface of osteoclasts and signal through rac1 to directly promote osteoclastogenesis. Conversely, as osteoblasts also express neuropilin-1 (331),
SEMA3B may act in an autocrine fashion to induce expression of cytokines such as RANKL by osteoblasts that, in turn, promote osteoclastogenesis and
137 resorption. A more thorough understanding of semaphorin signaling osteoblasts
and osteoclasts is required to elucidate the mechanism through which SEMA3B
stimulates osteoclast formation and bone resorption.
In summary, this study demonstrates that SEMA3B is a novel target of
1,25(OH)2D3 in osteoblastic cells. Transgenic mice that robustly overexpress
SEMA3B in osteoblasts displayed a profound reduction in trabecular bone mass and density. This defect appears to be caused by elevated osteoclastogenesis and bone resorption, indicating the SEMA3B may be directly or indirectly stimulating osteoclast formation and activity. The current study represents the first demonstration that a member of the semaphorin family regulates bone mineral density and uncovers a potential role for SEMA3B in modulating osteoclastogenesis and bone resorption.
ACKNOWLEDGMENTS
This work was supported in part by NIH Grants R01 DK50348 and R01 DK
53980 (to P.N.M.), by an award from the Medical Scientist Training Program NIH
Grant T32 GM007250 (to A.L.M.S.), and by a Pharmaceutical Manufacturers'
Association (PhRMA) Foundation Pre-Doctoral Fellowship (to A.L.M.S.). The
authors would like to thank Dr. Benoit de Crombrugghe for providing the
2300lacZ plasmid, Dr. John D. Minna for sharing his anti-SEMA3B antibody and
the pcDNA3-SEMA3B plasmid, Dr. Milan Uskokovic for providing 1,25(OH)2D3,
and Ms. Meika Moore for excellent technical assistance. We would also like to
138 thank members of the laboratories of Dr. John H. Nilson and Dr. Ruth A. Keri for
sharing their technical expertise and for helpful discussions. We acknowledge the
Case School of Medicine Comprehensive Cancer Center for performing the
demineralized bone histology. We are grateful to Ms. Patty Lott and her staff at
the University of Alabama at Birmingham, Center for Metabolic Bone Disease,
Histomorphometry and Molecular Analysis Core Laboratory, National Institutes of
Health Grant P30-AR46031, for performing the mineralized histology and
histomorphometry.
139 Figure IV-1
1,25(OH)2D3 induces SEMA3B expression in a time- and dose-dependent
manner through a mechanism that requires both active transcription and
protein synthesis.
A, MG-63 cells were treated for the indicated times with 10 nM 1,25(OH)2D3 or 10 nM cholecalciferol. B, MG-63 cells were treated with either ethanol vehicle control (-) or 0.1 to 100 nM 1,25(OH)2D3 for 6 hours. C, MG-63 cells were pre-
treated with methanol vehicle control (- ACTD) or 1 ug/ml actinomycin D (+
ACTD) for 1 hour. Cells were then treated with ethanol control (Et) or 10 nM
1,25(OH)2D3 (1,25) for 6 hours. D, MG-63 cells were pre-treated with ethanol
control (-CHX) or 10 ug/ml cycloheximide (+ CHX) for 1 hour. Cells were then
treated with ethanol control (Et) or 10 nM 1,25(OH)2D3 (1,25) for 6 hours. mRNA
was analyzed by Northern blots for SEMA3B and for β-actin in all panels.
140 A B 10 nM 10 nM
1,25(OH)2D3 chol -0.1110100 Time (hr): 036122424 nM 1,25(OH)2D3:
SEMA3B SEMA3B
β-actin β-actin
C D -ACTD +ACTD -CHX +CHX Et 1,25 Et 1,25 Et 1,25Et 1,25
SEMA3B SEMA3B
β-actin β-actin
141 Figure IV-2
SEMA3B is expressed in the long bones of mice.
A, Tibiae from wild-type 8-week-old male mice were formalin-fixed, decalcified in formic acid, embedded in paraffin, and sectioned. Sections were immunostained for SEMA3B with α-SEMA3B-3288 (brown). B, Higher-magnification view of A. showing osteoblasts lining the marrow surface of the secondary ossification center (arrows) and osteocytes trapped in the mineralized matrix (arrowheads).
142 A
10X
B
20X
143 Figure IV-3
Establishment of transgenic mice overexpressing SEMA3B in osteoblasts.
A, Schematic of transgene. The osteoblast-selective 2.3-kb mouse pro-α 1(I)
collagen promoter was used to drive human SEMA3B expression in osteoblasts.
B, Transgene is expressed robustly in line 2 of transgenic mice. Total humerus
RNA from transgenic (TG) and non-transgenic (NTG) mice was analyzed by
Northern blot for SEMA3B and β-actin expression.
144 A
2.3 α1(I)col sema3B
B TG
FEMALE MALE Line: 22 1 2112
SEMA3B
β-actin
145 Figure IV-4
Transgenic mice have reduced body weight and shorter bones.
A, Photograph of 31 day old male non-transgenic (NTG) and transgenic (TG)
littermates from line 2. B, Growth curve of male NTG and TG mice from line 2.
Data represent mean + SEM. n = at least 5 in each group. *, p < 0.01 by
student’s t-test. C, pQCT measurement of left tibia length in line 2. n = 8 in each
group. *, p < 0.01 by student’s t-test.
146 A B 20 NTG NTG TG 18 * 16 TG 14 * 12 * 10 * * 8 * 6 * body weight (g) 4 2 0 0 5 10 15 20 25 30 age (days)
C 8.2 8.1 8 7.9 7.8 7.7 * 7.6 7.5 tibia length (cm) 7.4 7.3 7.2 NTG TG
147 Figure IV-5
Altered cranial morphology in some transgenic mice.
A, Newborn NTG and TG littermates from line 2. Note the domed head in the TG
mouse. B, Dissected skulls of 4-week female NTG and TG littermates from line
2. Note that some TG skulls displayed a normal morphology (left TG skull), while others displayed a doming and bulging of the cranial bones (right TG skull).
148 A
NTG TG
B
TG NTG TG
149 Figure IV-6
Transgenic mice have decreased bone mineral density. pQCT analysis of total, trabecular, and cortical bone density of tibae from 31 day old male NTG and TG littermates from line 2. Data represent mean + SEM. n =
7-8 in each group. *, p < 0.02 by student’s t-test.
150 1000 ) 3 800 * NTG 600 TG
400 * 200 Bone Density (mg/cm * 0 Total Trabecular Cortical
151 Figure IV-7
Transgenic mice have diminished trabecular bone.
A, μCT cross-sectional images of representative tibiae from 31 day old male NTG
and TG littermates from line 2. B, Quantitative μCT shows decreased trabecular
bone volume. BV/TV, trabecular bone volume expressed as a percentage of
total tissue volume; BS/BV, bone surface expressed as a percentage of total
trabecular volume; Trab. #, number of trabeculae per mm; Trab. Th., trabecular
thickness; Trab. spacing, trabecular spacing; Conn. Dens., connectivity density of
trabecular bone. Data represent mean + SEM. n = 8 in each group, *, p < 0.01 by student’s t-test.
