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This is to certify that the
dissertation entitled

PATHOGENESIS AND TREATMENT OF TYPE 1 DIABETIC
OSTEOPOROSIS '

presented by

KATHERINE JEAN MOTYL

has been accepted towards fulfillment
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PATHOGENESIS AND TREATMENT OF TYPE 1 DIABETIC OSTEOPOROSIS
By

Katherine Jean Motyl

A DISSERTATION

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

DOCTOR OF PHILOSOPHY
Physiology

2010

ABSTRACT
PATHOGENESIS AND TREATMENT OF TYPE 1 DIABETIC OSTEOPOROSIS
BY
Katherine Jean Motyl
Type 1 diabetes (T1-diabetes) is a chronic condition characterized by

hypoinsulinemia and hyperglycemia. Patients with T1-diabetes are susceptible to
complications, including osteoporosis. Bone loss from diabetes increases risk of
fracture and impairs fracture healing. Understanding the mechanism of diabetic
osteoporosis is important for choosing the best therapies. Reduced bone
formation (not altered resorption) is responsible for diabetic bone loss.
Additionally, rodent models of T1-diabetes (streptozotocin and the non-obese
diabetic mouse) have increased marrow adipocyte number compared normal,
suggesting mesenchymal stem cell lineage selection favors the osteoblast over
the adipocyte in diabetic conditions. In Chapter 2, we characterized the bone
phenotype of mice during diabetes induction and found that markers of
osteoblasts and adipocytes were altered as early as two days after blood glucose
levels began to rise. Because diabetes is associated with inflammation, we
measured serum and bone pro-inflammatory cytokine levels and found increases
in cytokines that coincided with reduced osteoblast activity. Diabetes is also.
associated with reduced serum leptin, a hormone secreted by adipocytes that is
capable of promoting bone formation in conditions of unloading. In Chapter 3, we
treated diabetic mice with leptin and found that chronic subcutaneous leptin

infusion prevented diabetic marrow adiposity but not bone loss, suggesting that

marrow adiposity is not required for diabetic osteoporosis. In Chapter 4, we
induced diabetes in mice lacking CCAAT/enhancer binding protein beta
(C/EBPB), a transcription factor that promotes adiposity and prevents osteoblast
differentiation. We found that under diabetic conditions, the lack of C/EBPB
actually enhanced adipocyte differentiation in bone and increased bone
resorption, which is not classically observed in T1 -diabetes. In Chapter 5, we
found reduced Wnt1 Ob, a strong activator of osteoblast differentiation and
inhibitor of adipogenesis, in diabetic bone. To test whether Wnt10b was a factor
influencing bone changes in diabetes, we induced T1 -diabetes in mice with
overexpression of Wnt1 Ob in osteoblasts. Wnt10b mice were protected from
bone loss but not marrow adiposity in diabetes. Finally, in Chapter 6 we
examined the ability of intermittent parathyroid hormone (PTH), the only anabolic
osteoporosis therapy available to patients, to promote bone formation in
diabetes. We found that PTH increased osteoblast maturation and prevented
osteoblast death, even under diabetic conditions. Bone density levels of diabetic
mice returned to those of untreated controls, even after bone loss had already
occurred. Taken together, Chapters 2-5 demonstrate that bone marrow fat is
likely not necessary for diabetic bone loss. Additionally, Chapter 5 and 6 suggest
that anabolic therapies targeting Wnt10b/wnt signaling could improve bone

density in diabetic patients.

Dedicated to my husband Chris and our beautiful son Henry for their
unconditional love and encouragement through the good times and the better

times. I love you both.

ACKNOWLEDGMENTS

Dr. Laura McCabe is deserving of a very large portion of this section. I still
remember the day that I met her in her office and begged her to let me rotate in
the lab. She did, of course (she has trouble saying “no”) and the rest is history. It
has been a thrill to be a part of the research that she is so passionate about. She
has always been approachable and grounded, and a source of wisdom and
encouragement for life and lab. Thank you Laura.

My committee members, Dr. Ron Meyer, Dr. Narayannan Parameswaran,
Dr. Gloria Perez, and Dr. Richard Schwartz have given me useful input and
guidance over the past five years. Thank you for all of the time you have spent
helping me to achieve this goal.

Regina Irwin and Lindsay Martin, my current lab mates, have always given
me somethingto laugh about. Thank you forthe time you have given and
continue to give to this research. Also, thank you to the other graduate students,
undergraduate students, volunteers and rotation students who have donated
their time to these worthy projects. Thank you to the Department of Physiology
for their support.

Thank you to my family Chris and Henry for keeping me smiling through
this entire process. Thank you to my parents, Ken and Del Frank and to my in-
Iaws Dave and Michelle Motyl for encouraging me to do my best and for

supporting me in all of the decisions that l have made.

TABLE OF CONTENTS

LIST OF TABLES .................................................................................................. ix
LIST OF FIGURES ................................................................................................. x
LIST OF ABBREVIATIONS .................................................................................. xii
CHAPTER 1. LITERATURE REVIEW ........................................................ 1
1.1. BONE ................................................................................................... 1
1.1.1. Molecular composition of bone ............................................... 1
1 1 2. Cellular composition of bone .................................................. 2
1.1.3. Bone development and anatomy ............................................ 5
1.1.4. Regulation of osteoblast differentiation .................................. 7
1 1 5. Adipocyte differentiation ......................................................... 9
1 1 6. Osteoclasts ............................................................................. 9
1.1.7. Mineral homeostasis ............................................................ 12
1.2. OSTEOPOROSIS .............................................................................. 13
1 3 GLUCOSE HOMEOSTASIS AND DIABETES MELLITUS ................ 14
1.3.1. Glucose homeostasis ........................................................... 14
1.3.2. Diabetes mellitus definition and classifications .................... 14
1.3.3. Complications of diabetes mellitus ....................................... 16
1.3.4. Animal models of T1-diabetes and T2-diabetes ................... 17
1.4. BONE PHENOTYPE IN DIABETES MELLITUS ................................ 19
1.4.1. Type 2 diabetic bone phenotype .......................................... 19
1.4.2. Type 1 diabetic osteoporosis ................................................ 20
1.4.3. Mechanisms of type 1 diabetic bone loss ............................. 21
1.4.4. Treatment of T1 -diabetic bone loss ...................................... 30
1.5. SUMMARY ......................................................................................... 32
1.6. REFERENCES ................................................................................... 33
CHAPTER 2. BONE INFLAMMATION AND ALTERED GENE EXPRESSION
WITH TYPE I DIABETES EARLY ONSET ................................................. 47
2.1. ABSTRACT ........................................................................................ 48
2.2. INTRODUCTION ................................................................................ 49
2.3. MATERIALS AND METHODS ........................................................... 52
2.3.1. Streptozotocin mouse injections ........................................... 52
2.3.2. Genotyping ........................................................................... 52
2.3.3. Plasma measurements ......................................................... 53
2.3.4. RNA Analysis ....................................................................... 54
2.3.5. Micro-computed tomography (uCT) analysis ....................... 56
2.3.6. Statistical analysis ................................................................ 57
2.4. RESULTS ........... _. ............................................................................... 58
2.5. DISCUSSION ..................................................................................... 78

vi

2.6. ACKNOWLEDGEMENTS .................................................................. 84

2.7. REFERENCES ................................................................................... 86
CHAPTER 3. LEPTIN TREATMENT PREVENTS TYPE I DIABETIC MARROW
ADIPOSITY BUT NOT BONE LOSS IN MICE ..................................................... 98

3.1. ABSTRACT ........................................................................................ 99

3.2. INTRODUCTION .............................................................................. 100

3.3. MATERIALS AND METHODS ......................................................... 102

3.3.1. Animals ............................................................................... 102
3.3.2. Serum measurements ........................................................ 103
3.3.3. Bone Histology and Histomorphometry .............................. 104
3.3.4. Micro Computed Tomography (uCT) Analyses .................. 104
3.3.5. RNA Analyses .................................................................... 105
3.3.6. Statistical Analyses ............................................................ 106
3.4. RESULTS ......................................................................................... 107
3.4.1. Serum leptin, glucose and insulin ....................................... 107
3.4.2. Food consumption and body mass .................................... 110
3.4.3. Diabetic marrow adiposity was prevented by leptin
treatment.........; ............................................................................ 113
3.4.4. Leptin did not prevent diabetic bone loss ........................... 116
3.4.5. Leptin treatment suppressed bone resorption in diabetes.124

3.5. DISCUSSION ................................................................................... 127

3.6. ACKNOWLEGEMENTS ................................................................... 134

3.7. REFERENCES ................................................................................. 135

CHAPTER 4. CCAAT/ENHANCER BINDING PROTEIN BETA-DEFICIENCY
ENHANCES TYPE 1 DIABETIC BONE PHENOTYPE BY INCREASING

MARROW ADIPOSITY AND BONE RESORPTION ...... ’ ................................... 142
4.1. ABSTRACT ...................................................................................... 142
4.2. INTRODUCTION .............................................................................. 143
4.3. MATERIALS AND METHODS ......................................................... 146

4.3.1. Animals ............................................................................... 146
4.3.2. RNA Analyses .................................................................... 147
4.3.3. Bone histology and histomorphometry ............................... 148
4.3.4. Micro Computed Tomography (uCT) Analyses .................. 149
4.3.5. Statistical Analyses ............................................................ 150
4.4. RESULTS ......................................................................................... 150
4.5. DISCUSSION ................................................................................... 169
4.6. ACKNOWLEDGEMENTS ................................................................ 174
4.7. REFERENCES ................................................................................. 175

CHAPTER 5. MICE WITH OVEREXPRESSION OF WNT1OB ARE RESISTANT
TO BONE LOSS BUT NOT MARROW ADIPOSITY FROM TYPE 1

DIABETES ......................................................................................................... 182
5.1 . ABSTRACT ...................................................................................... 182
5.2. INTRODUCTION .............................................................................. 183

vii

5.3. MATERIALS AND METHODS ......................................................... 187

5.3.1. Leptin and PTH treatment .................................................. 188
5.3.2. OC-Wnt10b mice and diabetes induction ........................... 188
5.3.3. Micro-computed tomography (uCT) analyses .................... 189
5.3.4. RNA analyses ..................................................................... 190
5.3.5. Serum measurements ........................................................ 192
5.3.6. Bone histology and histomorphometry ............................... 192
5.3.7. Statistical Analyses ............................................................ 193
5.4. RESULTS ......................................................................................... 193
5.5. DISCUSSION ................................................................................... 209
5.6. ACKNOWLEDGEMENTS ................................................................ 212
5.7. REFERENCES ................................................................................. 214

CHAPTER 6. OSTEOPOROSIS FROM TYPE 1 DIABETES IS REVERSED BY
INTERMITI'ENT PARATHYROID HORMONE STIMULATION OF BONE

REMODELING AND REDUCTION OF OSTEOBLAST APOPTOSIS ............... 222
6.1 . ABSTRACT ...................................................................................... 222
6.2. INTRODUCTION .............................................................................. 223
6.3. MATERIALS AND METHODS ......................................................... 227

6.3.1. Diabetes induction .............................................................. 227
6.3.2. PTH Treatment ................................................................... 228
6.3.3. Micro-computed tomography (uCT) analyses .................... 228
6.3.4. Bone histology and histomorphometry ............................... 229
6.3.5. RNA analyses ..................................................................... 231
6.3.6. Statistical Analyses ............................................................ 232
6.4. RESULTS ......................................................................................... 232
6.4.1. Diabetes induction and body composition .......................... 232
6.4.2. PTH Counteracted Diabetic Bone Loss .............................. 235
6.4.3. PTH increased bone formation in a dose-dependent
manner ......................................................................................... 240
6.4.4. Effect of PTH on Bone Resorption ..................................... 244
6.4.5. Osteoblast viability is improved by PTH treatment ............. 246
6.4.6. Diabetic bone loss can be reversed by PTH ...................... 250
6.5. DISCUSSION ................................................................................... 254
6.5.1. Both low and high dose PTH treats trabecular bone loss from
diabetes ........................................................................................ 254
6.5.2. PTH restores bone density after it has occurred ................ 256
6.5.3. Diabetic response to PTH is stronger ................................. 257
6.5.4. Reduction of osteoblast death: basal and diabetes
induced ......................................................................................... 258
6.5.5. Cortical bone effects ........................................................... 259
6.6. SUMMARY ....................................................................................... 260
6.7. ACKNOWLEDGEMENTS ................................................................ 260
6.8. REFERENCES ................................................................................. 262

viii

LIST OF TABLES

Table 1. Serum and tissue mass measurements ............................................... 109
Table ‘2. Trabecular uCT measurements ............................................................ 118
Table 3. Tibia cortical uCT measurements ......................................................... 121

Table 4. [Body and tissue masses of 28 day diabetic and untreated wild type and
C/EBPB mice .................................................................................................. 155
Table 5. Trabecular bone uCT measurements from the tibia of 28 day diabetic

4.
and untreated wild type and C/EBPB mice ...................................................... 162

Table 6. Femur trabecular uCT measurements in control and diabetic, wild type
and OC-Wnt10b mice ........................................................................................ 203

Table 7. Blood glucose, muscle and fat composition of control and diabetic,
vehicle and PTH treated mice at 40 dpi ............................................................. 234

Table 8. ”CT analysis of control and diabetic, vehicle and PTH treated mice at 40
dpi ...................................................................................................................... 239

LIST OF FIGURES

Figure 1. Bone cells .............................................................................. 4

Figure 2. Three-dimensional micro-computed tomography image of a
mouse tibiofibula ................................................................................. 6

Figure 3. Differentiation of mesenchymal stem cells to osteoblasts and
adipocytes .......................................................................................... 8

Figure 4. Osteoclast activation through the RANKLIOPG pathway ............. 11
Figure 5. B-catenin signaling in the presence and absence of wnt Iigands..29

Figure 6. Time course of general body parameters during the onset of type I

diabetes in BALBIc mice ................................................................................... 61
Figure 7. Significant bone phenotype changes are evident at 5 dpi ............ 64
Figure 8. Analyses of osteoclast regulators, markers and activity .............. 67
Figure 9. Serum cytokine levels increase during the early onset of

diabetes .............................................................................................................. 70
Figure 10. IL-1 Ra, LT-B, TNF-a and IFN-y mRNA levels are increased in
diabetic bone ...................................................................................................... 72
Figure 11. Cytokine mRNA levels are increased predominantly during the
early stage (5 dpi) of the onset of diabetes ..................................................... 74
Figure 12. IFN-y deficiency does not prevent diabetic bone loss .................. 77

Figure 13. Leptin treatment prevented diabetic hyperphagia compared to

vehicle-treated mice, but did not prevent weight loss ................................. 112
Figure 14. Leptin treatment prevented T1-diabetic marrow adiposity ......... 115
Figure 15. Leptin did not prevent T1-diabetic bone loss .............................. 120
Figure 16. Cortical bone thickness did not differ when corrected for body

mass changes .................................................................................................. 123
Figure 17. Leptin treatment suppressed bone resorption diabetes ............ 126