152 A
NTG TG B 0.14 80 * 0.12 0.10 60 0.08 40 0.06
0.04 * BS/BV (%)
BV/TV (%) 20 0.02 0.00 0 NTG TG NTG TG
4.0 0.06 3.0 0.04 * 2.0 *
1.0 0.02 Trab. # (1/mm) Trab. Th. (mm) 0.0 0.00 NTG TG NTG TG ) 0.6 3 140 120 0.5 * 100 0.4 80 0.3 60 0.2 40 * 0.1 20 Trab. spacing (mm) 0.0 Conn. Dens. (1/mm 0 NTG TG NTG TG
153 Figure IV-8
Normal osteoblast function but potentially increased bone resorption in transgenic mice.
A, Decreased osteoblast number, but normal osteoblast function in transgenic mice. Histomorphometry measurements were performed on sections of tibiae from NTG and TG 31 day old male mice that had been double-labeled with calcein and tetracycline. N.Ob/BS, number of osteoblasts per mm of bone surface; MAR, mineral apposition rate; BFR, bone formation rate. B, Trend toward increased osteoclast number and resorption in TG mice. N.Oc/BS, number of osteoclasts per mm of bone surface; ES/BS, percentage of bone surface involved in resorption. Data represent mean + SEM. n = 4 in each group, *, p = 0.05 by student’s t-test.
154 A
16 N.Ob/BS 3.0 MAR (μm/day) 0.6 BFR (μm/day) 14 2.5 0.5 12 2.0 0.4 10 * 8 1.5 0.3 6 1.0 0.2 4 0.5 0.1 2
0 0.0 0.0 NTG TG NTG TG NTG TG
B 8 0.8 N.Oc/BS ES/BS 7 0.7 6 0.6
0.5 5
0.4 4
0.3 3
0.2 2
0.1 1
0.0 0 NTG TG NTG TG
155
Figure IV-9
Transgenic osteoblasts show increased differentiation and mineralization
in vitro.
Primary osteoblasts were isolated from the calvaria of newborn transgenic or non-transgenic pups. Osteoblasts were stimulated to differentiate by supplementing the culture medium with 50 μg/ml ascorbic acid and 10 mM β- glycerophosphate. A, Alkaline phosphatase activity in cellular lysates taken at the indicated times was determined by measuring conversion of p-nitrophenyl phosphate to p-nitrophenol. Activity was normalized to protein concentration.
Data represent the mean of triplicate well + SEM. B, Mineralized nodules were
visualized by staining fixed cells in triplicate with Alizarin red S dye. Data
represent the mean of triplicate well + SEM.
156 A 3
NTG 2.5 TG 2
1.5
1
ALP activity/ng protein 0.5
0 day 6 day 15 day 25
B 5
4
3
2 # nodules/well 1
0 NTG TG
157 Figure IV-10
Transgenic osteoblasts stimulate increased osteoclastogenesis.
Primary osteoblasts derived from newborn transgenic or non-transgenic mice were stimulated to differentiate by supplementing the culture medium with 50
μg/ml ascorbic acid and 10 mM β-glycerophosphate for 15 days. Osteoclast precursors derived from the spleens of wild-type adult mice were added to the osteoblast cultures and grown for an additional 8 days in the presence of 100 nM dexamethasone and 10 nM 1,25(OH)2D3. Multinucleated osteoclasts were visualized by staining for TRAP activity as described in Materials and Methods.
A, Representative fields of TRAP-stained co-cultures from NTG and TG osteoblasts and spleen cells. B, Number of TRAP+ multinucleated (> 3 nuclei) cells per well. Data represent the mean of triplicate wells + SEM.
158 A
NTG TG
B 500
400
300
200
# osteoclasts/well 100
0 NTG TG
159 CHAPTER V
INVESTIGATING THE FUNCTION OF THROMBOMODULIN, A 1,25(OH)2D3- REGULATED ANTICOAGULANT PROTEIN, IN OSTEOBLASTS AND BONE
INTRODUCTION
1,25(OH)2D3 is the most bioactive, or hormonal, form of vitamin D. This hormone signals through the vitamin D receptor (VDR), a member of the nuclear receptor superfamily of ligand-activated transcription factors (254, 255). 1,25(OH)2D3-
bound VDR heterodimerizes with retinoid X receptor (RXR), associates with the
promoter regions of target genes, and recruits a multiprotein complex including
coactivators and the core transcriptional machinery to initiate transcription of
1,25(OH)2D3-regulated genes. A major function of 1,25(OH)2D3 is to preserve
bone mineralization by promoting calcium absorption from the intestine.
1,25(OH)2D3 also acts locally in the bone by modulating the activity of
osteoblasts, the bone-forming cells, and osteoclasts, the bone-resorping cells
(232, 256). Despite these functions of 1,25(OH)2D3, relatively few target genes,
such as those encoding extracellular matrix proteins and osteoclast stimulating
factors (236, 243, 319, 320), have been described. The present study
characterizes the regulation and function of thrombomodulin as a 1,25(OH)2D3- induced gene in osteoblastic cells.
Thrombomodulin is a transmembrane protein that primarily functions as an anticoagulant at the surface of vascular endothelial cells (345, 346).
Thrombomodulin binds thrombin, the central enzyme in the coagulation cascade
160 that cleaves fibrinogen and converts it into fibrin. By interacting with thrombin, thrombomodulin interferes with its protease activity and prevents fibrin formation.
Thrombin-bound thrombomodulin then activates protein C, which proteolytically destroys activated factors V and VIII of the clotting cascade. Thus, thrombomodulin functions at numerous steps in the clotting pathway to prevent excess blood coagulation (345, 346).
In the adult, thrombomodulin is expressed outside the vascular system, including the brain meninges, the lung, and keratinocytes (347). Developmental studies indicate that thrombomodulin is expressed in the neuroepithelium, lung, and bone (347). Since it is expressed in several non-vascular tissues during development, it is likely that thrombomodulin may exert other functions outside of its prominent role in hemostasis. This hypothesis is supported by studies in thrombomodulin-deficient mice. Thrombomodulin-null embryos die before establishment of a functioning cardiovascular system (348). However, mice harboring a point mutation in thrombomodulin that abolishes its anticoagulant activity are still viable (349), implying that the developmental requirement for thrombomodulin is not related to its regulation of the clotting cascade. Thus, it is likely that thrombomodulin acts in numerous other tissues through mechanisms unrelated to its anticoagulation activity.
Growing evidence supports a role for thrombomodulin in osteoblast differentiation. One study indicated that thrombomodulin protein levels are
161 induced by 1,25(OH)2D3 in human osteoblastic cells (350). Transcriptional profiling showed that thrombomodulin expression increases during sonic hedgehog-stimulated osteoblastic differentiation in murine pluripotent mesenchymal cells (351). Similarly, increases in thrombomodulin mRNA expression were detected by microarray analysis in murine MC3T3 osteoblasts stimulated to differentiate with bone morphogenic protein (352). Collectively, these studies indicate that thrombomodulin may be involved in osteoblast differentiation since multiple factors that promote osteoblast differentiation stimulate thrombomodulin expression. However, neither the transcriptional regulation nor functional implications of thrombomodulin expression patterns were examined in these reports.
In the current study, we demonstrate that 1,25(OH)2D3 strongly increased thrombomodulin mRNA levels and protein levels in osteoblastic cells through a mechanism that required both active transcription and de novo protein synthesis.
Cells stably overexpressing thrombomodulin displayed an increase in proliferation, suggesting that thrombomodulin promotes osteoblast cell growth.
Finally, we have developed an osteoblast-selective thrombomodulin knockout mouse model to study the in vivo functions of thrombomodulin in the bone.
Although preliminary analyses of these animals revealed no obvious bone abnormalities, further studies are clearly required to determine if these animals have any alterations in bone development or maintenance.
162 MATERIALS AND METHODS
Cell culture. MG-63 human osteosarcoma cells were maintained in
modified essential media supplemented with 10% fetal bovine serum. For
experiments using 1,25(OH)2D3 compounds, cells were grown in media supplemented with charcoal-stripped bovine calf serum for three days prior to the experiment.
Northern blot analysis. mRNA was isolated from MG-63 cells with the
FastTrack system (Invitrogen, Carlsbad, CA) according to the manufacturer’s
instructions. mRNA was separated on a formaldehyde/agarose gel and
transferred to a Duralon membrane (Stratagene, La Jolla, CA) by capillary action.
α32P-labeled probes were synthesized using the Prime-A-Gene kit (Promega,
Madison, WI) according to the manufacturer’s instructions and hybridized to the
blots using standard methods.
Western blot analysis. Protein extracts were prepared by solubilizing cells in
SDS buffer. 250 ng of purified human thrombomodulin was also used a positive
control (American Diagnostica, Greenwich, CT). Protein was separated by SDS-
PAGE and subjected to western blot analysis using standard methods.
Thrombomodulin was detected by incubating the membrane with a 1:100 dilution of a monoclonal anti-thrombomodulin antibody (American Diagnostica).
163 Transfection and stable lines. pcDNA3-thrombomodulin was constructed by
subcloning the human thrombomodulin coding sequence from puc19TM15
(ATCC, Manassas, VA) into pcDNA3 (Invitrogen). MG-63 cells were transfected
with Fugene (Roche Applied Science, Indianapolis, IN). After 48 hours, cells
were plated in G418-containing media (1 mg/ml) to select for transfected cells for
two weeks. G418-resistant colonies were expanded and analyzed for
thrombomodulin expression by western blots as described above.
Proliferation assay. MG-63 cells were plated at a density of 2 X 104 cells/well of a six well plate. Viable cells were quantified by counting trypan blue- excluding cells with a hemocytometer daily for five days. Data presented represent the mean of triplicate wells -/+ standard deviation of a representative experiment performed twice.
Animals and μCT imaging. All experiments involving mice were approved by the Institutional Animal Care and Use Committee (IACUC) at Case Western
Reserve University. Thrombomodulin-loxP (TMlox) mice in the C57Bl/6
background (353) were kindly provided by Dr. Harmut Weiler of the Department
of Physiology of the Medical College of Wisconsin. Mice harboring a transgene
containing the cre recombinase under the control of the 2.3-kb proximal fragment
of the mouse pro-alpha 1(I) collagen promoter (cre) in the FVB/NTac background
(354) were obtained from the Mutant Mouse Regional Research Center of the
University of California at Davis. Mice were genotyped by PCR as described for
TMlox (353) and for cre (354). Cre mice were back-crossed one generation into
164 C57Bl/6, and then bred with homozygous TMlox (TMlox/lox) mice. TMlox/+/cre- positive progeny were crossed with TMlox/lox mice to generate experimental mice:
TMlox/lox/cre-negative and TMlox/lox/cre-positive mice that were approximately
82.5% C57Bl/6 and 17.5% FVB/NTac. These animals were weighed and then
killed at 5 weeks of age by CO2 asphyxiation. Bones were dissected and fixed in
10% neutral-bufferered formalin overnight at room temperature, and then transferred to 70% ethanol and stored at 4° C. Micro-computerized tomography
(μCT) was performed by the laboratory of Dr. Kimerly Powell from the
Department of Biomedical Engineering at the Lerner Research Institute of the
Cleveland Clinic Foundation using a custom designed imaging system. Images
with a spatial resolution of 18 μm were obtained by collecting one-hundred and
eighty 512 X 512 12-bit projection radiographs at 1° intervals around one-half of
the tibiae. Final images were processed and reconstructed using custom in-
house software.
RESULTS
1,25(OH)2D3 induces thrombomodulin expression in a time- and dose-dependent
manner. A preliminary microarray screen showed that thrombomodulin
mRNA was increased by 10 nM 1,25(OH)2D3 after 6 hours by approximately 15
fold (see Introduction). This response was confirmed by Northern blot analysis.
Whereas thrombomodulin mRNA was barely detectable in vehicle treated cells, it
was induced as early as three hours by 10 nM 1,25(OH)2D3 (Fig. 1A).
Thrombomodulin expression continued to rise and was maximal at 12 hours.
165 This induction was specific for 1,25(OH)2D3 since neither the precursor molecule
vitamin D3 (cholecalciferol) nor the metabolite 24,25(OH)2D3 affected
thrombomodulin expression (Fig. 1A and data not shown). 1,25(OH)2D3 also increased thrombomodulin mRNA levels in a dose-dependent manner (Fig. 1B).
As little as 1 nM 1,25(OH)2D3 stimulated thrombomodulin expression, and mRNA
levels continue to increase up to 100 nM 1,25(OH)2D3.
1,25(OH)2D3-mediated induction of thrombomodulin requires both active
transcription and de novo protein synthesis. To determine if the 1,25(OH)2D3 stimulation of thrombomodulin expression was a transcriptional response, MG-63 cells were treated with actinomycin D, an inhibitor of mRNA synthesis. As shown in Figure 2A, actinomycin D treatment completely abolished 1,25(OH)2D3- mediated induction of thrombomodulin mRNA levels. In contrast, inhibition of protein synthesis with cycloheximide had little effect on the stimulation of thrombomodulin expression by 1,25(OH)2D3. These results indicate that
1,25(OH)2D3 increases steady-state levels of thrombomodulin mRNA through a mechanism that requires active transcription but does not require de novo protein synthesis.
1,25(OH)2D3 induces thrombomodulin expression at the protein level. To confirm that the induction of thrombomodulin mRNA by 1,25(OH)2D3 is
concomitant with an increase protein levels, western blot analysis was performed
on 1,25(OH)2D3-treated MG-63 cells. As shown in Figure 3, in the absence of
166 ligand stimulation, thrombomodulin protein was undetectable. However, upon
1,25(OH)2D3 stimulation a robust thrombomodulin protein signal was observed,
indicating that 1,25(OH)2D3 also increases protein levels of thrombomodulin.
Stable lines overexpressing thrombomodulin have an increased proliferation rate.
To understand the functional implications of 1,25(OH)2D3-mediated induction of
thrombomodulin expression in MG-63 cells, we created stably-transfected cell
lines that maintain a high level of thrombomodulin expression in the absence of
1,25(OH)2D3 (Fig. 4A). Compared to control stable lines, these cells proliferate at a more rapid rate (Fig. 4B). These observations indicate that thrombomodulin may promote osteoblastic cell proliferation.