Figure 18. CIEBPB expression is increased in diabetic bone in conjunction
with aP2 expression and is followed by increased marrow adiposity ........ 152

Figure 19. Diabetes induction did not differ between CIEBPB knockout and
wild type mice .................................................................................................. 154

Figure 20. Diabetic marrow adiposity was increased by CIEBPB
deficiency ......................................................................................................... 158

Figure 21. Adipogenic transcription factors are elevated in diabetic CIEBPB'
" compared to wild type diabetic bone .......................................................... 160

Figure 22. Absence of CIEBPB exacerbated T1 -diabetic bone loss without
altering osteocalcin expression ..................................................................... 164

Figure 23. CIEBPB knockout causes increased bone resorption in type 1
diabetes ............................................................................................................ 168

Figure 24. Wnt10b mimics bone volume fraction changes in diabetes and in
PTH and leptin treatments .............................................................................. 196

Figure 25. Overexpression of Wnt10b did not prevent diabetes induction by
streptozotocin .................................................................................................. 197

Figure 26. Mice with Wnt10b overexpression were protected from diabetes-
induced trabecular bone loss ......................................................................... 200

Figure 27. Diabetes reduces bone formation in both wild type and OC-
Wnt10b mice, but does not alter resorption .................................................. 207

Figure 28. Overexpression of Wnt10b did not alleviate diabetic marrow

adiposity ........................................................................................................... 208
Figure 29. PTH treatment counteracted trabecular bone loss from T1-

diabetes ............................................................................................................ 238
Figure 30. PTH promotes bone formation in diabetes .................................. 243
Figure 31. High dose PTH promotes bone resorption in diabetic mice ...... 245
Figure 32. PTH ameliorates diabetes-induced osteoblast death ................. 249

Figure 33. PTH promotes bone formation in diabetic mice when initiated
after diabetic bone loss is detectable ............................................................ 253

xi

AP-1
AGE
ATP
BADGE
BMC
BMD
BMI
BMU
BVF
bZIP
bea1
C/EBP
CRP
CSF1 R

Dlx
DNA
dpi
FABP4
Fra-1
de
GDM
GLUT
GSK38

Hb A1c
HD
IFNJY
InsZ
IRKO
KO
LAP
LEF
LIP
LRP
M-CSF
MSC
Msx
NF-KB
OPG
PICP
PPARy
Pref-1
PTH

LIST OF ABBREVIATIONS

activator protein 1

advanced glycation end product
adenosine triphosphate
bisphenol-A—diglycidyl ether
bone mineral content

bone mineral density

body mass index

basic multicellular unit

bone volume fraction

basic region-Ieucine zipper
core binding factor 1

CCAAT enhancer binding protein
C-reactive protein

colony stimulating factor 1 receptor
dickkopf

distaless

deoxyribonucleic acid

days post injection

fatty acid binding protein 4
Fos-related antigen 1

frizzled

gestational diabetes mellitus

' glucose transporter

glycogen synthase kinase 3 beta

glycated hemoglobin

homeodomain

interferon gamma

insulin 2

insulin receptor knockout

knockout

liver activating protein

lymphoid enhancer factor

liver inhibitory protein

low density Iipoprotein receptor-related protein
macrophage colony stimulating factor
mesenchymal stem cell

meshless

nuclear factor kappa-B

osteoprotegrin

procollagen carboxy-terminal extension peptide
peroxisome proliferator-activated receptor gamma
preadipocyte factor 1

parathyroid hormone

xii

RANK

RAN KL
ROS

Runx2

Ser

sFRP
SREBP
STZ
T1-diabetes
T2-diabetes
TCF

TNFa
TRAP

Thr

WT

receptor activator of nuclear factor kappa-B
receptor activator of nuclear factor kappa-B ligand
reactive oxygen species

runt related transcription factor 2

seflne

secreted frizzled-related protein

sterol regulator element binding protein
streptozotocin

type 1 diabetes mellitus

type 2 diabetes mellitus

T-cell factor

tumor necrosis factor alpha
tartrate-resistant acid phosphatase
threonine

wildtype

xiii

CHAPTER 1

1. LITERATURE REVIEW

1.1. BONE

Bone is the structural connective tissue comprising the skeleton of
reptiles, mammals, amphibians and some fish (i.e. higher vertebrates) (1). Highly
dynamic, bone is constantly remodeling in response to physiological stimuli.
Remodeling (the process of bone formation and resorption) ensures that bone
fulfills its basic functions of structural support and calcium homeostasis. Defects
in remodeling lead to either too much or too little bone, both of which can have

debilitating consequences.

1.1.1. Molecular composition of bone

Bone is made of a mineralized matrix of collagen proteins containing 99%
of the calcium, 85% of the phosphate, and 50% of the magnesium in the body
(2). Outside of bone, calcium is important for a variety of cellular functions
including muscle contraction and neurotransmitter release and its levels are
therefore tightly controlled (2). The major protein component of the bone matrix is
type 1 collagen, with smaller contributions from other collagens. Non-collagenous

proteins in the bone matrix include osteocalcin, which is secreted by osteoblasts

(bone forming cells), binds calcium, and is clinically used as a blood marker of
osteoblast activity (2). Other proteins in the matrix are important for cell

attachment, collagen fibril formation, and sensing of mechanical stress (2).
1.1.2. Cellular composition of bone

Embedded in the bone matrix are osteocytes, and on the surface of bone
are bone lining cells, osteoblasts, and osteoclasts (Figure 1). Osteoblasts, bone
lining cells, and osteocytes are derived from mesenchymal stem cells.
Osteoblasts are responsible for the deposition of bone matrix, which is
subsequently mineralized either by osteoblast derived matrix vesicles, or through
processes initiated by collagen molecules themselves, although these processes
of mineralization are poorly understood (1). Osteocytes are mature osteoblasts
that occupy circular structures called lacunae and are thought to be responsible
for some maintenance of the bone matrix (1). Bone lining cells are flat and cover
the surface of the bone but do not form or resorb bone (1). Multinucleated
osteoclasts are derived from hematopoietic precursors and are responsible for
the resorption of mineralized bone (1). Osteoclasts, nearby osteoblasts that
deposit osteoid and mineral in their wake and the more distal bone lining cells
together comprise one basic multicellular unit (BMU).

In addition to the above mentioned bone cells, the interior of the bone
compartment contains bone marrow, which is comprised of cells of the

hematopoietic and mesenchymal lineages, and is the site of the majority of

hematopoiesis (blood production). Therefore, bone also contains blood and
lymphatic vessels, leukocytes and erythrocytes, as well as nerves and

adipocytes.

activated
osteoclast

 

 

Figure 1. Bone cells. Cuboidal osteoblasts (bone forming), elongate bone lining
cells, and multinucleated osteoclasts (bone resorbing) line the surface of the
bone. Osteocytes arise from osteoblasts that have become embedded in the
bone.

1.1.3. Bone development and anatomy

Bones have an outer shell of cortical (dense) bone and inner trabecular
(spongy, cancellous) bone in the compartment containing bone marrow (Figure
2). The periosteum and endosteum refer to the layer of cells lining the outer and
inner bone surfaces, respectively. During embryonic development, bone
formation is either endochondral or intramembranous. Vertebrae and long bones
(i.e. femur and tibia) arise from endochondral bone formation while flat bones
(i.e. calvaria and mandible) arise from intramembranous bone formation. During
endochondral bone formation, chondrocytes (derived from mesenchymal stem
cells) form a cartilaginous matrix, which is followed by vascularization and
osteoblast matrix deposition and mineralization (2). Bone lengthening occurs
through continued endochondral bone formation at the site of the cartilaginous
growth plate, also known as the metaphysis. This growth plate becomes
mineralized in young adults when growth halts. Unlike in humans, the growth
plates of mice never completely mineralize and bone growth continues into
adulthood, albeit at a much slower rate. The diaphysis is the region of bone
between both metaphyses, and the epiphyses are the ends of the bone.
Intramembranous bone formation occurs when osteoblasts do not follow a
cartilage matrix template and directly deposit and mineralize the bone matrix.
Long bones can also have intramembranous bone formation at sites where
cartilage does not provide a matrix template (i.e. in the periosteum and during

remodeling in the trabeculi of adults) (2).

proximal

 

growth
‘ {— plate _>

trabecular
bone

marrow
compartment
tibiofibular
junctions

distal

 
  
 

   
 

3i- epiphysis

F metaphysis

- diaphysis

 

 

Figure 2. Three-dimensional micro-computed tomography image of a
mouse tibiofibula. Left: complete bone, right: longitudinal section demonstrating

structures of the interior of the bone.

1.1.4. Regulation of osteoblast differentiation

Differentiation of MSCs to the osteoblast lineage is completely dependent
on expression of runt related transcription factor 2 (Runx2) (also known as
bea1, core binding factor 1) (1) which is induced by Indian hedgehog (lhh).
secretion from prehypertrophic chondrocytes (3, 4) (Figure 3). Runx2 binding
sites are present in the promoters of several genes expressed in mature
osteoblasts, such as osteocalcin and type I collagen (3). Activator protein 1 (AP-
1) family transcription factor (i.e. c-Fos, Fos-related antigen 1 (Fra-1), and c-Jun)
binding sites also exist on the promoters for osteocalcin, alkaline phosphatase
and other bone genes (1, 5). The meshless (Msx) and distaless (Dlx)
homeodomain (HD) proteins regulate skeletal development as well, and some
(i.e. Dlx5) may work independently of Runx2 (6, 7). Activation of osteoblast
differentiation may occur at the expense of other cell types that arise from the
same precursors, such as adipocytes. Other transcriptional regulators (such as
CCAAT enhancer binding protein beta (C/EBPB) and T-cell factor
(T CF)IIymphoid enhancer factor (LEF)) are likely crucial for understanding MSC
fate and will be further discussed in the context of diabetic bone defects (See

Section 1.4.3).

type 1 collagen
osteocalcin

     
 
 

L..Ill.l‘r . .

Runx

        

, [pres . ' "

   

/ iibsteoblai‘Sti -.
marrow
stromal
cell
cram
CIEBPo
pre-
adipocyte —'> adipocyte
CIEBPa.
PPARyZ
Pref-1

aP2

Figure 3. Differentiation of mesenchymal stem cells to osteoblasts and
adipocytes.

1.1.5. Adipocyte differentiation

Adipocytes are also derived from mesenchymal stem cells, and are
present to varying degrees in the bone marrow (depending on location, age, and
other factors). The adipocyte portion of the bone marrow is often called yellow
marrow, while the blood component is red marrow. C/EBPB and C/EBP8 are
transiently expressed during early adipocyte differentiation followed by
expression of peroxisome proliferator-activated receptor gamma (PPARy) 2 and
C/EBPa (8-10) (Figure 3). Preadipocyte factor 1 (Pref-1) is not expressed in
mature adipocytes and is therefore used as a marker for preadipocytes (11).
Alternately, fatty acid binding protein 4 (FABP4, also called aP2) is a marker for
mature adipocytes. Sterol regulatory element binding protein (SREBP) also plays
a role in adipocyte differentiation and regulates several genes important for lipid

metabolism (12).

1.1 .6. Osteoclasts

Osteoclasts, responsible for bone resorption, attach to the bone surface
through a ruffled border surrounded by a sealing zone (Figure 1). The membrane
that is in contact with bone secretes acid and enzymes responsible for degrading
mineral and matrix. Recruitment of an inactive osteoclast precursor requires both
macrophage colony stimulating factor (M-CSF) and receptor activator of nuclear
factor kappa-B ligand (RANKL) (1). After stimulated to differentiate by M-CSF

(through the colony stimulating factor 1 receptor, CSF1 R), osteoclastic

precursors express the membrane bound receptor activator of nuclear factor
kappa-B (RANK), which binds RAN KL to further stimulate osteoclast maturation
and activity. RANKL is expressed on the cell membrane of osteoblasts, which
also secrete osteoprotegrin (OPG), a decoy receptor for RAN KL that inhibits it’s
binding to RANK and thus inhibits osteoclast maturation. Thus, the ratio of
RANKL to OPG is important for regulating degree of bone resorption (Figure 4)
(1). Active osteoclasts secrete tartrate-resistant acid phosphatase (TRAP) 5b into
the serum, which can be quantitated and used to measure whole body bone

resorption status.

10

hematopoietic
- stem cell

osteoclast
progenitor

    
 

RANK

activated
osteoclast

OPG

Figure 4. Osteoclast activation through the RANKIJOPG pathway. M-CSF
embedded in the matrix or on osteoblasts stimulates hematopoetic precursors to
express RANK, becoming osteoclast progenitors. RANKL and M-CSF then
activate osteoclasts. OPG is the decoy receptor for RANKL.

1.1.7. Mineral homeostasis

Whole body calcium and phosphate levels are tightly regulated. Too much
could cause spontaneous precipitation in the tissues and too little could result in
multiple organ dysfunction (2). In its active form, parathyroid hormone (PTH) is
an 84-amino acid peptide secreted from chief cells of the parathyroid gland in
response to low blood calcium. PTH has an extremely short half-life of 2 minutes
once it is secreted, allowing for nearly instantaneous regulation of calcium (2).
PTH itself is stable for longer in solution, but in the body it is quickly metabolized
by the liver and kidneys (2). Although most of the calcium entering the kidney is
resorbed, PTH further stimulates active calcium resorption in the distal

convoluted tubule and the connecting tubule by increasing expression of calcium

transporters, as well as through increased synthesis of vitamin D (1 ,25(OH)2D)

(2, 13). Vitamin D itself promotes expression of calcium transporters in the kidney
and is also necessary for intestinal calcium absorption (2).

PTH effects on bone are dependent on the method of administration and
are not completely understood (2). Minute-to-minute regulation of calcium levels
by PTH are thought to be primarily through its effects on osteocytes, which may
be responsible for small amounts of mineral resorption (1). However,
preosteoblasts, osteoblasts and bone lining cells also express PTH receptors
and exhibit responses to PTH depending on conditions. lnterrnittent PTH (daily
injections) promotes osteoblast differentiation and inhibits osteoblast and

osteocyte apoptosis, perhaps through upregulation of Runx2 (14). Additionally, in

12

vitro PTH treatment activates TCF/LEF-dependent transcription, which is
activated by Wnt signaling (15). A very recent report indicates that the anabolic
effects of intermittent PTH are dependent on T-lymphocyte expression of Wnt10b
(16). PTH may also inhibit production of sclerostin, which inhibits bone formation
by inhibiting Wnt and bone morphogenic protein (BMP) signaling (17). Because
no PTH receptors are present on cells of the osteoclast lineage, net bone
resorption from chronic PTH treatment is likely due to secreted osteoblast factors

(RANKL and M-CSF) activating osteoclast activity (1).

1.2. OSTEOPOROSIS

Osteoporosis is a severe reduction of bone mass, which leaves patients at
risk for fractures. Bone remodeling is altered in osteoporosis such that there is
decreased bone formation, increased resorption, or both. Clinically, osteoporosis
is defined as having a bone mineral density (BMD) less than 2.5 standard
deviations lower than the average population BMD for that location. This number
is also referred to as the T-score. Patients are deemed osteopenic when their T-
score is between -1 and -2.5, and normal if it is greater than -1 (1). Osteoporotic
fractures are a serious health concern because they cost more than $18 billion
annually in the United States and often leave patients hospitalized, with '
decreased mobility, and at higher risk for contracting secondary infections like
pneumonia (18, 19). Therefore, effective diagnosis and treatment of osteoporosis

is essential to the overall health of the patient. However, osteoporosis can occur

13

through different mechanisms depending on the cause (which can range from
aging, menopause and unloading, to being secondary to diseases like diabetes
and inflammatory bowel disease). Understanding the mechanism of osteoporosis

is essential to developing successful therapies and preventative regimens.

1.3. GLUCOSE HOMEOSTASIS AND DIABETES MELLITUS

1.3.1. Glucose homeostasis

Glucose is the most commonly used fuel for generation of energy rich ATP
in the tissues. Blood glucose levels are regulated by the hormones insulin and
glucagon. High blood glucose levels (such as after a meal) stimulate insulin
secretion from the pancreas into the blood. Insulin then binds insulin receptors on
target tissues (liver, muscle, fat), which initiates an intracellular signaling cascade
that results in increased plasma membrane expression of GLUT4, the insulin
responsive glucose transporter. GLUT4 mediates the uptake of glucose into the
cell, where it is stored in the form of glycogen in liver and muscle, or lipid in
adipose tissue. Alternatively, low blood glucose levels (during fasting or exercise)

trigger glucagon secretion from pancreatic a-cells. Glucagon stimulates liver

glycogenolysis, gluconeogenesis, and subsequent increased glucose production

(2).

1.3.2. Diabetes mellitus definition and classifications

14

Diabetes mellitus is a metabolic disease in which a defect in insulin action
(either by lack of insulin or tissue insulin resistance) results in impaired glucose
homeostasis. Diabetes is a broad term that encompasses type 1 diabetes
mellitus (T 1-diabetes), type 2 diabetes mellitus (T 2-diabetes) and gestational
diabetes mellitus (GDM).

T1-diabetes is characterized by hypoinsulinemia and subsequent
hyperglycemia caused by death of insulin-secreting pancreatic (El-cells.
Depending on the etiology of the disease, T1-diabetes has been divided into type
1A (immune mediated and more prevalent) and type 18 (not immune-mediated
and less prevalent). In type 1A diabetes, immune destruction of pancreatic [3-
cells is confirmed by the presence of anti-islet autoantibodies in humans (2).
Type 1A diabetes is heterogeneous, with several loci linked to the disorder (20).
The only environmental factor demonstrated to induce type 1A diabetes in
humans is congenital rubella infection, although other factors may contribute and
could be difficult to identify (2, 21).

T1 -diabetes generally occurs in'adolescents or young adults and must be
controlled by regular blood glucose monitoring and insulin delivery. Better
education, control of the disease, and treatment of life-threatening complications
have helped T1 -diabetes patients to live longer. Although patients may be
considered to have blood glucose under control, no insulin delivery method is as
precise as the pancreas, therefore fluctuations in blood glucose levels continue,

which puts patients at risk for additional complications.

15

T2-diabetes is associated with resistance to insulin signaling in the tissues
(pancreas, liver, muscle and brain) and increased insulin production by the
pancreas. Long-term high demand for insulin results in eventual pancreatic B—cell
death, after which patients are T1-diabetic and dependent on insulin injections for
glucose control. Risk factors for T2-diabetes include obesity (either genetic, or as
a result of poor diet and inactivity) and fetal malnutrition or over nutrition (2).
Gestational diabetes is a form of insulin resistance that occurs during pregnancy

and it usually reverses itself after birth.

1.3.3. Complications of diabetes mellitus

Osteoporosis from T1-diabetes is the main focus of this dissertation and
will be discussed at length in Section 1.4. Other complications from diabetes
include retinopathy, nephropathy, peripheral neuropathy and impaired wound
healing. Hyperglycemia induces elevated reactive oxygen species (ROS) and
advanced glycation end products (AGE), which can cause cell death and are
thought to perpetrate many diabetic complications. These complications also
arise in part from changes in microvasculature. Macrovascular changes from
both T1- diabetes and T2-diabetes leave patients at higher risk for cardiovascular

disease than non-diabetics (2). Risk for each complication increases with

increasing glycated hemoglobin (Hb A10, a measure of glucose control) (2, 22),

therefore tight control of blood glucose is important. However, no treatment for

16

diabetes can control glucose as well as a functional pancreas, so complications

from diabetes cannot simply be treated with insulin.

1.3.4. Animal models of T1-diabetes and T2-diabetes

Animal models of diabetes are important for studying not only the disease
itself, but also its many complications, including osteoporosis. Most rodent
models of type 1 diabetes are either pharmacologic or spontaneous (23), while
type 2 diabetes can be modeled pharmacologically, with spontaneous genetic
rodents, and with administration of a high-fat diet. Pharmacologic agents used to
mOdeI type 1 diabetes include streptozotocin, alloxan, dithizone, vacor and 8-
hydroxyquinolone (23). Widely used to study diabetic complications,
streptozotocin (STZ) is a glucose mimetic that diffuses into insulin secreting
pancreatic B-cells through the non-insulin responsive glucose transporter GLUT2
(24). The effects of STZ include DNA damage and subsequent [ES-cell death (24).
In mice, five daily low dose intraperitoneal injections (40 mg/kg body weight)
induce significantly detectable non-fasting hyperglycemia within three days of the
first injection (See Chapter 2) (25) and a full type 1 diabetic phenotype of
hyperglycemia and hypoinsulinemia after twelve days (26). A single large dose
(5060 mg/kg) of STZ is effective at inducing type 1 diabetes in rats (24).
Injecting 100 mg/kg STZ soon after the birth of rats induces mild hyperglycemia
and impaired glucose tolerance by 8-10 weeks of age and is, therefore, a

pharmacologic method of studying type 2 diabetes in rodents. Pharmacologic

17

induction of diabetes can be performed in any age of rodent and in animals
(primarily mice) with genetic manipulations, making it a convenient and easily
controlled model. However, to confirm results from STZ and other pharmacologic
models, investigators often turn to spontaneous models of diabetes mellitus.

Spontaneous mouse models of type 1 diabetes include the non-obese

diabetic (NOD) model and the Inez“.Aklta mouse. The NOD mouse generally

develops diabetes sometime between 12 and 30 weeks of age (23, 27), making it
difficult to study complications that are affected by age, such as osteoporosis.
Additionally, only 60% of male mice actually develop a diabetic phenotype (23),

which increases the number of animals necessary for a complete study. The

+ - ‘ . . .
lnsZ lAkita mouse is heterozygous for the insulin gene. Demand on the pancreas

for insulin production from only one allele leads to B-cell death within 4-6 weeks
of life (28). Because it occurs early and consistently, diabetes induction is much
easier to predict and better for examining diabetic complications at early time
points. Additionally, these mice can be used to understand complications in
young mice that are still growing, which is when type 1 diabetes generally
develops in humans. Spontaneously diabetic BioBreeding (BB) rats develop type
1 diabetes around 12 weeks of age, which is during puberty, making the time of
induction representative of human disease (23). Pathogenesis in BB rats is a
result of autoimmune attack of the pancreas, similar to humans (23).
Unfortunately, BB rats develop severe diabetes and ketoacidosis and survival is
dependent on insulin treatment (23), which is similar to the human situation, but

adds complexity to data interpretation.

18

ll

Non-pharmacologic models of T2-diabetes include the leptin deficient
oblob and leptin receptor deficient db/db mice. The absence of leptin or leptin
signaling (respectively) induces increased food consumption to the point of
developing obesity, impaired glucose tolerance, increased insulin production, and
eventual pancreatic beta cell death due to high demand on the pancreas (29).
The Zucker (fa/fa) rat is also a model of leptin resistance, similar to db/db mice
(23, 30). Several other spontaneous rodent models exist that each have slight
variations in diabetes induction, severity, and phenotype of complications,
making them useful for studying the human condition, which is in itself
heterogeneous (23). T2-diabetes can also be induced by administering research

animals with a high fat diet (31).

1.4. BONE PHENOTYPE IN DIABETES MELLITUS

1.4.1. Type 2 diabetic bone phenotype

In human studies, T2-diabetes has been associated with increased risk of
nonvertebral fracture, despite increased cortical thickness and increased
trabecular bone mineral density (BMD) (1). Additionally, some fractures in type 2
(and type 1) diabetic patients are thought to be due to increased falls as a result
of neuropathy and poor vision, but this does not completely explain the increased
fracture risk phenotype (32). Adjusting for body mass index (BMI) reduces the

correlation between bone density and diabetes, however it does not completely

19

eliminate statistical significance (1). Hyperinsulinemia associated with insulin
resistance has been thought to perpetrate increased bone density that cannot be
correlated to body mass (33). Medications, age and progression of T2-diabetes
(all of which can have bone effects) have complicated interpretations of human
studies. It is likely that more rapid. bone loss occurs in T2-diabetic patients over
with aging, especially if pancreatic B-cell function ceases, and insulin secretion
stops (32).

The bone phenotype of mouse models of T2-diabetes is variable
depending on the model examined. Leptin/leptin receptor—deficient mice have
high bone mass early in life, then because of the deficiency in the leptin pathway,
become obese, insulin resistant, and eventually lose B-cell function (34, 35).
When diabetic, these mice have reduced bone formation and increased marrow
adiposity in the long bones, but do not have bone loss or increased marrow
adiposity in the vertebrae (36-39). This is not a perfect model of T2-diabetes,
since leptin is known to have direct effects on the skeleton. Leptin deficient mice
will be discussed in further detail in Sections 1.4.3.2.1. Alternately, in high fat diet
induced obesity and diabetes more closely mimics the human etiology, but in
these mice, bone loss is primarily due to increased resorption, whereas it

remains unclear how resorption is altered in T2-diabetic patients (32, 40, 41).

1.4.2. Type 1 diabetic osteoporosis

20

Osteoporosis is a serious complication of T1-diabetes, leaving patients at
risk for bone loss, fracture and impaired fracture healing (42-44). Young patients
diagnosed with T1-diabetes have reduced growth, which correlates with the level
of glycemic control (1, 45, 46). Serum osteocalcin is significantly reduced in
diabetic patients of all ages, suggesting reduced bone formation (47). In the
same study, serum procollagen carboxy-terminal extension peptide (PICP), a
marker of bone resorption, was unchanged.

Rodent models of T1-diabetes have a bone phenotype similar to that of
humans. In both STZ-mice and rats, T1-diabetes causes bone loss and impaired
bone healing (48-54). Non-obese diabetic (NOD) mice are susceptible to
development of autoimmune T1 -diabetes and display a similar bone phenotype
to that of STZ-diabetic mice (55). Spontaneously diabetic BioBreeding (BB) rats
also have bone loss due to decreased bone turnover and impaired intestinal

calcium absorption (56, 57).

1.4.3. Mechanisms of type 1 diabetic bone loss

Understanding the mechanism(s) of reduced bone formation in type 1
diabetes is essential for determining what treatments will be most effective at
improving bone density, strength and healing in patients (26). We do not believe
that hypoinsulinemia directly contributes to reduced osteoblast activity because
euglycemic insulin receptor knockout (IRKO) -L1 mice that do not express insulin

receptor (and therefore have no insulin signaling) in osteoblasts do not have

21

reduced bone density or reduced bone remodeling normally seen in T1-diabetes
(58). Other potential mechanisms of reduced bone formation include bone
inflammation, altered lineage selection of MSCs, hyperglycemialhyperosmolarity
and reactive oxygen species (ROS). It is important to keep in mind that reduced
osteoblast activity could be a result of not only impaired differentiation, but also
increased apoptosis. We are currently examining the extent to which increased
apoptosis contributes to diabetic bone loss, and it will be briefly discussed in

relation to PTH treatment in Chapter 6.

1.4.3.1. Characterization of early bone metabolism and bone inflammation

While our lab has demonstrated bone loss and reduced osteoblast
markers 5 days after confirmation of diabetes (equivalent to 17 days after the first
STZ injection (days post injection, dpi)) (48, 59), understanding temporal
changes in bone during diabetes induction may help elucidate the mechanism of
diabetic osteoporosis. Hyperglycemia has been associated with increased
inflammatory markers (C-reactive protein (CRP), TNFa, and adiponectin), which
can increase osteoblast apoptosis, ROS and impair maturation (60-63). lf
inflammation was in part responsible for impaired bone formation, it is
presumable that we would be able to detect it either systemically or locally in
bone at some point before bone changes are apparent. Therefore, in Chapter 2
we examined the bone metabolism of control and STZ mice at several time

points between 1 and 17 days after the first STZ injection (days post injection,

22

dpi). Here we also characterized proinflammatory cytokine markers in bone and
serum, and tested whether deficiency of one of the altered cytokines, interferon

gamma (lFN-y) could ameliorate diabetic bone loss.

1.4.3.2. ls diabetes-induced marrow fat bad for bone?

In addition to reduced bone formation and unchanged or reduced
resorption in type 1 diabetic bone, STZ and NOD mice have an obvious increase
in bone marrow adipocyte number (26, 48, 55, 59, 64-66). STZ mice have
increased expression of peroxisome proliferator-activated receptor gamma
(PPARy) 2 in bone, a transcription factor important for adipocyte maturation (48).
Because adipocytes are derived from the same stem cells that give rise to
osteoblasts, the increased marrow adiposity suggests that differentiation of
MSCs may be shunted away from the osteoblast lineage and toward the
adipocyte lineage under diabetic conditions (48). It is also possible that
adipocytes secrete factors (i.e. tumor necrosis factor alpha (T NFa)) into the
marrow microenvironment that reduce osteoblast differentiation or increase
osteoblast apoptosis. High marrow fat has also been implicated in bone loss

models of aging (67, 68) and unloading (69).

1.4.3.2.1. Leptin

23

Interestingly, increased adiposity is not observed in male or female
vertebrae, although it is present in tibia, femur and calvaria (66). In this respect
the T1-diabetic bone phenotype resembles that of leptin-deficient mice (39) and
T1-diabetic mice also have suppressed leptin (66). Leptin is a small protein (16
kDa) secreted by adipocytes and involved in bone mass regulation. The effects
of leptin on bone are complex; it can stimulate or inhibit bone formation
depending upon bone location and whether leptin is functioning directly on
osteoblasts (through receptors (70, 71)) or indirectly through the hypothalamus
(34, 36, 37, 71, 72). In children and during adolescence, decreases in serum
leptin levels, associated with reduced food intake and some disease conditions,
are thought to contribute to reduced bone formation and growth (36, 73).
Absence of leptin in mice also results in bone loss as well as increased bone
marrow adiposity (39, 74). Similarly, increased leptin levels, as observed in
obesity (75), are correlated with increased bone mass (76). Several studies have
tested the potential therapeutic benefits of leptin treatment on bone loss and
marrow adipocyte accumulation. In vitro studies demonstrate that leptin promotes
bone marrow stromal cells to exhibit an osteoblast rather than adipocyte
phenotype (70, 71, 77). Consistent with this finding, subcutaneous infusion of
leptin with osmotic mini-pumps reduces marrow adiposity and increases bone
mass in oblob mice (38). Similarly, leptin treatment reduces bone loss from
ovariectomy (78) and tail suspension (79). In sum, these studies suggest the
efficacy of leptin treatment to restore bone density under conditions of bone loss.