Creating an osteoblast-selective thrombomodulin knockout model. Knockout
animals are powerful tools to study the function of individual genes in a
physiological relevant context. Since thrombomodulin-deficient mice die very
early in embryogenesis (348), it is not possible to study the function of
thrombomodulin in the bone using this model. Therefore, we created osteoblast-
selective conditional knockout mice by crossing mutant mice carrying a thrombomodulin-floxed allele (353) to transgenic mice that express the cre recombinase under the control of the osteoblast-selective 2.3 kb promoter region mouse pro-alpha 1(I) collagen gene (354). 5-week old conditional knockout
(TMlox/lox/cre-positive) mice have body masses similar to the cre-negative control
(TMlox/lox/cre-positive) mice (Fig. 5). To examine if deletion of thrombomodulin in
167 osteoblasts affects bone development and mineralization, μCT imaging was performed on 5-week female TMlox/lox/cre-negative and TMlox/lox/cre-positive mice.
This preliminary analysis did not reveal any striking abnormalities in the
TMlox/lox/cre-positive mice (data not shown).
DISCUSSION
Microarray analysis has elucidated numerous potential 1,25(OH)2D3-regulated
genes in several cell types (266, 282-287, 355). Such studies have provided the
framework to begin functionally characterizing target genes and to more fully
understand the diverse effects of 1,25(OH)2D3. Our microarray screen identified
thrombomodulin as a 1,25(OH)2D3-induced gene in human MG-63 osteoblastic
cells. In addition to characterizing the regulation of thrombomodulin, the current
study has begun to elucidate the function of thrombomodulin in osteoblasts using
both in vitro and in vivo approaches.
Northern blot analyses confirmed the preliminary microarray data and showed
that 1,25(OH)2D3 increased thrombomodulin mRNA levels in a time- and dose-
dependent manner (Fig. 1). This increase in mRNA levels was reflected by a
concomitant increase in protein levels (Fig. 3). These findings are consistent
with a previous report that described a 1,25(OH)2D3-mediated induction of
thrombomodulin protein levels in osteoblastic cells (350). Thrombomodulin also
has been identified as a 1,25(OH)2D3 target gene in keratinocytes (355),
myelogenous leukemia cells (356), and monocytes (357).
168
Although it is clear from these studies and from our findings that thrombomodulin
levels increase upon 1,25(OH)2D3 stimulation in multiple cell types, the exact
mechanisms underlying this induction remain unknown. Blockade of
transcription with actinomycin D completely abolished 1,25(OH)2D3-mediated
induction of thrombomodulin, indicating that 1,25(OH)2D3 increases
thrombomodulin mRNA levels through a transcriptional mechanism (Fig. 2A).
However this response did not require de novo protein synthesis (Fig. 2B),
suggesting that 1,25(OH)2D3 directly regulates thrombomodulin expression by binding to and activating its promoter. One report regarding the effect of
1,25(OH)2D3 on thrombomodulin expression concluded that a retinoid response
element (RARE) in the thrombomodulin promoter represented a composite
element that also mediated the 1,25(OH)2D3 response (356). However, when
this RARE was cloned upstream of a reporter gene and stimulated with
1,25(OH)2D3 in myelogenous leukemia cells, less than a 1.5-fold increase in
promoter activity was observed. In our studies, a 3.4 kb fragment of the
thrombomodulin promoter that contains this RARE does not respond to
1,25(OH)2D3 in osteoblastic cells (data not shown). It is likely that the true
vitamin D response element (VDRE) lies elsewhere in thrombomodulin promoter.
Consequently, further analyses are required to fully understand how 1,25(OH)2D3 controls thrombomodulin expression in various cell types.
169 To determine the impact of constitutively elevated thrombomodulin protein levels
on osteoblastic cell function, stable MG-63 cell lines were created that
overexpress thrombomodulin (Fig. 4A). These lines displayed an increased
proliferation rate when compared to control vector-transfected stable line (Fig.
4B). These data suggest that thrombomodulin promotes cellular proliferation in
osteoblastic cells. However, most other studies suggest that thrombomodulin
inhibits proliferation. For example, overexpression of thrombomodulin blocks
proliferation of endothelial cells (358) and melanoma cells (359) in culture. In
most human cancers, loss of thrombomodulin expression is correlated with more
malignant tumors and poorer prognoses (360). The thrombomodulin promoter is
methylated in gastric cancer cell lines (361) and in several melanomas (362).
Furthermore, overexpression of thrombomodulin in melanoma cells restricts the growth of these cells in nude mice (363). Only one additional study indicates a role for thrombomodulin in stimulating cell growth. These authors found that antisense-mediated knockdown of thrombomodulin expression in A549 lung adenocarcinoma cells results in diminished cell proliferation (364). Given the variety of cell types analyzed in these reports, it is possible that thrombomodulin acts in a cell-type selective manner to either positively or negatively regulate proliferation. Our findings that thrombomodulin promotes cell proliferation are somewhat surprising given that 1,25(OH)2D3 inhibits cell growth in osteoblasts
(see Figure II-6 in Chapter II). However, since 1,25(OH)2D3 induces a number of other genes that negatively control proliferation (259, 260), thrombomodulin may
170 function to feedback on other 1,25(OH)2D3-stimulated pathways to limit the extent
of cell cycle arrest.
The function of thrombomodulin in osteoblasts is completely unexplored. Two
recent microarray studies indicate that thrombomodulin expression rises during
osteoblastic differentiation (351, 352). It is notable that these studies utilized two
distinct models of osteoblast differentiation, highlighting the potential of
thrombomodulin as a key gene involved in this process. In the current study, we
have created an osteoblast selective thrombomodulin knockout model to
examine the effect of thrombomodulin in bone. Our preliminary analyses did not
reveal any obvious abnormalities in the body weight of both sexes or bone
structure of females of the TMlox/lox/cre-positive mice (Fig. 5 and data not shown).
However, the μCT imaging performed was not quantitative but rather qualitative,
providing an overall assessment of the morphology of the bones. Further
quantitative analyses are required to carefully evaluate the bone for alterations in
cortical and trabecular structure and mineral density. Additionally, we have only
examined 5-week females in the current study. A more complete analysis should
include males and other ages since the phenotype could be sex- or age-
dependent or both.
In summary, this study demonstrates that thrombomodulin is a 1,25(OH)2D3 target gene in osteoblastic cells. 1,25(OH)2D3-mediated induction involved active
transcription and did not require new protein synthesis. Osteoblastic cell lines
171 stably overexpressing thrombomodulin displayed an accelerated proliferation rate
suggesting that this protein promotes osteoblastic cell growth. Finally, we report
establishment of an osteoblast-selective thrombomodulin knockout mouse model. Although these mice did not have any obvious bone abnormalities, they represent an important tool to further analyze the in vivo function of
thrombomodulin in the bone.
172 Figure V-1
1,25(OH)2D3 induces thrombomodulin expression in a time- and dose-
dependent manner.
A, MG-63 cells were treated for the indicated times with 10 nM 1,25(OH)2D3 or 10 nM cholecalciferol (chol). mRNA was analyzed by Northern blots for thrombomodulin and β-actin. B, MG-63 cells were treated with ethanol vehicle control (-) or 0.1 to 100 nM 1,25(OH)2D3 for 6 hours. mRNA was analyzed by
Northern blots for thrombomodulin and β-actin.
173 A 10 nM 10 nM
1,25(OH)2D3 chol time (hr): 0 3 6 1224 24
TM
β-actin
B
nM 1,25(OH)2D3: - 0.1 1 10 100
TM
β-actin
174 Figure V-2
1,25(OH)2D3 induction of thrombomodulin requires active transcription but
not de novo protein synthesis.
A, MG-63 cells were pre-treated with methanol vehicle control (- ACTD) or 1
ug/ml actinomycin D (+ ACTD) for 1 hour. Cells were then treated with ethanol
control (Et) or 10 nM 1,25(OH)2D3 (1,25) for 6 hours. mRNA was analyzed by
Northern blots for thrombomodulin and β-actin. B, MG-63 cells were pre-treated
with ethanol control (-CHX) or 10 ug/ml cycloheximide (+ CHX) for 1 hour. Cells were then treated with ethanol control (Et) or 10 nM 1,25(OH)2D3 for 6 hours. mRNA was analyzed by Northern blots for thrombomodulin and β-actin.
175 A -ACTD + ACTD
Et 1,25 Et 1,25
TM
β-actin
B - CHX + CHX Et 1,25 Et 1,25
TM
β-actin
176
Figure V-3
1,25(OH)2D3 increases protein levels of thrombomodulin.
MG-63 cells were treated with 10 nM 1,25(OH)2D3 for the indicated times. Cell extracts and purified human thrombomodulin (as a positive control) were separated by SDS-PAGE and subjected to western blot analysis with a monoclonal anti-human thrombomodulin antibody.
177 Time (hr): 0 24 48 Purified Ligand (10-8 M): Et Et 1,25 Et 1,25 TM
WB: anti-TM
178 Figure V-4
Stable MG-63 cell lines overexpressing thrombomodulin display an
increased proliferation rate.
A, MG-63 cells were transfected with either empty vector (pCDNA) or pcDNA3-
thrombomodulin (TM), and stable transfectants were selected for two weeks.
Lines were screened for thrombomodulin expression by western blot analysis
using a monoclonal anti-human thrombomodulin antibody. B, Control (pcDNA) or
thrombomodulin (TM) stable lines were seeded at equal densities and grown for
five days. Trypan blue-excluding viable cells were counted each day for five days using a hemocytometer.
179 A
TM TM Purifed pcDNA 150 251 TM 1,25: -+ - + -+
WB: anti-TM
B
50 TM251
) 40 4 TM150
30 pcDNA100
20 pcDNA101 Cell Count (X 10 10
0 012345
Days in Culture
180 Figure V-5
Mice with an osteoblast-selective knockout of thrombomodulin weigh the same as control littermates.
TMlox/lox/cre-negative (lox/negative) and TMlox/lox/cre-positive (lox/positive) mice were weighed at five weeks of age. Data represent the mean + standard deviation. n > 4 in each group. No significant differences were detected between the lox/negative and lox/positive groups within each sex (p > 0.05 by student’s t-test).
181 lox/negative 25 lox/positive 20
15
10
Body Weight (g) Body Weight 5
0 Females Males
182 CHAPTER VI
SUMMARY AND FUTURE DIRECTIONS
The current project had identified several 1,25(OH)2D3-regulated genes in osteoblastic cells. A combination of in vitro and in vivo studies has begun to
elucidate the molecular mechanisms governing the regulation of these genes and
has revealed several novel functions of target genes in osteoblasts and in the
bone. Importantly, I have developed numerous tools that will be useful in
providing a more complete picture of how 1,25(OH)2D3 regulates a multitude of
genes and the pleiotropic effects of such target genes in skeletal cells and tissue.
MN1
As described in Chapter II, 1,25(OH)2D3 induces the expression of MN1. This transcription factor functions as a coactivator for VDR and inhibits osteoblastic cell proliferation. 1,25(OH)2D3-mediated increases in MN1 mRNA levels are likely mediated through a direct transcriptional response since they are dependent on active mRNA synthesis but not on expression of new proteins (Fig.
II-2). MN1 cooperates with the SRC family coactivators to synergistically stimulate VDR-mediated transcription (Fig. II-5). Overexpression of MN1 potently inhibits osteoblastic cell proliferation by modulating the cell cycle (Fig. II-
6 and Fig. II-7).
183 The exact mechanism through which 1,25(OH)2D3 stimulates MN1 transcription is not clear. Sequence analysis of the presumed 5’ proximal regulatory region of the human MN1 locus reveals no canonical VDREs, but the region does contain half-site elements that may mediate the 1,25(OH)2D3 response. A 2 kb fragment of the potential promoter region has been cloned upstream of a luciferase reporter gene, but it does not appear to respond to 1,25(OH)2D3 in MG-63 osteoblastic cells (X. Zhang, unpublished observations). It is possible that the response element lies elsewhere, such as further upstream in the promoter sequence or in exonic or intronic sequences. Identification of such sequences will require cloning of larger promoter fragments into reporter genes. For instance, a bacterial artificial chromosome containing the whole genomic sequence of MN1 could be tested for responsiveness to 1,25(OH)2D3 (365).
Then the exact responsive sequence or sequences could be further narrowed down.
Although we have shown that MN1 enhances the transcriptional activity of VDR
(Fig. II-3) potentially through cooperation with SRC proteins (Fig. II-5), the molecular details of these effects remain unknown. Co-immunoprecipitation analyses with overexpressed proteins in COS-7 cells have failed to provide consistent evidence of an interaction between MN1 and either VDR or SRCs (A.
Sutton, C. Gu, and X. Zhang, unpublished observations). Further, GST pull- down assays with isolated proteins indicated that MN1 does not directly interact with VDR (X. Zhang, unpublished observations). Mammalian two-hybrid studies
184 have proven uninterpretable due to the strong intrinsic transactivation activity of
MN1 in these assays (X. Zhang, unpublished observations). Although these
approaches are powerful tools for studying protein-protein interactions, they rely
on overexpressed proteins or high concentrations of purified proteins. Thus, they
may not represent physiologically relevant conditions under which weaker
protein-protein interactions can be influenced by the formation of intricate
multiprotein complexes in the native chromatin environment of transcriptional
regulatory regions. Therefore, to better determine if MN1 associates with the
VDR transcriptional complex, chromatin immunoprecipitation (ChIP) assays
should be performed in osteoblastic cells. Currently, polyclonal antibodies against MN1 are being generated that will be useful in these and other studies.
ChIP assays are powerful approaches to study the association of endogenous proteins with hormone-responsive regions of native genes (69). Given the stimulatory effects of MN1 on VDR transactivation, I expect that ChIP analyses will reveal an association between MN1 and VDREs. These studies will also be important in understanding how MN1 and SRCs functionally cooperate to regulate VDR-mediated transcription. Once the presence of MN1 in the VDR complex is established, further protein-protein interaction studies can be undertaken to elucidate the exact molecular contacts that exist between MN1 and other VDR-associated proteins and to map the domains of MN1 important for these interactions. Analysis of the primary sequence of MN1 does not indicate any LXXLL-containing motifs that are present in other nuclear receptor co-
185 activators (51, 68) indicating that a novel mechanism of interaction with the VDR
complex may govern the coactivator activity of MN1.