Therefore, we tested the ability of leptin to treat diabetic bone loss in Chapter 3.

24

1.4.3.2.2. C/EBPB and MSC lineage selection

In order to test altered MSC lineage selection as a mechanism of diabetic
bone loss, we previously inhibited PPARy2 with bisphenoI-A-diglycidyl ether
(BADGE) in control and diabetic mice (59). BADGE was effective at reducing
diabetic hyperlipidemia and bone marrow adipocyte accumulation. However,
BADGE did not prevent reduced bone formation (osteocalcin and Runx2
expression) or reduced bone density from T1-diabetes. These results suggest
that marrow adipocyte differentiation is not linked to reduced osteoblast bone
formation. However, PPARyZ expression occurs later than C/EBPB and C/EBP8
in adipocyte differentiation (Figure 3), therefore we could not exclude that MSCs
differentiated into preadipocytes but could not fully mature in BADGE treated
diabetic mice. In order to address this issue, we examined diabetic bone loss in
mice deficient in C/EBPB, an early adipocyte transcription factor, in Chapter 4.

CCAAT enhancer binding proteins (C/EBPs) are members of the basic
region-leucine zipper (bZIP) class of transcription factors. A key regulator of
adipocyte lineage selection C/EBPB, it is transiently expressed during early
adipocyte differentiation with C/EBP8, and then followed by expression of
PPARyZ and C/EBPa (8-10). C/EBPB exists in three isoforms: transcriptionally
active LAP-1 and LAP-2 and generally inactive LIP (10). Overexpression of
C/EBPB alone induces adipocyte differentiation in NIH-3T3 fibroblasts, while LIP

overexpression can prevent it (80, 81). Total deficiency of C/EBPB protects mice

25

from obesity and reduces body fat mass (82-84). Combined knockout of C/EBPB
and CIEBP6 leads to an even greater block to the adipocyte phenotype and
adiposity (85).

C/EBPB has been shown to affect osteoblast differentiation and bone
density. These effects are dependent on when (early vs. late differentiation),
where (local vs. systemic), and which (LAP vs. LlP, or both) CIEBPB is
expressed. C/EBPB is normally expressed during early and late stages of
osteoblast differentiation, with decreased expression during middle stages (86).
Complete knockout of C/EBPB in mice results in decreased total body mass and
total bone mineral density (BMD) (83). Similarly, targeted expression of the
C/EBPB inactive form, UP, to preosteoblasts and osteoblasts results in

osteopenia in transgenic mice due to decreased bone formation (87).

1.4.3.2.3. Wnt/B-catenin signaling

Wnt/B—catenin signaling through TCF/LEF-induced transcription is a potent
regulator of bone formation and adipocyte differentiation (88, 89). Wnts are
secreted ligands that bind low density lipoprotein receptor-related protein 5/6
(LRP5/6) and frizzled (de) membrane receptors (Figure 5). Dimerization of the
LRP and de receptors in response to wnts initializes a signaling cascade that
inhibits glycogen synthase kinase 3 beta (GSK38). Without the wnt signal (or in
the presence of endogenous pathway inhibitors such as dickkopf (Dkk) proteins),

GSK38 actively phosphorylates cytoplasmic B-catenin, targeting it for

26

degradation. When GSK38 is inactive (as in the presence of a wnt ligand),
transcriptionally active B-catenin (dephosphorylated on serine (Ser) 37 or
threonine (Thr) 41) accumulates in the cytoplasm and translocates to the nucleus
where it initiates transcription of genes with TCF/LEF binding sites, such as
Runx2 (90, 91).

Mice null for LRP5 have low bone density and blindness, consistent with
the rare recessive disorder, osteoporosis-pseudogIioma syndrome (OPPG), while
gain of function of LRP5 results in high bone mass due to increased osteoblast
activity (92-97). Although complete loss of function of LRP6 is fatal, loss of one
allele causes more bone loss in mice deficient in LRP5 by exacerbating reduced
bone formation (98). Similarly, modulation of Wnt10b affects only bone formation,
not resorption (99). Wnt10b knockout (-KO) mice have low bone density while
mice overexpressing Wnt10b (Wnt10b-Tg) have high bone density and low
marrow adiposity (99). These mice have attenuated bone loss from aging and
ovariectomy (99). In vitro activation and inhibition of wnt signaling results in
inhibition and activation of adipogenesis, respectively (100, 101), through
regulation of expression of PPARy and C/EBPo (102, 103). Wnt1, 5a and 7b
have also been shown to promote osteoblast and/or suppress adipocyte
differentiation (98).

Endogenous agonists and antagonists can further regulate wnt signaling.
Dally protein enhances the interaction between wnts and des, while Dkks and
secreted frizzled-related proteins (SP RP) antagonize it. Additionally, sclerostin,

prevents the de-LRP interaction. Targeted inhibition of Wnt agonists and

27

antagonists is a relatively new and promising area of interest for treatment of
osteosclerotic and osteoporotic diseases (89). We therefore examined Wnt
pathway family member changes in diabetic bone, and examined the ability of

overexpression of Wnt10b to counteract osteoporosis in STZ mice in Chapter 5.

28

jl-catenln

jl-Catenin
his.

 

Wnt ligands absent

I
l
I
I
I
I
I
I
I
I
I
I
I
I
g
: GSK3B
I
I
I
I
l
l
I
I
I
I
, ‘ I
Wntligands present :

Figure 5. B-catenin signaling in the presence and absence of wnt ligands. In
the presence of wnt ligands (left), B-catenin accumulates in the cytoplasm and
translocates to the nucleus, where it can activate TCF/LEF responsive gene
transcription. In the absence of wnt ligands or when Dkk binds frizzled (right), 8-
catenin is targeted for degradation by GSK3B.

29

1.4.4. Treatment of T1-diabetic bone loss

1.4.4.1 . Antiresorptive treatments

Most of the treatments for osteoporosis are antiresorptive, meaning they
work by inhibiting osteoclast activity. Bisphosphonates (the major antiresorptive
therapy) have a similar structure to inorganic pyrophosphate and is incorporated
into bone (104). When osteoclasts encounter bisphosphonates embedded in
bone, resorption is halted, which has recently been associated with increased
fracture risk (105-I107). Additionally, one of the well-characterized side effects of
bisphosphonates is osteonecrosis of the jaw (bisphosphonate-related
osteonecrosis, BON) (108, 109). Recent clinical evidence suggests that diabetes
is a risk factor for BON: compared to the total patient population receiving
bisphosphonates for osteoporosis, diabetic patients receiving bisphosphonates
were nearly 5 times more likely to be diagnosed with BON (110). Other
antiresorptive treatments for osteoporosis include hormone replacement therapy
and selective estrogen receptor modulators (SERMs), and calcitonin. However,
because bone loss from T1-diabetes is due to reduced osteoblast bone
formation, and not increased resorption, therapies that target bone formation

directly may be most appropriate.

1.4.4.2. Anabolic intermittent PTH

30

Anabolic treatments for osteoporosis are those that promote bone
formation, rather than inhibit resorption. Currently, the only anabolic treatment in
use is a truncated form of endogenous PTH. Also called teriparatide, PTH(1-34)
is administered intermittently by daily subcutaneous injections and can cause a
net increase in bone formation and reduction of fracture risk (111-113). In
humans, PTH(1-34) is used to treat only severe osteoporosis because of
elevated risk of osteosarcoma in rats treated with higher than approved human
doses (114, 115). However, use of PTH(1-34) in a broader patient group may
prove to be appropriate, especially when osteoporosis is caused by bone
formation defects, as in T1-diabetes.

There are no reports examining the efficacy of PTH(1-34) treatment for
T1-diabetic bone loss in humans. In laboratory animals, intermittent PTH(1-34)
treatment has proven to be anabolic (111), and is effective at increasing bone
formation in models of unloading, ovariectomy, and alcohol consumption (113,

1 16—120). Additionally, 4-week intermittent PTH(1-34) treatment of STZ-diabetic
rats improves bone density parameters 4, 6 and 8 weeks after diabetes induction
(121). However, the effects of PTH(1-34) therapy on mouse models of diabetes-
induced bone loss have not been determined. In Chapter 6, we tested whether
intermittent PTH(1-34) treatment could be beneficial for T1-diabetic patients, by
examining the ability of PTH(1-34) to prevent and reverse diabetes-induced bone

loss in STZ mice.

31

1.5. SUMMARY

Understanding the mechanism of any disease or disease complication is
imperative for proper treatment. Using mouse and cell culture models, our past
studies have demonstrated that hyperglycemia and T1-diabetes reduce
osteoblast differentiation and increase adipocyte differentiation in the marrow.
The following chapters characterize the time course of bone phenotype changes
in diabetes, with regard to osteoblast/adipocyte differentiation and inflammation
in bone. Additionally, I examined the effects of two commercially available
treatments (leptin and intermittent PTH) and two genetic manipulations (C/EBPB
knockout and overexpression of Wnt1 Ob) on diabetic bone changes. My
experiments have provided us with new insight into the mechanisms of diabetic
bone loss (which are most likely multi-faceted) and suggested how best to treat it
with the therapies available today. Finally, these chapters provide a launching
pad for additional investigation into bone changes from diabetes, which will likely

provide new insights into our understanding of bone itself.

32

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46

CHAPTER 2

Motyl KJ*, Botolin S*, Irwin R*, Appledorn DM, Kadakia T, Amalfitano A,
Schwartz RC, McCabe LR 2009 Bone inflammation and altered gene
expression with type I diabetes early onset. J Cell Physiol 218:575-583

*authors contributed equally.

47

CHAPTER 2

2. BONE INFLAMMATION AND ALTERED GENE EXPRESSION WITH TYPE I
DIABETES EARLY ONSET

2.1 . ABSTRACT

Type I diabetes is associated with bone loss and marrow adiposity. To
identify early events involved in the etiology of diabetic bone loss, diabetes was
induced in mice by multiple low dose streptozotocin injections. Serum markers of
bone metabolism and inflammation as well as tibial gene expression were
examined between 1 and 17 days post injection (dpi). At 3 dpi, when blood
glucose levels were significantly elevated, body, fat pad and muscle mass were
decreased. Serum markers of bone resorption and formation significantly
decreased at 5 dpi in diabetic mice and remained suppressed throughout the
time course. An osteoclast gene, TRAP5 mRNA, was suppressed at early and
late time points. Suppression of osteogenic genes (runx2 and osteocalcin) and
induction of adipogenic genes (PPARyZ and aP2) were evident as early as 5 dpi.
These changes were associated with an elevation of serum cytokines, but more
importantly we observed an increase in the expression of cytokines in bone,
supporting the idea that bone, itself, exhibits an inflammatory response during
diabetes induction. This inflammation could in turn contribute to diabetic bone

pathology. Mice deficient in IFN-y (one of the key cytokines elevated in bone and

48

known to be involved in bone regulation) did not prevent diabetic bone pathology.
Taken together, our findings indicate that bone becomes inflamed with the onset
of T1-diabetes and during this time bone phenotype markers become altered.
However, inhibition of one cytokine, lFN-y was not sufficient to prevent the rapid

bone phenotype changes.

2.2. INTRODUCTION

Type I diabetes is a complex disorder associated with multiple long term
complications. It evolves with progression to nearly complete beta cell.
destruction and establishment of a hyperglycemic state. Unfortunately, at the
time clinical symptoms present, pancreatic B-cells are irreversibly and almost
completely damaged. This makes it difficult to monitor and understand the early
events in the etiology of diabetic complications. Data on the progression of early
changes is scarce and comes from experimental models of induced diabetes
such as streptozotocin injected diabetic mice. In this model, animals are injected
daily with streptozotocin, for 5 consecutive days. Streptozotocin is a nitrosourea
derived drug from Streptomyces achromogenes which causes pancreatic B-islet
infiltration of T cells and macrophages in rodents, similar to the histology of
human type I pancreatic biopsies (1, 2). Monitoring of the glycemic profile in
fasting rats following streptozotocin (80 mg/kg by intraperitoneal injection)
treatment demonstrates a rapid response with evidence of hyperglycemia and

reduced plasma insulin levels within two hours (3). This response is followed by

49

a drop in blood glucose levels and high plasma insulin levels at 6 hours, but
ultimately a progressive state of chronic hyperglycemia ensues (3).

It is known that one of the complications of type I diabetes is bone loss
and ongoing population studies continue to support these observations in
humans (4-13). Similarly, rodent type I diabetes models also exhibit significant
bone loss (14-22). Examination of bone metabolism markers in humans and
rodent models suggest that diabetic bone loss is primarily due to a defect in bone
formation rather than resorption (17, 18, 23-25). In fact, analyses of serum
markers of bone resorption indicate that, if anything, resorption is actually
decreased in parallel with the decrease in osteoblast activity (18, 25, 26).
However, it remains unknown if osteoclast activation occurs early in the disease

progression and contributes to the significant bone loss seen at later time points.

The above findings support the notion that type I diabetes promotes a true
suppression of bone formation most likely through a reduction of osteoblast
numbers and/or maturation. Previously, we demonstrated that streptozotocin-
induced diabetes in BALBlc male mice is associated with bone loss and
suppression of serum osteocalcin and osteocalcin mRNA levels in bone (18). In
addition, visible changes in bone histology marked by increased marrow .
adiposity were observed and further confirmed by increased mRNA levels of
markers of adipocyte maturation, PPARy2 and aP2 (18). The elevated
expression of adipogenic markers and decreased expression of osteogenic
markers was apparent as early as 17 and 19 days after the first injection of

streptozotocin (18, 27). Considering the present literature on the timing of early

50

changes in streptozotocin induced diabetes, and our previous data, we
hypothesize that the commitment to a suppressed osteoblast phenotype and
increased bone marrow adiposity in diabetes occurs early during the onset of
diabetes. Here we demonstrate that osteoclast activity is not increased during
the progression of diabetes as and is actually significantly reduced by 5 dpi, as
determined by serum PYD and TRAP5 mRNA, although we did observe an early
decrease and an rapid increase at 3 dpi in RANKL expression. We also found a
reduction in serum osteocalcin and mRNA levels of osteogenic markers and an
increase in adipocyte markers within 5 to 7 days dpi (18, 27). Systemic and local
cytokine expression increased during this time, indicating that bone is a site of
inflammation during disease onset. These findings indicate that diabetes-
induced changes in bone phenotype and inflammation occur rapidly and in
parallel with changes in metabolic status. Of the cytokines elevated in bone RNA
samples, IF N-y (known to contribute to the regulation of bone remodeling (28-
37)) was found to be consistently elevated early and during bone phenotype
adaptation to diabetes. However, IF N-y deficiency was not sufficient to prevent
diabetic bone loss, suggesting that other cytokines, factors or combinations of

factors may need to be targeted to prevent this diabetes complication.

51

2.3. MATERIALS AND METHODS

2.3.1. Streptozotocin mouse injections

Adult (15 week old) male wild type or lFN-y-l- mice (BALB/c strain;

Jackson Laboratories, Bar Harbor, ME) were intraperitoneally injected daily with
streptozotocin (40 pig/g body weight in 0.1 citrate buffer) for 5 days (38, 39).
Controls were injected with citrate buffer alone. Twenty-four hours after the first

injection, animals were considered as 1 day post injection. Wild type mice were
“New“ 1' 3' 5. 7. 9’ 11: 17 or 32 days post the first injection (dpi). IFN-Y-I-

mice were harvested 32 dpi, after bone loss was visible with mCT. All mice were
kept on a light/dark (12hl12h) cycle at 23°C, and received food (standard lab
chow) and water ad Iibitum. Mice were euthanized and tibiae were immediately
removed, freed from soft tissue, snap-frozen in liquid nitrogen, and stored at -
80°C (for RNA analyses). Animal studies were conducted in accordance with the

Michigan State University Institutional Animal Care and Use Committee.

2.3.2. Genotyping

Knockout of lFN-y was confirmed by RT-PCR as described below with
primers specific for the wild type and knockout gene locus, according to the
protocol provided by The Jackson Laboratory. VIfild type primers were 5'-AGA
AGT AAG TGG AAG GGC CCA GAA G-3' and 5'-AGG GAA ACT GGG AGA

52

GGA GAA ATA T-3’. Knockout primers were 5’-TCA GCG CAG GGG CGC CCG
GTI' C‘I'l' T—3’ and 5’-ATC GAC AAG ACC GGC TTC CAT CCG-3’. PCR
amplicons were separated on a 2% agarose DNA gel with a 100 bp DNA ladder

(Invitrogen, Calsbad, CA).

2.3.3. Plasma measurements

Blood was obtained from mice at the time of euthanasia and blood serum
prepared from each sample by centrifugation for 5 min at 3000 rpm. Serum was
stored frozen at -20°C. Glucose concentration in serum samples was
determined using a Glucose Assay Kit (Sigma, Saint Louis, MO). Serum
osteocalcin levels were measured using Mouse Osteocalcin EIA Kit (Biomedical
Technologies Inc. Stoughton, MA, USA) according to manufacturer instructions.
Serum PYD was measured using Metra® PYD kit (Quidel Corporation, San
Diego, CA, USA) according to manufacturer instructions. Quantitative
determination of non-esterified (free) fatty acids in mouse serum was performed
according to manufacturer instructions using Wako NEFA C test kit (Wako
Chemicals USA, Inc. Richmond, VA, USA). Serum cytokines were measured
using the Bio-Plex multiplex mouse cytokine 23-plex bead-based assay
according to manufacture protocol (Bio-Rad, Hercules, CA). Standard curves

were run for each cytokine and serum concentrations were calculated.

53

2.3.4. RNA Analysis

Whole tibias were crushed under liquid nitrogen conditions using a
Bessman Tissue Pulverizer and RNA extracted using the method of
Chomczynski and Sacchi as previously described (40, 41). RNA integrity was
verified by formaldehyde-agarose gel electrophoresis. Synthesis of cDNA was

performed by reverse transcription with 2 pg of total RNA using the Superscript II
kit with oligo dT(12-13) primers as described by the manufacturer (Invitrogen,

Carlsbad, CA). cDNA (1 ul) was amplified by PCR in a final volume of 25 III
using the i0 SYBR Green Supermix (Bio-Rad, Hercules, CA) with 10 pmol of
each primer (Integrated DNA Technologies, Coralville, IA). Osteocalcin was
amplified using 5’-ACG GTA TCA CTA TTI' AGG ACC TGT G-3’ and 5’-ACT
TI'A 'ITT TGG AGC TGC TGT GAC-3’ (42). Runx2 was amplified using 5’- GAC
AGA AGC TTG ATG ACT CTA AAC C-3’ and 5’- TCT GTA ATC TGA CTC TGT
CCT TGT G-3’ (43). PPARyZ was amplified using 5’-TGA AAC TCT GGG AGA
'I‘I'C TCC TG-3’ and 5’-CCA TGG TAA 'I'I'T CTI' GTG AAG TGC-3’ (44).
Adipocyte fatty acid-binding protein 4 (aP2) was amplified using 5'-GCG TGG
AAT TCG ATG AAA TCA-3’ and 5’-CCC GCC ATC TAG GGT TAT GA-3’ (45).
TRAP5 was amplified using 5'-AAT GCC TCG ACC TGG GA—3' and 5'-CGT AGT
CCT CCT TGG CTG CT-3' (46). RANKL was amplified using 5’-'ITI' GCA GGA
CTC GAC TCT GGA G-3' and 5’-TCC CTC CTI' TCA TCA GGT TAT GAG-3'

according to Zhao et al. (47). OPG was amplified using 5’-GAA GAA GAT CAT

54

CCA AGA CAT TGA C-3’ and 5’-TCC ATA AAC TGA GTA GCT TCA GGA G-3’.
lL-6 was amplified using 5’-ATC CAG TTG CCT TCT TGG GAC TGA-3’ and 5’
TAA GCC TCC GAC TTG TGA AGT GGT-3’. The following cytokine primers and
their PCR protocols are previously described: lL-1 Ra (48), LT-j3 (49), IL-1a (50),
lFN-y (51), MCP-1 (52), KC (52), MlP-1B (53) and TNF-a (54). Cyclophilin,
which was not modulated under diabetic conditions, was used as a control for
RNA levels; it was amplified using 5'-ATT CAT GTG CCA GGG TGG TGA C-3’
and 5’-CCG TTI' GTG ‘ITT GGT CCA GCA-3’ (55, 56) and exhibited similar
kinetics of amplification compared to other genes examined. In some cases
HPRT was also used as a non-modulated control gene and was amplified using
5’-AAG CCT AAG ATG AGC GCA AG-3’ and 5’-TTA CTA GGC AGA TGG CCA
CA-3’ (57). Real time PCR was carried out for 40 cycles using the iCycler (Bio-
Rad, Hercules, CA) and data were evaluated using the iCycler software. Each
cycle consisted of 95°C for 15 seconds, 60°C for 30 seconds (except for runx2
and osteocalcin, which had an annealing temperature of 65°C) and 72°C for 30
seconds. RNA-free samples, a negative control, did not produce amplicons.
Melting curve and gel analyses (sizing, isolation and sequencing) were used to
verify single products of the appropriate base pair size.

Cytokine expression was also examined by ribonuclease protection assay

(RPA) using the multi template probe sets mCK—2b and mCK-3b according to the

manufacturer instructions (BD Biosciences, California BD RionuantTM assay).

Briefly, RNA (5 gig) was hybridized to 1 x 106 cpm of the probe in 10 iii of

hybridization buffer. Samples were denatured at 95°C, incubated at 56°C for 16

55

hours, treated with RNase (19.2 ngl100 pl) for 45 minutes at 30°C, and incubated
with proteinase K (27 mg/18ul) for 15 minutes at 37°C. Precipitated RNA was
resuspended in 5 pl of gel loading buffer and resolved on a ~8 M urea, 4.75%
polyacrylamide gel. Radiolabeled probe (alone) was used as a marker for each
cytokine band. Controls include samples without RNA and a) with RNase to
demonstrate efficient RNA degradation and b) without RNase to detect general

degradation of probe.

2.3.5. Micro-computed tomography (pCT) analysis

Tibias were scanned using a GE Explore Locus pCT system at a voxel

resolution of 20 pm obtained from 720 views. Each run included control and
diabetic, wild type and IF N-y'l' bones, and a calibration phantom to standardize

grayscale values and maintain consistency. Based on auto threshold and
isosurface analyses of multiple bone samples, a fixed threshold (1400) was used
to separate bone from bone marrow. Trabecular bone analyses were done in a
region of trabecular bone defined at 0.17 mm (approximately 1% of the total
length) immediately distal to the growth plate of the proximal tibia extending 2
mm toward the diaphysis, and excluding the outer cortical shell. Trabecular bone
volume fraction (BVF) was computed by GE Healthcare MicroView software or

visualization and analysis of volumetric image data.

56

2.3.6. Statistical analysis

All statistical analyses were performed using Microsoft Excel data analysis

program for t-test analysis. Values are expressed as a mean a: SE.

57

2.4. RESULTS

Previously, we demonstrated that multiple low dose streptozotocin-
induced diabetes caused changes in body weight, serum parameters and bone
histology, as well as gene expression resulting in bone loss (18). We also
reported that these changes were evident five days after confirmation of diabetes
(58). Based on the protocol for diabetes induction (daily injections of
streptozotocin for 5 days followed by a 7 day period prior to confirmation of
diabetes), the day 5 time point in our past studies was actually 17 days after the
animals received their first injection of streptozotocin (17 dpi). The marked
changes that we observed at this time suggested that the events involved in the
changes in bone and serum parameters must occur earlier. To identify the
progression of these events, we injected mice with streptozotocin and examined
serum, muscle and bone parameters at 1, 3, 5, 7, 9, 11, and 17 dpi of
streptozotocin or citrate buffer (vehicle control).

The STZ-injected mice exhibited weight loss at 3 dpi which plateaus by 5
dpi (Figure 6). Blood glucose levels were significantly increased at 3 dpi and
continued to rise over the course of diabetes induction (Figure 6). The
immediate change in body mass was not associated with elevated serum free
fatty acids (Figure 6), but was directly associated with the loss of femoral fat pad
weight and size, which were significantly reduced at 3 dpi and reached maximum

reduction in size at 5 dpi (Figure 6). The mass of the tibialis anterior muscle was

58

also significantly reduced with the onset of diabetes (Figure 6), indicating a

reduction in mass of several mesenchymal derived tissues.

59

Figure 6. Time course of general body parameters during the onset of type I
diabetes in BALBIc mice. Diabetes was induced by 5 daily streptozotocin
injections beginning on day 0. Body, muscle (tibialis), and fat pad (femoral) mass
and serum parameters (glucose and free fatty acids (FFA)) were measured at 1,
3, 5, 7, 9, 11, and 17 days post injection (dpi) of streptozotocin (triangles) or
citrate buffer (vehicle control, squares). Values represent averages obtained from
56 mice per condition per time point +/- SE. *p<05.

60

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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Previously, we demonstrated that at 17 dpi, diabetic mice exhibited
decreased serum levels of osteocalcin (bone formation marker), decreased
expression of mature osteoblast markers and increased expression of adipocyte
markers in bone. To determine when the onset of these changes first occurred,
we collected serum and isolated tibia RNA from 8-11 mice per condition at 1, 3,
5, 7, 9, 11, and 17 dpi of streptozotocin or citrate buffer (vehicle control). At 5
dpi, the onset of diabetes significantly decreased serum osteocalcin levels
indicating that bone formation was suppressed immediately (Figure 7A).
Consistent with the suppression of serum bone formation markers, osteocalcin
mRNA levels were significantly reduced in diabetic tibias at 5 dpi. Osteocalcin
mRNA levels also trended to decrease within 24 hours of the first streptozotocin-
injection. Similar changes were seen with runx2 mRNA levels. Specifically,
runx2 expression was significantly suppressed at 1 and 5 dpi and beyond.
Interestingly, we found that adipocyte markers were equally responsive and that
induction of diabetes upregulates aP2 and PPARy2 mRNA levels as early as 5
and 7 dpi, respectively (Figure 7B). This correlates with the elevation of blood

glucose and other early events of diabetes that we observed.

62

Figure 7. Significant bone phenotype changes are evident at 5 dpi. A.
Serum osteocalcin (OC) levels were determined in mice injected with citrate
buffer (controls, squares) or streptozotocin (triangles) at 1, 3, 5, 7, 9, 11, and 17
dpi. Values are averages +/- SE obtained from 8-11 mice per condition. *p<0.05.
B. Total RNA was extracted from tibia isolated from citrate buffer (vehicle control,
squares) and streptozotocin (triangles) injected mice at 1, 3, 5, 7, 9, 11, and 17
dpi. Levels of osteocalcin (OC), runx2, PPARy2 and aP2 mRNAs were
determined by real time RT-PCR and are expressed relative to cyclophilin
(housekeeping gene). Values are averages +/- SE obtained from 5-6 mice per
condition per time point. *p<0.05.

63

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Our past studies have indicated that serum and RNA markers of
osteoclast maturation and activity are decreased in T1-diabetic mice. To
determine if these changes are apparent early in T1-diabetes onset we
measured several osteoclast parameters. Figure 8 demonstrates that expression
of osteoclast regulators, osteoprotegrin (OPG) and RANKL, in bone during the
time course was variable. At 1 dpi RANKL was suppressed but was then levels
significantly increased at day 3 and then returned and remained at normal levels
after 5 dpi. OPG levels did not change early on, but were reduced at later time
points. The ratio of the two parameters (not shown) was highly variable in both
control and T1 -diabetic mice. Therefore we examined a serum marker of
resorption, PYD levels, and found a significant decrease at all time points studied
(Figure 8). Serum was examined because T1-diabetic mice exhibit polyurea and
potentially nephropathy, both of which could have a major impact on urine
measurements. As a further assessment of osteoclast activity we measured
TRAP5 mRNA levels in bone (Figure 8) and found that they were decreased at

early time points (1 and 3 dpi) and at 17 dpi, consistent with our previous reports.

65

Figure 8. Analyses of osteoclast regulators, markers and activity. Total
RNA was extracted from tibia isolated from citrate buffer (vehicle control,
squares) and streptozotocin (triangles) injected mice at 1, 3, 5, 7, 9, 11, and 17
dpi. Levels of osteoprotegrin (OPG), RANKL, and TRAP5 mRNAs were
determined by real time RT-PCR and are expressed relative to HPRT
(housekeeping gene). Values are averages +/- SE obtained from 5-6 mice per
condition per time point. *p<0.05. Serum isolated at time points 1, 5, 11 and 17
was analyzed for pyridinoline crosslink PYD levels. Values are averages +/- SE
obtained from 8-11 mice per condition. *p<0.05.