Previous studies have shown that other classes of coactivators are required for
1,25(OH)2D3-dependent transcriptional activation by expression of dominant
negative proteins or peptides (77) or through biochemical depletion of select
proteins in vitro (64). Similar approaches should be deployed to determine if
MN1 is also an essential factor in VDR-mediated transactivation. These studies
could be performed though at least three approaches. First, total levels of MN1
expression could be knocked down using RNA interference (RNAi) gene
silencing technology (366). Next, if specific domains of MN1 are identified that
interact with VDR-associated proteins, these protein regions could be exploited in
a dominant-negative manner to disrupt endogenous MN1 function. We could
then test the effects of decreased MN1 expression or disrupted MN1 function on
VDR-mediated transcriptional activation using reporter gene studies. To
examine the effect of disrupting MN1 on endogenous VDR function, a stable
inducible RNAi approach (367) could be utilized to efficiently knockdown MN1
expression in various cells and determine if 1,25(OH)2D3 can still stimulate
expression of target genes such as 24-hydroxylase. Finally, and most
importantly, with the recent development of MN1-null mice (290), 1,25(OH)2D3 gene responses could be tested in primary osteoblastic cells derived from wild- type and MN1-null embryos. Based on the initial findings presented in Chapter II,
I expect that disrupted MN1 expression will impair 1,25(OH)2D3-mediated
186 transcriptional responses, further solidifying a role for this novel transcriptional
coactivator in modulating VDR-regulated gene expression.
Our data indicate that overexpression of MN1 causes osteoblastic cell growth
arrest by delaying entry into the S phase of the cell cycle (Fig. II-6 and Fig. II-7).
It is possible that MN1 is one factor induced by 1,25(OH)2D3 that mediates the
potent growth inhibitory effects of this hormone (Fig. II-6). To test this
hypothesis, cells with decreased levels of MN1 or cells derived from MN1
knockout animals, as described above, could be used to determine if MN1 is
required, at least in part, for the anti-proliferative effects of 1,25(OH)2D3. Since previous studies (269, 270) and ours have demonstrated that MN1 exhibits transcriptional activity, it is likely that MN1 regulates genes that are involved in cell cycle control. Potential target genes include the cyclin-dependent kinase inhibitors p21WAF1/cip1 and p27kip1, two genes already shown to be induced by
1,25(OH)2D3 (261, 262). In fact, MN1 may cooperate with VDR through its
coactivator activity to regulate such target genes. This possibility could be
explored by testing the ability of MN1 to enhance 1,25(OH)2D3-mediated
stimulation of p21WAF1/cip1 and p27kip1 promoter activity. Alternatively, MN1 may
act independently of its coactivator activity to modulate expression of genes
involved in proliferation and cell cycle progression. Such target genes could be
discovered through global gene expression profiling changes in MN1-deficient
cells, using either cells derived from MN-null embryos (290) or RNAi-mediated knockdown of MN1 expression in cell lines.
187
C/EBPβ
Chapter III demonstrates that C/EBPβ is regulated by 1,25(OH)2D3 in osteoblastic cells (Fig. III-1) and functionally interacts with VDR to stimulate expression of 24(OH)ase (Fig. III-2). In fact, we demonstrate, using C/EBPβ-null cells, that C/EBPβ is essential for the maximal activation of the 24(OH)ase
promoter by 1,25(OH)2D3 (Fig. III-2). Further analysis of C/EBPβ-deficient cells
shows that this protein is required for in vitro osteoblastic differentiation and
mineralization (Fig. III-3). Finally, a mouse with a targeted deletion in both
C/EBPβ and VDR showed dramatically undermineralized bones, while single
knockout bones did not appear grossly abnormal (Fig. III-4). These findings
indicate that, in addition to cooperatively enhancing gene expression, C/EBPβ
and VDR may functionally collaborate in vivo to modulate bone mineral density.
Both reporter gene analysis and RT-PCR amplification of endogenous
24(OH)ase levels have established the essential function of C/EBPβ in
1,25(OH)2D3-mediated stimulation of 24(OH)ase expression. However, deletion
of C/EBPβ more prominently diminished the promoter activity of 24(OH)ase as
compared to the 24(OH)ase mRNA levels (Fig. III-2). As such, it is likely that
other mechanisms are operating in the context of the full-length native
24(OH)ase gene to regulate its transcriptional activation. More detailed
molecular analyses are in order to fully elucidate how VDR and C/EBPβ
cooperate. In vitro studies indicate that VDR and C/EBPβ can directly interact
188 (P. Dhawan and S. Christakos, unpublished observations). These data suggest
a model in which VDR and C/EBPβ are tethered to the promoter through their
respective DNA-binding domains and induce a folding of the regulatory region to
interact with one another. The identity of other proteins that associate with VDR
and C/EBPβ in this context will add another layer of complexity to this
transcriptional mechanism.
It will also be important determine if the cooperative activity between VDR and
C/EBPβ is specific to select subset of 1,25(OH)2D3 target genes or if C/EBPβ
represents a general coactivator for VDR in regulating all 1,25(OH)2D3-induced
genes. Of special interest to our laboratory is the potential involvement of
C/EBPβ in regulating expression of some of the genes identified in the current
work. SEMA3B represents the most prominent candidate gene given that
1,25(OH)2D3-mediated induction of SEMA3B is entirely dependent on synthesis of other protein factors (Fig. IV-1). Indeed, preliminary evidence suggests that
C/EBPβ regulates the SEMA3B promoter (see below).
To examine the potential cooperative interplay between VDR and C/EBPβ in
vivo, mice with a targeted deletion in both genes were engineered. μCT analysis revealed that while single VDRKO and C/EBPβ KO bones had no obvious abnormalities, the VDRKO/C/EBPβKO bone had markedly decreased cortical and trabecular bone (Fig. III-4). These studies suggest that VDR and C/EBPβ
may functionally compensate for one another in regulating bone mineral density.
189 Although these findings are exciting, it is important to note that only one animal of
each genotype was analyzed in this preliminary study. Unfortunately, we can
generate only limited numbers of C/EBPβKO animals due to the increased perinatal lethality of the null allele in the C57BL/6 genetic background (C.
Croniger, unpublished observations). In a mixed background of 129/Sv/Ev and
C57Bl/6, approximately 50% of the C/EBPβ-null mice reach adulthood (306). To better investigate the effects of disrupting C/EBPβ and VDR on bone density, we should breed these mice into the mixed background to increase the number of animals in the study. Furthermore, the current work utilized qualitative μCT imaging. A more accurate and detailed assessment of the bone density and volume can be obtained with quantitative pQCT and μCT measurements, as was performed in Chapter IV. Similarly, these sensitive methods may reveal more subtle abnormalities in the single C/EBPβKO bones. These studies are especially important to perform considering our in vitro data that supports a dramatic mineralization defect in C/EBPβKO osteoblasts (Fig. III-3). Finally, given the marked decrease in mineralization and the extreme fragility of the
VDRKO/C/EBPβKO bone, it is likely that these mutant bones are structurally weak and prone to fractures. The strength of these bones could be evaluated by mechanical testing (368). In fact, Dr. Clare Rimnac of the Department of
Mechanical and Aerospace Engineering at Case Western Reserve University has experience with characterizing the biomechanic properties of bone and would represent a useful collaborator for these studies.