66

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67

We hypothesized that T1-diabetes associated immunologic responses
resulting from early pancreatic beta cell destruction could contribute to changes
observed in bone phenotype. Therefore, we examined the levels of 23 different
cytokines in the serum of control and diabetic mice. The serum profiles during
the onset of type I diabetes varied for each cytokine with some showing no
modulation (IL-2, -3, -5, and lL-12 (p70)), some trending toward an increase at
early time points (p< 0.10; IL-18, lL-9, lL-10, lFN-y and KC), some showing
significant increases at 3 dpi (p<0.05; lL-1a, lL-6, lL-12 (p70), lL-13, IL-17, MCP-
1, MlP-18, and TNF-a) and some showing a significant elevation at later time
points (at 5, 7 and/or 9dpi; IL-17, MCP-1, MlP-1a, MlP-1B, Rantes, and TNF-a)
compared to vehicle treated mice. Figure 9 shows only the serum cytokines that
were significantly elevated by greater than 2-fold at any time point during the
onset of type I diabetes. Cytokines whose levels are statistically higher than
vehicle treated mice prior to and/or during changes in gene expression in bone

(ie, IL-1a, lL-6, lFN-y, TNF-a) could be potential contributors to this process.

68

Figure 9. Serum cytokine levels increase during the early onset of diabetes.
Serum cytokine levels were determined using the Bio-Plex multiplex mouse
cytokine 23 plex bead-based assay (Bio-Rad). Of the 23 cytokines the 7
cytokines shown exhibited the greatest induction at any one time point. Values
are averages +/- SE obtained from 3-6 mice per condition per time point.
*p<0.05.

69

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Locally elevated cytokine gene expression in bone could have an even
greater impact on bone remodeling, since theoretically cytokine concentrations
would be highest at sites of secretion. Therefore, to determine if cytokine
expression is modulated in diabetic bone, we measured mRNA levels of multiple
cytokines using RNase protection assays on RNA isolated from mouse tibias at 5
dpi, the time point at which the earliest suppression of osteoblast gene
expression was seen. Figure 10 demonstrates clear increases in TNF-a, lFN-y,
lL-1Ra and LT-B mRNA levels. Real time RT-PCR confirmed the induction of
. these cytokines at 5 dpi (Figure 11). Cytokine RNA levels were not elevated
prior to this point (data not shown). Examination of additional cytokines revealed
that IL-6 expression was also elevated at 5 dpi while IL-1a was the only cytokine
elevated at 17 dpi. MlP-18, KC and MCP-1 mRNAs were unchanged or
decreased in diabetic bone. The induction of lL—6, TNF-a, IL-1 Ra and LT-B in
diabetic mouse bone was transient and returned to control levels by 7 dpi and.
Only the elevation of lFN-y expression was found to be sustained over a period of
several days, but by day 9 levels were back to control values (not shown) and by
day 17 levels were lower than controls (similar to the LT-B profile). Treatment of
osteoblasts (MC3T3-E1 cells) or isolated calvaria in vitro with STZ at
concentrations up to 0.5 mg/ml did not cause an increase in cytokine expression

(data not shown) supporting a role for diabetes onset (not STZ) in bone

inflammation.

71

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diabetic bone. Representative autoradiograph of an RNase protection assay
using RNA isolated from control (C) and diabetic (D) mouse whole tibias at 5 dpi.

*represents signals significantly increased as determined by densitometry.
*p<0.05.

72

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Figure 11. Cytokine mRNA levels are increased predominantly during the
early stage (5 dpi) of the onset of diabetes. Total RNA extracted from whole
tibias from control (square) and diabetic (triangle) mice during the progression of
diabetes onset. Cytokine mRNA levels were determined by real time RT-PCR
and expressed relative to HPRT levels. Values are averages +/- SE obtained
from 3-6 mice per condition per time point. *p<0.05.

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Given that 1) IFN-y induction was the most sustained of the cytokines we
examined and 2) IFN-y has been demonstrated to contribute to bone pathologies,

we examined the role of lFN-y in diabetic bone loss. To do this, diabetes was

induced in mice that lack expression of lFN-y (IF N-y'l'). The absence of IFN-y

was confirmed by genotyping wild type and lFN-y'l' mice using primers specific

for the wild type or knockout cassette sequences. Specifically, a 320 base pairs
(bp) amplicon is indicative of the defective gene insert with a premature stop
codon while a 260 bp amplicon represents amplification of the wild type gene

(Figure 12A). As shown in Figure 118, streptozotocin was capable of inducing
diabetes in both wild type and lFN-y'l' mice as evidenced by an elevation in blood
glucose levels above 300 mg/dL (Figure 12B). Bones were examined 32 days
after the first streptozotocin injection so that bone loss, if it occurred, would be

great enough to be detected by micro-computed tomography analysis. As shown

in Figure 12C, lFN-y deficiency was unable to prevent diabetes-induced loss of
bone volume fraction (which was not significantly different from wild type mice).

Correspondingly, osteocalcin mRNA levels were suppressed and aP2 levels

Were increased in diabetic IFN- y'l' mice similar to wild type mice (Figure 12C)

and early lineage markers were similarly altered (decreased runx2 and increase

p PARyZ, not shown).

75

Figure 12. IFN-q deficiency does not prevent diabetic bone loss. Diabetes

was induced in wild type and lFN-y 'I' (lFN-y deficient) mice by 5 daily
streptozotocin injections. Bones were analyzed at 32 dpi. A. DNA amplicons from

a PCR reaction with primers specific to wild type or lFN-{l' mice were separated

on a 2% agarose gel. Wild type and lFN-y'l' amplicons ran at the expected
molecular weight of 260 bp and 320 bp, respectively. B. Diabetes was induced in

WT and IFN-y-l' mice as indicated by blood glucose levels greater than 300
mgldL. Values represent mean :1: SE; n > 9 mice per condition. *p<0.05
compared to genotype-matched non-diabetic controls. C. Osteocalcin (OC) and
aP2 gene mRNA levels (obtained by real time RT-PCR analysis of tibial RNA and
quantitated relative to HPRT levels) and tibial bone volume fraction (BVF) were
determined and expressed as % change in diabetic compared to control values

for each genotype, wild type (WT, white bars) and lFN-y'l' (gray bars). Values
represent mean :1: SE; n > 9 mice per condition.

76

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2.5. DISCUSSION

Type I diabetes is associated with significant bone loss (8, 18, 22),
increased bone marrow adiposity (18) and increased fracture risk (9, 59).
Changes in bone phenotype are reported at 2 or 4 weeks post-confirmation of
diabetes. In our previous studies, we looked as early as 5-7 days after the
confirmation of diabetes, which amounts to 17 days after the first injection of
streptozotocin (18, 27). Here we utilized the suppression of pancreatic B-cell
function by multiple low dose streptozotocin injection to analyze rapid changes in
bone that occur at the onset of diabetes prior to the mice attaining maximal
hyperglycemia.

Our findings demonstrated that changes in body mass, blood glucose
levels, fat and muscle mass, as well as bone phenotype occurred before or by 5
days after the first injection of streptozotocin. Pighin et al. (60) also reported
reduced fat pad mass in mice 6 days after injection of streptozotocin. The
adipose tissue Iipolysis that we observed suggests that free fatty acids were
being mobilized as an energy source during the onset of insulin deficiency (60,
61). However, we were unable to detect an increase in serum free fatty acids at
early time points as reported by others (60), but increases were observed at later
time points 17, 19, 26, 33, and 40 dpi (corresponding with days 5, 7, 14, 21 and
28 days after diabetes confirmation) consistent with previous reports (18, 27).

Possibly, at early time points there was a rapid uptake of mobilized fats by

78

tissues in our mouse model. Also, we observed a 3 mM, 6 mM, and then 16 mM
increase in blood glucose levels within 3, 5 and 11 days of streptozotocin
injection, respectively. This is in contrast to Pighin et al. (60) who found that
blood glucose levels were not significantly elevated until day 12. The metabolic
differences between these studies (glucose and lipid levels in the blood) could
stem from differences between mouse strains (C57BU6J versus BALB/c) and/or
differences in streptozotocin effectiveness in early pancreatic B-cell destruction.

Only a handful of studies have examined the early effects of
streptozotocin-induced diabetes on gene expression. Wang et al. (62) reported
some of the earliest changes: a significant reduction of both GLUT2 protein and
mRNA expression in pancreatic islets isolated from streptozotocin-injected mice
4 days after the first streptozotocin injection. In our studies, we saw decreased
runx2 and osteocalcin mRNA levels and increased PPARy2 and aP2 at 5 dpi that
were maintained out to day 17 and are consistent with differences we see at later
stages (ie: day 40 dpi) (18, 27). This suggests that the immediate changes in
mouse blood glucose levels and overall metabolic state may in turn dramatically,
immediately, and chronically influence osteoblast and adipocyte maturation.

In our studies, streptozotocin was used to induce diabetes and allow
identification of the early onset of T1 -diabetes-associated changes in bone
phenotype, something that cannot be done in models of spontaneous T1-
diabetes occurrence. One cannot exclude that secondary effects of
streptozotocin could contribute to our findings. However, STZ alone was unable

to induce a cytokine response in bone in vitro. In addition, studies in other

79

models of diabetes including non-obese T1-diabetic mice (NOD) (22) and in
virally induced diabetic animals (23) demonstrate similar bone pathologies to the
STZ-induced T1-diabetic mouse model. For example, NOD mice exhibit
decreased bone formation and increased marrow adiposity similar to the multiple
low dose streptozotocin model (63). While spontaneously diabetic mouse
models allow analyses without pharmacologic treatments, our studies
demonstrate the utility of the streptozotocin model in the examination of changes
occurring at the onset of diabetes prior to maximum states of hyperglycemia.
This time period would be difficult to study in spontaneously diabetic mice.

While our past studies did not find an enhancement of osteoclast number
and/or activity in T1-diabetic mice at 17, 19, 26, 33, and 40 dpi (18, 27), the
question remained: “does bone resorption/osteoclast activation occur during the
early onset of diabetes prior to these time points?” To answer this question we
monitored the expression of osteoclast markers, activity, and regulators. We did
observe and increase in RANKL at 3 dpi which could be related to early
increases in cytokines that perhaps are not detectable in whole bone RNA
analyses, however, TRAP5 levels were significantly decreased at this time point
and by 5 dpi levels of both parameters were not different from controls. Serum
PYD levels, on the other hand, were significantly decreased in diabetic animals 5
days after the first streptozotocin injection and remained reduced through day 17.
Systemic levels of PYD represent the amount of bone resorption going on in all
bones, which may explain why they are more consistent throughout the time

course. While some studies examine urinary markers of resorption, results can

80

be confounded by secondary effects of T1-diabetes on kidney function and
urinary volume, which is why we focused on serum parameters. However, our
past studies have demonstrated reduced or not modified osteoclast activity by
both serum and urinary parameter measures at later time points (18). Our
findings of reduced or no change in osteoclast activity are consistent with the
majority of reports demonstrating reduced or no change in resorption parameters
in T1-diabetic mouse or rat bones (16-18, 21, 58, 64-69). In addition, in vitro
studies show that high glucose and advanced glycosylation end products can
suppress osteoclast activation and bone resorption (70, 71) and, in T1-diabetic
patients, serum bone resorption markers are unaltered (11, 72, 73) or decreased
(74, 75) compared to controls. However, there are a few contrasting reports
suggesting that advanced glycosylation end products and STZ induced diabetes
can enhance osteoclast activity (76, 77). Differences between studies could
result from the severity of diabetes, different cell lines, mou'se versus rat effects,

and potentially gender differences.

Induction of T1-diabetes in spontaneous models and in streptozotocin-
induced models is associated with pancreatic inflammation. Consistent with this,
our diabetic mice exhibited significant increases in serum cytokine levels
including lL-1a, IFN-y and TNF-a at early time points (3 dpi) and for several
cytokines at later time points (9 dpi). Studies in rats demonstrate increases in
serum lL-18 and TNF-a within 5 days of the first injection of streptozotocin (78).
Similarly, mononuclear/macrophage cells isolated from diabetic mice and

humans exhibit increased IFN-y, TNF-a, lL-18, and lL-6 secretion and/or gene

81

expression compared to controls (79-81). The cytokine profiles are thought to
represent a major T helper (T h)1 cellular immune response that outweighs the
Th2 response (81, 82). Studies examining the influence of Th1 cells on
osteoblast activity indicate that osteoblasts cultured with Th1 cytokine-
conditioned medium exhibit suppression of alkaline phosphatase activity (a
marker of osteoblast maturation) whereas Th2 cytokine-conditioned medium did

not affect alkaline phosphatase activity (30).

Indicative of local bone inflammation, the mRNA levels of several
cytokines were elevated in bone, including LT-B, IL-6, IF N-y and TNF-a.
However, not all of the cytokines elevated in diabetic mouse serum (MCP-1, KC,
MlP-1I3) were elevated in bone. This could be the result of a subpopulation of
immune cells residing in the bone marrow and/or interactions between immune
and bone cells that leads to a unique cytokine profile and/or secondary cytokine
inductions at distal sites. Cytokines such as TNF-a, lL-1 and IFN-‘y are known to
increase osteoblast nitric oxide production and induce apoptosis, and to
decrease growth and maturation in vitro (83-94). In vitro studies also
demonstrate that hyperglycemia is capable of increasing macrophage expression
of lL-1, lL-6 and TNF-a and could therefore contribute to the elevations that we
observe (80). A related induction of the inflammatory response is seen in
hyperglycemic type II diabetic mice in response to P. gingivalis challenge or
fracture conditions; specifically, local diabetic mouse bone exhibits greater

expression of MCP-1, MIP, and TNF-a mRNA compared to control mice (95).

82

Because of the increased cytokine expression during early bone
phenotype changes, we hypothesize that inflammation and cytokine production
are necessary for induction of diabetic bone loss. Under conditions of increased
cytokine production, such as rheumatoid arthritis (without corticosteroid
treatment), bone formation and resorption rates are decreased similarly to T1-
diabetic bones (18, 63, 96, 97). Of the cytokine fingerprint seen in diabetic mice,
we were particularly interested in lFN-y since it is the only cytokine found to be
significantly induced in bone within 24 hours of diabetes induction, is also
elevated at 5 and 7 dpi, is shown to suppress osteoblast growth, maturation and
increase apoptosis in vitro (98, 99), and is demonstrated to regulate bone density
in vivo. Of particular interest is the finding that the ability of Th1 cytokine-
conditioned medium to suppress osteoblast maturation is lost when given in

combination with lFN-y neutralizing antibody (30).

To test whether lFN-y is a key mediator of diabetic bone loss, we induced
diabetes with streptozotocin in lFN-y knockout mice. Based on cell culture

models and the finding that lFN-y treatment increases resorption in patients with
osteopetrosis (33), as well as the observation that lFN-yR-I' mice are protected

from OVX-induced bone loss (100), we hypothesized that IF N-y-deficiency would
reduce the severity of the diabetic bone phenotype. However, we found that
Osteocalcin and aP2 gene expression patterns were similar in both genotypes,
a nd that the magnitude of diabetic bone loss (% change in bone volume fraction)
in the knockout was similar to that of the wild type mice. From these findings we

conclude that IF N-y is not solely responsible for streptozotocin-induced diabetic

83

osteoporosis. This is consistent with reports demonstrating negative (not

protective) skeletal effects in lFNyR1-null (lFN-yR'l') mice, including increased

resorption and susceptibility to arthritic bone loss (32, 101, 102). Bone loss could
result from the loss of IFN-y inhibition of lL-1-induced osteoclast activity (103,
1 04).

In summary, the streptozotocin-induced diabetes mouse model of bone
loss, marked by decreased bone formation, is well suited to studying early
changes in bone phenotype/gene expression. Our studies demonstrate that
changes (suppression of osteogenic genes and induction of adipogenic genes)
are evident within 5 days of the first injection of streptozotocin, a time prior to
observable hyperlipidemia and maximum hyperglycemia. During this period,
there is systemic as well as local bone inflammation, suggesting a potential role
for inflammation in the observed pathological bone changes. Our studies
indicate that lFN-y exhibits early and prolonged elevation during this period, but
deficiency of this single cytokine was not sufficient to prevent bone loss in
diabetic mice. Identifying and understanding the role of factors present at the
early stages of diabetes onset will provide a foundation to identify the cause of

diabetic bone loss and effective preventative treatments.

2.6. ACKNOWLEDGEMENTS

We would like to thank Lindsay Martin for her insightful comments and

Alison Bauer for her help with the lFN-y mice. This work was funded by grants

84

from the National Institutes of Health (RO1DK061184) and the American
Diabetes Association (7-07-RA—105) to LRM. A.A. was supported by the National
Institutes of Health grants RO1DK069884 and P01CA078673, as well the

Osteopathic Heritage Foundation. The authors have no financial conflicts.

85

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promoter. J Immunol 148:788-794

Nanes MS 2003 Tumor necrosis factor-alpha: molecular and cellular
mechanisms in skeletal pathology. Gene 32121-15

Zhou FH, Foster BK, Zhou XF, Cowin AJ, Xian CJ 2006 TNF-alpha
mediates p38 MAP kinase activation and negatively regulates bone
formation at the injured growth plate in rats. J Bone Miner Res 2121075-
1088

Graves DT, Kayal RA 2008 Diabetic complications and dysregulated
innate immunity. Front Biosci 13:1227-1239

Compston JE, Vedi S, Croucher PI, Garrahan NJ, O'Sullivan MM 1994
Bone turnover in non-steroid treated rheumatoid arthritis. Ann Rheum Dis
53:163-166

McCabe LR 2007 Understanding the pathology and mechanisms of type I
diabetic bone loss. J Cell Biochem 102:1343-1357

Smith DD, Gowen M, Mundy GR 1987 Effects of interferon-gamma and
other cytokines on collagen synthesis in fetal rat bone cultures.
Endocrinology 12022494-2499

Chen RM, Chen TL, Chiu WT, Chang CC 2005 Molecular mechanism of
nitric oxide-induced osteoblast apoptosis. J Orthop Res 232462-468

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Cenci S, Toraldo G, Weitzmann MN, Roggia C, Gao Y, Qian WP,
Sierra 0, Pacifici R 2003 Estrogen deficiency induces bone loss by
increasing T cell proliferation and lifespan through lFN-gamma-induced
class II transactivator. Proc Natl Acad Sci U S A 100:10405-10410

Takayanagi H, Sato K, Takaoka A, Taniguchi T 2005 Interplay between
interferon and other cytokine systems in bone metabolism. Immunol Rev
208:181-193

Manoury-Schwartz B, Chiocchia G, Bessis N, Abehsira-Amar O,
Batteux F, Muller S, Huang 8, Boissier MC, Fournier C 1997 High
susceptibility to collagen-induced arthritis in mice lacking lFN—gamma
receptors. J Immunol 158:5501-5506

Gowen M, Mundy GR 1986 Actions of recombinant interleukin 1,
interleukin 2, and interferon-gamma on bone resorption in vitro. J Immunol
1 3622478-2482

Takayanagi H 2005 Mechanistic insight into osteoclast differentiation in
osteoimmunology. J Mol Med 832170-179

97

 

 

CHAPTER 3

Motyl KJ, McCabe LR 2009 Leptin treatment prevents type I diabetic marrow
adiposity but not bone loss in mice. J Cell Physiol 2182376-384

98

CHAPTER 3

3. LEPTIN TREATMENT PREVENTS TYPE I DIABETIC MARROW ADIPOSITY
BUT NOT BONE LOSS IN MICE

3.1. ABSTRACT

Leptin is a hormone secreted by adipocytes that is implicated in the
regulation of bone density. Serum leptin levels are decreased in rodent models of
type 1 (T1-) diabetes and in diabetic patients. Whether leptin mediates diabetic
bone changes is unclear. Therefore, we treated control and T1-diabetic mice with
chronic (28 day) subcutaneous infusion of leptin or saline to elucidate the
therapeutic potential of leptin for diabetic osteoporosis. Leptin prevented the
increase of marrow adipocytes and the increased aP2 expression that we
observed in vehicle-treated diabetic mice. However, leptin did not prevent T1-
diabetic decreases in trabecular bone volume fraction or bone mineral density in
tibia or vertebrae. Consistent with this finding, markers of bone formation
(osteocalcin RNA and serum levels) in diabetic mice were not restored to normal
levels with leptin treatment. Interestingly, markers of bone resorption (T RAP5
RNA and serum levels) were decreased in diabetic mice by leptin treatment. In
summary, we have demonstrated a link between low leptin levels in T1-diabetes
and marrow adiposity. However, leptin treatment alone was not successful in

preventing bone loss.

99

3.2. INTRODUCTION

Leptin is a small protein (16 kDa) secreted by adipocytes and involved in
bone mass regulation. The effects of leptin on bone are complex; it can stimulate
or inhibit bone formation depending upon bone' location and whether leptin is
functioning directly on osteoblasts (through receptors (1, 2)) or indirectly through
the hypothalamus (2-6). In children and during adolescence, decreases in serum
leptin levels, associated with reduced food intake and some disease conditions,
are thought to contribute to reduced bone formation and growth (3, 7). Absence
of leptin in mice also results in bone loss as well as increased bone marrow
adiposity (8, 9). Similarly, increased leptin levels, as observed in obesity (10),
are correlated with increased bone mass (11). Several studies have tested the
potential therapeutic benefits of leptin treatment on bone loss and marrow
adipocyte accumulation. In vitro studies demonstrate that leptin promotes bone
marrow stromal cells to exhibit an osteoblast rather than adipocyte phenotype (1,
2, 12). Consistent with this finding, subcutaneous infusion of leptin with osmotic
mini-pumps reduces marrow adiposity and increases bone mass in ob/ob mice
(13). Similarly, leptin treatment reduces bone loss from ovariectomy (14) and tail
suspension (15). In sum, these studies suggest the efficacy of leptin treatment to
restore bone density under conditions of bone loss.

Diabetes mellitus (types 1, 2 and gestational forms) is a metabolic disease
marked by hyperglycemia due to defective insulin action (lack of insulin or tissue

insulin resistance), which results in impaired glucose uptake by insulin

100

responsive cells. Type 1 (T1) diabetes is generally first diagnosed in adolescents
or young adults and affects nearly one million children and adults in the United
States. In T1-diabetes, insulin-secreting pancreatic B-cells are destroyed as a
result of autoimmunity. This causes severe hyperglycemia that must be
controlled by insulin delivery in humans. In addition to hyperglycemia, weight
loss can occur in patients that do not have an adequate insulin therapy. T2-
diabetes, generally diagnosed in adults, also results in hyperglycemia but insulin
levels, body weight, and serum lipids are increased. Whereas effects of T2-
diabetes on bone remain controversial due to increased fracture risk and
increased bone mineral density in these patients (16), it is recognized that
osteoporosis is a serious complication of T1-diabetes, leaving child and adult
patients at risk for bone loss, fracture and impaired fracture healing (17-20).

Previously we demonstrated that streptozotocin-induced T1 -diabetic mice
exhibit weight loss, decreased body fat mass and bone loss (21). Bone
formation by osteoblasts is decreased while resorption is unchanged or
decreased in diabetic humans and rodents (21-23). Interestingly, both STZ-
induced and spontaneous mouse models of T1-diabetes exhibit an increase in
bone marrow adiposity despite the loss of peripheral fat depots (21, 24, 25). A
reciprocal relationship between bone volume/density and marrow adipocyte
number (consistent with altered mesenchymal stem cell lineage selection) has
been implicated as a mechanism for diabetic (21, 24, 26, 27), age-related (28,
29), and unloading-induced (30) bone loss. However, while an increase in

adiposity was observed in tibia, femur and calvaria in T1-diabetic mice, vertebrae

101

did not exhibit an increase in adiposity (26). In this respect, the T1-diabetic bone
phenotype resembles that of leptin—deficient mice, which have increased marrow
adiposity and bone loss in the femur but no change in marrow adiposity in the
vertebrae (8). Because of this finding, we measured fed serum leptin and found
that it was decreased in both male and female T1—diabetic mice (26), consistent
with reports in STZ-diabetic rats (31).

These findings led us to hypothesize that decreased serum leptin levels
contribute to T1-diabetic bone loss and altered mesenchymal lineage selection.
Therefore we treated control and diabetic mice with leptin and examined bone
parameters. Consistent with our hypothesis, we observed reduced bone marrow

adiposity in leptin-treated diabetic mice; however, bone loss was not prevented.

3.3. MATERIALS AND METHODS

3.3.1. Animals

Forty 9-week old BALB/c mice were obtained from Harlan Sprague
Dawley (Indianapolis, IN). Mice were maintained on standard lab chow and had
food and water ad Iibitum. At 14 weeks, mice were divided into four treatment
groups: control + vehicle, control + leptin, diabetic + vehicle and diabetic + leptin.
Mice were anesthetized with isofluorane and implanted subcutaneously with
Alzet mini-osmotic pumps (model 2004, Durect Corporation, Cupertino, CA) that

contained either 0.9% sterile saline vehicle (n=20) or 1.3 mg/mL leptin (Amylin,

102

San Diego, CA) (n=20). Osmotic pumps had a mean pumping rate of 0.21 i 0.01
pL/hour and therefore delivered 6.6 pg leptin per day. Wounds were closed with
staples and mice were given a one-time injection of 0.15 mg carprofen (Pfizer,
New York, NY). Starting on the day of implantation (day 0), mice were given five
consecutive daily intraperitoneal injections of 50 mg/kg streptozotocin (n=24) or
0.1 M citrate buffer pH 4.5 vehicle (n=16). Body mass and food intake were
monitored throughout the experiment. Staples were removed on day 9.

On day 28, mice were fasted 2-3 hours prior to being euthanized. Blood
glucose was measured at the time of harvest with an AccuChek Compact
glucometer (Roche, Nutley, NJ). Blood was collected at the time of harvest,
allowed to rest at room temperature for five minutes, then centrifuged at 4000
rpm for ten minutes. Serum was removed and stored at -80°C and pellet
discarded. Tibias were removed and either fixed in 10% formalin or frozen in
liquid nitrogen and stored at -80°C. Femoral fat pads and tibialis anterior muscles
were removed and weighed, fixed and frozen. All animal procedures were

approved by the Michigan State University Institutional Animal Care and Use
Committee.
3.3.2. Serum measurements

Serum was stored at -80°C and put through no more than one freeze/thaw

CYcle. Leptin was measured using the Assay Designs Mouse Leptin Titer Zyme

kit (Ann Arbor, MI) according to the manufacturer protocol. Insulin was measured

103

using a Crystal Chem Inc. Ultra Rat Insulin ELISA kit (Downers Grove, IL)

according to the manufacturer protocol.

3.3.3. Bone Histology and Histomorphometry

Bones were fixed in 10% formalin and transferred to 70% EtOH after 24
hours. Fixed samples were processed on an automated Therrno Electron
Excelsior tissue processor for dehydration, clearing and infiltration using a routine
overnight processing schedule. Samples were then embedded in Surgipath
embedding paraffin on a Sakura Tissue Tek II embedding center. Paraffin blocks
were sectioned at 5 pm on a Reichert Jung 2030 rotary microtome. Slides were
stained with hematoxylin and eosin. Visible adipocytes, greater than 30 pm, were
counted in the tibia trabecular region ranging from the proximal growth plate to 2

mm away distally.

3.3.4. Micro Computed Tomography (pCT) Analyses

Fixed tibias were scanned using a GE Explore Locus pCT system at a
voxel resolution of 20 pm obtained from 720 views. Beam angle of increment was
0.5 and beam strength was set at 80 peak kV and 450 pA. Each run included
Control and diabetic, saline and leptin-treated bones and a calibration phantom to
standardize grayscale values and maintain consistency. Based on autothreshold

and isosurface analyses of multiple bone samples, a fixed threshold (1400) was

104

used to separate bone from bone marrow. Cortical bone analyses were made in
a defined 2 x 2 x 2 mm cube in the mid-diaphysis immediately proximal to the
distal tibial-fibular junction, with the exception of cortical bone mineral content
(BMC) and bone mineral density (BMD), which were made in a 0.1 x 0.1 x 0.1
mm cube. Trabecular bone analyses were done in a region of trabecular bone
defined at 0.17 mm (~1% of the total length) distal to the growth plate of the
proximal tibia extending 2 mm toward the diaphysis, and excluding the outer
cortical shell. Cortical BMC, BMD, moment of inertia (MOI), thickness, perimeter
and area, and trabecular BMC, BMD, volume fraction (BVF), and thickness

(T bTh) values were computed by a GE Healthcare MicroView software
application for visualization and analysis of volumetric image data. Cortical
isosurface images were taken from a section immediately proximal to the tibial-
fibularjunction measuring 0.3 mm thick. Trabecular isosurface images were
taken from a cylindrical region in the tibia immediately distal to the proximal

growth plate measuring 0.8 mm in length and 0.8 mm in diameter.

3.3.5. RNA Analyses

Immediately after euthanasia, tibias were cleaned of muscle and
connective tissue, snap frozen in liquid nitrogen and stored at -80 °C. Frozen
tibias were crushed under liquid nitrogen conditions with a Bessman Tissue
Pulverizer (Spectrum Laboratories, Inc., Rancho Dominguez, CA). RNA was

isolated with Tri Reagent (Molecular Research Center, Inc., Cincinnati. OH) and

105

integrity was assessed by formaldehyde-agarose gel electrophoresis. cDNA was
synthesized by reverse transcription with Superscript II Reverse Transcriptase Kit
and oligo dT(12-18) primers (Invitrogen, Carlsbad, CA) and amplified by real time
PCR with iQ SYBR Green Supermix (Biorad, Hercules, CA) and gene-specific
primers synthesized by Integrated DNA Technologies (Coralville, IA). HPRT
mRNA levels do not fluctuate in diabetes or with increased serum leptin and were
used as an internal control. HPRT was amplified using 5’-AAG CCT AAG ATG
AGC GCA AG-3’ and 5’-TTA CTA GGC AGA TGG CCA CA-3’ (32). aP2 was
amplified using 5'-GCG TGG AAT TCG ATG AAA TCA-3' and 5'-CCC GCC ATC
TAG GGT TAT GA-3' (33). Osteocalcin was amplified using 5'-ACG GTA TCA
CTA TTT AGG ACC TGT G-3' and 5'-ACT TTA TTT TGG AGC TGC TGT GAC-3'
(34). TRAP5 was amplified using 5'-AAT GCC TCG ACC TGG GA-3' and 5'-CGT
AGT CCT CCT TGG CTG CT-3' (35).