190 SEMAPHORIN3B
As discussed in Chapter IV, 1,25(OH)2D3 regulates SEMA3B expression in
osteoblastic cells in vitro (Fig. IV-1). The SEMA3B protein is also expressed in
osteoblasts in vivo, suggesting a potential role for this signaling system in bone
(Fig. IV-2). To further explore the function of SEMA3B in the skeleton, a
transgenic mouse model that selectively overexpresses SEMA3B in osteoblasts
was created (Fig. IV-3). These animals are severely osteopenic potentially due
to enhanced osteoclast formation and bone resorption (Fig. IV-6 – Fig. IV-10).
Together, these data suggest that SEMA3B is a 1,25(OH)2D3-regulated gene that
stimulates osteoclastogenesis and bone resorption.
Both active transcription and de novo protein synthesis are required for
1,25(OH)2D3-mediated induction of SEMA3B mRNA levels (Fig. IV-1C and Fig.
IV-1D). These findings suggest that 1,25(OH)2D3 stimulates expression of other
protein factors that either cooperate with VDR or act alone to stimulate
expression of SEMA3B. Two separate putative promoters, based on the
expressed sequence tag databases, of SEMA3B have been cloned into to
reporter genes, but neither respond to 1,25(OH)2D3 treatment in MG-63 cells (A.
Sutton and X. Zhang, unpublished observations). However, overexpression of
C/EBPβ, but not C/EBPδ, causes a dose-dependent increase in the activity of the proximal promoter activity (X. Zhang, unpublished observations). Since
1,25(OH)2D3 also stimulates C/EBPβ expression (Fig. III-1) and VDR and
C/EBPβ cooperatively regulate 24-hydroxylase promoter activity (Fig. III-2), it is
191 possible that C/EBPβ and VDR act in a similar manner to enhance SEMA3B
expression. Further studies, such as examining the 1,25(OH)2D3-mediated
induction of SEMA3B expression and testing the activity of the SEMA3B
promoters in C/EBPβ-null osteoblasts, are required to establish the potential role
of C/EBPβ in modulating SEMA3B expression.
Along these same lines, although we have clearly established that 1,25(OH)2D3 induces SEMA3B expression in human MG-63 osteoblastic cells, it is unclear whether 1,25(OH)2D3 upregulates SEMA3B mRNA levels in mouse osteoblastic
cells. A separate microarray analysis demonstrated that SEMA3B is potently
regulated by 1,25(OH)2D3 in human squamous cell carcinoma cells (266),
indicating that this regulation is not likely tissue-selective. However, the
regulation may indeed be species-specific. Preliminary RT-PCR data indicated
that 1,25(OH)2D3 may increase murine SEMA3B expression in primary
osteoblasts (A. Sutton, unpublished observations), but these experiments must
be repeated with a more quantitative approach. Furthermore, 1,25(OH)2D3-
mediated regulation of SEMA3B should be analyzed throughout the
differentiation profile of primary osteoblasts as they may be more or less
responsive depending on their maturation level.
Regardless of whether 1,25(OH)2D3 stimulates SEMA3B expression in murine
osteoblasts, our transgenic mouse model suggests that overexpression of this
protein in osteoblasts dramatically impairs bone mineralization potentially through
192 stimulating bone resorption. Histomorphometric analysis shows a trend toward
increased osteoclastogenesis (Fig. IV-8), and these findings are supported by in
vitro co-culture data demonstrating that transgenic osteoblasts support enhanced
osteoclast differentiation (Fig. IV-10). We are currently in the process of
analyzing more animals with histomorphometry to provide statistical significance
to these data. Additionally, since the in vitro osteoclastogenesis assay was only
performed once, this study must be repeated to validate the results. Regardless,
the data presented in this study provide compelling evidence for the first time that
semaphorin signaling regulates bone mass.
Although these findings are intriguing, the obvious caveat exists that, of the two transgenic lines derived, only one expresses abundant levels of the transgene
(Fig IV-3). Since the high-expressing line has a severe osteopenic phenotype, it is likely, although not conclusive, that these effects are due to the increased levels of SEMA3B. To definitively rule out integration site effects, at least one additional transgenic line should be generated and analyzed for bone density defects. In contrast to the in vivo data, osteoblasts derived from both the low-
expressing and high-expressing lines of transgenic mice differentiate and
mineralize at an accelerated rate. These findings suggest that low levels of
SEMA3B expression are sufficient to stimulate osteoblast differentiation. In
contrast, higher levels of SEMA3B may be required to induce osteoclastic
activity. Since the in vitro co-culture assay demonstrating that SEMA3B transgenic osteoblasts support increased osteoclastic activity has only been
193 performed using cells derived from the high-expressing line, it will be important to
repeat these assay using osteoblasts from the low-expressing line. If my
hypothesis holds true then we should not observe enhanced osteoclastogenesis
in co-cultures with low-expressing osteoblasts.
Beyond simple increases in osteoclast number, the histomorphometric analysis
suggests that transgenic mice have an increased surface of bone involved in
resorption (ES/BS; Fig. IV-8). This increase could be explained by the elevated
osteoclast numbers, increased osteoclast activity, or both. To examine
osteoclast activity in vitro, pit-forming assays could be performed. In this assay,
osteoblasts derived from transgenic and non-transgenic mice are co-cultured
with osteoclast precursor cells on ivory slices and resorption lacunae are
visualized by staining (369). The number and size of these “pits” reflect the
activity of the osteoclasts in the culture.
The current work provides in vivo and in vitro evidence that SEMA3B regulates osteoclastogenesis, but the mechanism remains unclear. Most importantly, we must determine whether SEMA3B affects osteoclast differentiation directly or indirectly, by stimulating expression of additional osteoblast-derived factors that enhance osteoclastogenesis. I have attempted to address this issue by preparing conditioned medium from COS-7 cells that were transfected with a
SEMA3B expression vector. As SEMA3B is secreted into the medium, this approach provides a readily available source of soluble SEMA3B. However,
194 conditioned media from neither vector-transfected nor SEMA3B-transfected
COS-7 cells is capable of supporting in vitro osteoclastogenesis (A. Sutton,
unpublished observations). This conditioned medium is also unsuitable for this
assay since the level of SEMA3B secretion is impossible to control. Thus, to
better test if SEMA3B directly augments osteoclast differentiation, a more purified
source of SEMA3B should be utilized. Our laboratory has extensive experience
with the purification and characterization of baculovirus-expressed proteins (370),
so baculovirally-expressed SEMA3B will likely prove useful in future osteoclast
differentiation studies.
A few reports have indicated that semaphorins, neuropilin receptors, and the effector molecule rac1 are expressed in osteoclasts (331, 338, 344), suggesting that direct SEMA3B-mediated stimulation of osteoclastogenesis is possible since the components for a functional semaphoring signaling pathway are present in these cells. Further, inhibition of rac1 function by expression of a dominant negative protein or with neutralizing antibodies disrupts osteoclast differentiation and bone resorptive activity (343, 344), highlighting an essential role for this small GTPase in osteoclastogenesis. In light of these data, it is possible that
SEMA3B signals through rac1 to promote osteoclast differentiation. However, rac1 responds to multiple signaling inputs, so we must first test if rac1 is activated by SEMA3B. Purified SEMA3B, as indicated above, can be applied to osteoclasts and tested for its ability to activate rac1. The Wilson-Delfosse laboratory currently uses an approach that sensitively detects the activated GTP-
195 bound form of rac1 and other small G-proteins in response to various stimuli.
This technique could be utilized to determine if endocgenous rac1 is activated by
SEMA3B in osteoclasts. The ultimate outcome of rac1 signaling is often reorganization of the actin cytoskeleton, which has been observed in osteoclasts
(344), or gene expression changes. We could utilize phalloidin staining to monitor the cytoskeleton for structural changes following SEMA3B treatment in
osteoclasts. Likewise, as NFATc1 is considered the master transcriptional
regulator in osteoclast differentiation (209), the activity of this protein in response
to SEMA3B could be tested with a reporter gene assay that is currently in use in
our laboratory. Such studies will provide exciting mechanistic insight into the
semaphorin signaling pathway in osteoclasts and further solidify the functional
link between SEMA3B and osteoclastogenesis.
THROMBOMODULIN
Chapter V investigates the regulation and function of the anticoagulant protein
thrombomodulin in osteoblasts and in bone. 1,25(OH)2D3 appears to directly
regulate thrombomodulin expression at both the mRNA level (Fig. V-1 and Fig.
V-2) and protein level (Fig. V-3). To understand the function of thrombomodulin
in osteoblasts, cell lines were created that stably overexpress thrombomodulin
(Fig. V-4). These lines displayed an increased proliferation rate when compared
to vector-transfected lines (Fig. V-4). Finally, a mouse strain with an osteoblast- selective knockout of thrombomodulin was created to examine the potential in vivo effects of thrombomodulin in bone. However, preliminary μCT analysis of
196 tibiae from these animals did not reveal any striking defects, indicating that deletion of thrombomodulin in osteoblasts does not have a drastic effect on bone
morphology or density.
Although our study and numerous others (350, 355-357) have firmly established
that 1,25(OH)2D3 increases thrombomodulin expression in multiple cells types,
the molecular mechanisms involved in this response are not well-characterized.
A retinoid-responsive element (RARE) in the thrombomodulin promoter is weakly stimulated by 1,25(OH)2D3 in myelogenous leukemia cells (356). However, we
have shown that a 3.5 kb fragment of the thrombomodulin promoter that contains this RARE is not affected by 1,25(OH)2D3 in MG-63 cells (A. Sutton, unpublished
observations). Therefore, the VDRE may lie elsewhere in the promoter, in the
exonic sequences, or in the 3’ untranslated region of thrombomodulin. Clearly
further promoter mapping studies are required to understand how 1,25(OH)2D3 regulates thrombomodulin expression.
The expression and up-regulation of thrombomodulin by 1,25(OH)2D3 in
osteoblasts is somewhat surprising since this protein classically acts as
anticoagulant protein in the vasculature. The potential effects of thrombomodulin
in osteoblasts and the bone remain completely unexplored. Therefore, we
attempted to uncover the functional implications of thrombomodulin expression in osteoblasts by generating stable MG-63 cells lines that overexpress this protein.
These lines showed an increased rate of proliferation (Fig. V-4). This finding was
197 unexpected given the anti-proliferative effects of 1,25(OH)2D3 in osteoblastic cells
(Fig. II-6). Further analyses of the cell lines have shown that thrombomodulin overexpression does not affect osteoblastic differentiation as measured by alkaline phosphatase activity (A. Sutton, unpublished observations). Thus, this in vitro system may not prove useful in uncovering the true function of thrombomodulin in osteoblasts. In this study we also report establishment of a mouse model with an osteoblast-selective disruption of thrombomodulin to probe the in vivo functions of thrombomodulin in osteoblasts and in the bone.
Preliminary μCT imaging of these osteoblast-selective thrombomodulin knockout mice did not reveal any striking abnormalities in bone morphology or density.
However, since this initial assessment was qualitative rather quantitative, a detailed measurement of bone mineral density and bone volume by pQCT and quantitative μCT, as was performed in Chapter IV, is required to more accurately determine if osteoblastic deletion of thrombomodulin affects skeletal mineralization.
Even if these further studies do not support a function for thrombomodulin in
regulating osteoblast differentiation and bone density, it remains possible that osteoblast-expressed thrombomodulin may modulate other processes in the skeleton. Recently, several reports have uncovered an important role for osteoblasts in creating a microenvironment conducive to proper hematopoiesis
(371-373). These results are not surprising considering that osteoblasts line the surface of the marrow cavity and interface with hematopoietic stem cells.
198 Osteopontin, a bone matrix protein, is highly expressed by osteoblasts on the
surface of bony trabeculae that intercalate with the bone marrow (374). Targeted
inactivation of osteopontin in mice indicates that this protein functions in
negatively regulating hematopoietic stem cell proliferation (375, 376).
Osteopontin is cleaved by thrombin (377, 378) to generate fragments that bind to and signal through integrin receptors (379). It is possible that local bone thrombin, by cleaving osteopontin, may also inhibit hematopoietic stem cell proliferation. Furthermore, since thrombomodulin decreases thrombin activity, osteoblast-derived thrombomodulin may be involved in the hematopoietic microenviroment by indirectly promoting stem cell proliferation. Clearly, many more studies are required to test these hypotheses. The osteoblast-selective thrombomodulin knockout mouse developed in this work will provide an important tool for studying the potential contributions of thrombomodulin to the hematopoietic stem cell niche.
CONCLUSIONS
The studies presented in this dissertation have identified and characterized four
novel 1,25(OH)2D3-induced genes in osteoblastic cells: MN1, C/EBPβ, SEMA3B,
and thrombomodulin. First, our findings suggest that MN1 may participate in a
feed-forward mechanism whereby 1,25(OH)2D3 directly stimulates MN1
expression and, MN1, in turn promotes VDR-dependent transcription, eventually
inhibiting osteoblastic cell proliferation. Second, we demonstrate that C/EBPβ
and VDR cooperatively regulate both 24(OH)ase expression on a molecular level
199 and bone mineralization on an organismal level, and show that C/EBPβ is
required for osteoblastic cell differentiation in vitro. Third, our data uncover a
novel role for SEMA3B in negatively regulating bone mass through stimulating
osteoclastogenesis and bone resorption. Finally, we show that thrombomodulin may promote osteoblast proliferation, and we report development of an osteoblast-selective knockout model of thrombomodulin to further study the potentially novel function of this gene in the bone. As is the case with many global gene expression profiling studies, this work has likely posed more questions than it has answered. Nonetheless, the findings have provided significant insight into the regulated expression and function of MN1, C/EBPβ,
SEMA3B, and thrombomodulin in osteoblastic cells and within the more complex
environment of bone tissue.
200
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