3.3.6. Statistical Analyses

All measurements are presented as the mean :1: standard error of the

mean (SEM). Statistical significance was determined with a student’s t-test

(assuming equal variance) using Microsoft Excel (Microsoft Corporation).

106

3.4. RESULTS
3.4.1. Serum leptin, glucose and insulin

To determine if correction of T1-diabetic serum leptin levels could prevent
diabetic bone loss and marrow adiposity, osmotic pumps containing either leptin
(delivering 6.6 pg leptin per day) or saline (vehicle) were implanted
subcutaneously in BALB/c mice and T1-diabetes was induced by 5 daily
injections of streptozotocin. Leptin treatment was successful in raising fasting
serum leptin levels in both control and diabetic mice (Table 1). Similar to our past
results, serum leptin levels in vehicle-treated diabetic mice trended to be lower
than vehicle-treated controls, however in this case the difference did not reach
statistical significance. This result is likely due to the variability in control mouse
leptin levels observed in this study and the difference in fasted (this study) versus
fed (past study) serum leptin measurements (36). Analysis of blood glucose
levels demonstrated that leptin treatment of control mice causes a decrease in
blood glucose levels compared to untreated controls, as previously shown by
others in rats (37, 38). As expected, fasting blood glucose levels were increased
in diabetic compared to control mice in both normal (501 vs 161 mgldl,
respectively) and high elptin conditions (250 vs 94 mgldl, respectively) (Table 1).
Although the blood glucose level in leptin-treated diabetic mice was significantly
lower (by 50%) than vehicle-treated diabetic mice, the fold increase in blood

glucose levels compared to corresponding treatment controls was similar

107

(untreated diabetic/untreated control = 3 fold; leptin-treated diabetic/Ieptin-treated
control = 2.7 fold). Differences in blood glucose levels between leptin-treated and
untreated mice may result from the reduced food intake seen in both control
leptin-treated and diabetic leptin-treated mice. Examination of fasting serum
insulin levels demonstrated that diabetic vehicle-treated mice have lower fasting
insulin levels compared to control vehicle-treated mice. Leptin-treated diabetic
mice exhibited lower serum insulin levels compared to untreated diabetic mice;
however, insulin levels were not lower than leptin-treated controls. This is likely
due to leptin treatment effectively reducing insulin levels in non-diabetic mice
(Table 1), and enhancing insulin sensitivity, thereby reducing glucose-stimulated

insulin release from the pancreas, as previously shown (39-41).

108

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3.4.2. Food consumption and body mass

Food intake and body mass were monitored throughout the experiment.
As expected, leptin treatment decreased food intake in control mice and
suppressed T1-diabetic hyperphagia compared to untreated diabetic mice
(Figure 13). The decrease in food intake corresponded to a decrease in body
mass (Figure 13) in control but not diabetic mice. Differences were apparent
throughout the time course beginning at diabetes confirmation (not shown).

To determine if the weight loss was the result of fat and/or muscle mass
loss, these parameters were also examined. As expected and consistent with
our past studies, diabetic vehicle-treated mice lost 50% of femoral fat pad mass
compared to vehicle-treated controls (Table 1). Even more fat pad mass was
lost in control leptin versus vehicle-treated mice (64%), consistent with previous
reports (13, 42). No differences were seen between leptin-treated diabetic,
vehicle-treated diabetic or leptin-treated control mouse fat pad mass. However,
after normalizing for body mass changes, diabetic leptin-treated mice had
significantly lower fat pad mass than diabetic vehicle-treated mice (3.6 :l: 0.3
mg/g versus 4.5 :1: 0.4 mg/g, respectively). Normalizing fat pad mass for body
mass did not alter the statistical comparisons in any other case. Similar to our
previous findings, diabetic mice had decreased tibialis anterior muscle mass
(Table 1). Leptin treatment did not affect muscle weight in control mice, but did
Prevent diabetic muscle mass loss compared to leptin-treated controls.

However, when normalized for body weight, no statistical differences were

110

apparent between groups, indicating that all muscle mass changes were directly

proportional to body weight changes.

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Figure 13. Leptin treatment prevented diabetic hyperphagia compared to
vehicle-treated mice, but did not prevent weightless. Food intake and body
mass from vehicle-treated (white bars) and leptin-treated (gray bars), control and
diabetic mice were monitored throughout the experiment. Shown are average
values obtained 28 days after diabetes induction. Values are representative of all
t"ne points after diabetes was induced. Bars represent mean 1 standard error.
"25 Per condition. *p<0.05 by Student’s t-test.

 

 

112

3.4.3. Diabetic marrow adiposity was prevented by leptin treatment

T1-diabetic bone loss is associated with increased marrow adiposity,
implicating altered lineage selection of bone marrow stromal cells as a potential
mechanism. Because leptin-treatment has been shown to prevent marrow
adipocyte accumulation in ob/ob mice (13), we hypothesized that it should
prevent marrow adiposity in diabetic mice. As expected based on previous
reports (21, 24), untreated diabetic tibias exhibited an increase in marrow
adipocyte number and aP2 mRNA levels (a marker of mature adipocytes; Figure
14). Consistent with our hypothesis, leptin treatment completely prevented
adipocyte accumulation and aP2 mRNA induction in diabetic bone marrow

(Figure 14).

113

Figure 14. Leptin treatment prevented T1 -diabetic marrow adiposity. (A)
Representative hematoxylin and eosin stained sections of decalcificed tibias from
vehicle- (white bars) or leptin-treated (gray bars), control and diabetic mice. (B)
Marrow adipocyte counts in the proximal tibia, immediately distal to the growth
plate. (C) aP2 gene expression was measured by RT-PCR in whole tibia cDNA
synthesized from mRNA. aP2 levels were expressed relative to HPRT, an
unmodulated housekeeping gene control. Bars represent mean a; standard error.
n26 per condition. *p<0.05 by Student’s t-test.

114

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3.4.4. Leptin did not prevent diabetic bone loss

Given the inverse relationship between bone density and adiposity in
diabetes, we examined leptin-treated diabetic bone to see if preventing adiposity
would also prevent bone loss. Because leptin can affect long bone and vertebrae
differently, we examined both the proximal tibia and L2 vertebrae trabecular bone
(Table 2). In control mice, leptin treatment tended to reduce all bone parameters
examined in both tibia and vertebrae. Significant reductions (compared to vehicle
treatment) were seen in bone volume fraction (BVF) and trabecular thickness in
tibia but not vertebrae, consistent with location-dependent leptin effects (5, 8).
However, leptin-treated control vertebrae exhibited significant decreases in bone
mineral content and density (BMC, BMD). As expected, untreated diabetic mice
exhibited a decrease in trabecular bone mineral content and density (BMC, '
BMD), bone volume fraction (BVF), and trabecular thickness (TbTh) compared to
untreated controls at both sites (tibia and vertebrae). These decreases were
more prominent than the changes seen in untreated versus leptin—treated
controls. In contrast to our original hypothesis, leptin treatment did not prevent
the reduction in diabetic trabecular BMC, BMD or BVF in tibia. In vertebrae, leptin
treatment did not prevent the diabetes-induced decrease in BMC or BVF. Leptin
treatment did significantly increase diabetic vertebral BMD compared to vehicle-

treated diabetic mice to the point of not being different from leptin-treated

116

controls. Because diabetic- and leptin-treated mice lose body weight, we
examined changes in BVF relative to total body mass. When corrected for body
mass, the leptin-treated control tibia bone volume fraction (BVF/g) was similar to
vehicle-treated controls (Figure 15), suggesting the reduction in bone volume is
consistent with reduced body size and load. However, bone loss (BVF/g) was still
evident in both untreated and leptin-treated diabetic groups (Figure 15).

We further examined cortical bone parameters in the tibia metaphysis
(Table 3). Diabetic mice had significantly decreased cortical thickness, which can
be attributed to greater inner cortical bone perimeter and increased marrow area.
Leptin-treatment (in both control and diabetic mice) also reduced cortical
thickness in a similar manner, suggesting the decreased cortical bone could be
due to weight changes. When corrected for differences in body mass, cortical

thickness was not statistically significant different between groups (Figure 16).

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Figure 15. Leptin did not prevent T1 -diabetic bone loss. Fixed tibias were
scanned by pCT and the trabecular area immediately distal to the proximal
growth plate was analyzed for bone volume fraction. Top: Representative
isosurface images of trabecular bone in vehicle- and leptin-treated mice, control
and diabetic mice. Bottom: Bone volume fraction (BVF) was corrected for body
mass. Bars represent mean 1 standard error. n26 per condition. *p<0.05 by

Student's t-test.

119

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Figure 16. Cortical bone thickness did not differ when corrected for body
mass changes. Top: representative isosurface images of cortical bone
transverse sections immediately proximal to the tibial-fibular junction. Bottom:
Cortical thickness expressed per gram body mass. Bars represent mean 2
standard error. n26 per condition. *p<0.05 by Student’s t-test.

122

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3.4.5. Leptin treatment suppressed bone resorption in diabetes

To understand how the bone changes observed in diabetes and leptin-
treatment occurred, we examined whole tibia gene expression and serum for
markers of bone formation and resorption (Figure 17). Osteocalcin mRNA, a
marker of bone formation, was significantly decreased in vehicle-treated diabetic
bone compared to vehicle-treated control. Leptin-treatment had no effect on
osteocalcin gene expression in control mice. Interestingly, osteocalcin trended to
decrease in leptin-treated diabetic compared to leptin-treated control mice,
although this changed did not reach statistical significance. Additionally, the level
of osteocalcin mRNA was not different between vehicle-treated diabetic mice and
leptin-treated diabetic mice. Both leptin groups tended to have higher serum
osteocalcin levels than vehicle-treated controls, although this difference was not
statistically significant.

Reports indicated that T1-diabetes generally decreases or does not affect
bone resorption. In this study, vehicle-treated diabetic mice had similar levels of
bone resorption as vehicle-treated control mice (marked by TRAP5 gene
expression and serum TRAP5b levels, Figure 17). We examined serum TRAP5
(versus urine measurements) because diabetic mice develop polyluria and can
exhibit nephropathy, both of which can confound urine measurements. Similarly,
TRAP5 mRNA and serum TRAP5b were not different in leptin-treated compared
to vehicle-treated controls. Diabetes did however decrease serum TRAP5b in

leptin-treated mice compared to leptin-treated controls. Interestingly, diabetic

124

leptin-treated mice had reduced TRAP5 expression and serum TRAP5b
compared to vehicle-treated diabetic mice, suggesting that leptin is capable of

suppressing resorption in diabetic mice.

125

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Figure 17. Leptin treatment suppressed bone resorption diabetes. (A)
Osteocalcin and TRAP5 gene expression was measured by RT-PCR in whole
tibia cDNA synthesized from mRNA and were expressed relative to HPRT, an
unmodulated housekeeping gene control. (B) Osteocalcin and active TRAP5b
were measured in serum collected at 28 days with commercially available ELISA
kits. Bars represent mean 1 standard error. n26 per condition. *p<0.05 by
Student's t-test.

126

3.5. DISCUSSION

We (and others) found serum leptin levels decreased in diabetic rodent
models (26, 31, 36, 38). Additionally, we demonstrated that T1 -diabetic mice
have a location-dependent marrow adiposity phenotype similar to that of leptin-
deficient ob/ob mice: increased marrow adiposity in long bones with diabetes, but
no marrow adipocyte accumulation in the vertebrae (8, 26). Altered
mesenchymal stem cell lineage selection is thought to be one mechanism for T1-
diabetic bone loss and increased marrow adiposity. We therefore hypothesized
that treating diabetic mice with leptin (through an osmotic pump) could prevent
bone loss and marrow adipocyte accumulation. We found that although leptin
administration completely prevented T1-diabetic marrow adiposity, it did not
prevent bone loss in tibia or vertebrae.

As noted previously, serum leptin levels have been demonstrated to be
reduced in diabetic rodent models (26, 31, 36, 38) and in diabetic patients (43,
44). In this experiment, we observed a trend toward decreased serum leptin
levels in diabetic compared to control mice, but it did not reach significance as
we had observed previously (26) (Table 1). In part, we observed increased
variability in our control population. In addition, the current study obtained leptin
levels from fasted mice, whereas our previous measurements were obtained
from fed mice. Akirav, et al. (36) also demonstrate significant decreases in fed
serum leptin levels in streptozotocin-induced diabetic compared to control rats,

but found no difference between fasting diabetic and control rats. The latter was

127

the result of reduced leptin levels (to that of diabetic rat levels) in fasted control
rats. Although leptin levels are often decreased in T1 -diabetic patients (43, 44),
fasting serum leptin levels were reported to be unchanged in T1-diabetic children
and adolescents aged 6-16 years (45). We expect that if we had isolated serum
from fed mice, leptin levels would have been decreased.

To our knowledge, this is the first report demonstrating a role leptin in the
accumulation of marrow adipocytes in T1 -diabetes. We found that by replacing
the leptin that is normally lost by peripheral fat Iipolysis in T1—diabetes, we can
prevent the increase in adiposity in the marrow that is associated with T1-
diabetes. It is possible that subcutaneous leptin administration prevented the
increased lipid accumulation and marrow adipocyte differentiation that occurs in
diabetes (12) or alternatively acted directly or via sympathetic nervous system
(SNS) stimulation on marrow adipocytes to induce apoptosis (46, 47). The latter
is supported by a report indicating that central leptin administration to the rat
ventromedial hypothalamus causes adipocyte apoptosis in fat pads and in bone
marrow (4).

Several reports (21, 29, 48-50) demonstrate increased marrow adiposity
with bone loss suggesting that (1) marrow fat accumulation could have negative
effects on bone formation, (2) mesenchymal stem cells preferentially choose the
adipocyte lineage instead of the osteoblast lineage in disease conditions, and/or
(3) trabecular bone volume decreases, therefore fat accumulates, filling in the
empty space it has left behind (28, 51-53). However, in this experiment inhibiting

marrow adiposity with leptin treatment did not prevent diabetes-induced

128

decreases in BMC, BMD and BVF in tibia trabecular bone (Table 2, Figure 15).
Consistent with this finding, diabetes induced changes in serum bone remodeling
markers were not prevented by leptin treatment in rats (54).

Our results also indicated that leptin treatment reduced tibia trabecular
bone mass and several vertebral bone parameters (BMC and BMD) in control
mice. Other parameters showed trends toward decreases but were not
significant. The suppression in bone parameters in control mice was somewhat
unexpected since previous reports of chronic leptin treatment in wild type mice
suggest that under normal conditions leptin does not affect bone mass (9) and/or
may increase bone strength (2). To our knowledge we are the first to examine
the influence of leptin treatment on BALB/c mouse bone density, which may
account for some of the differences between studies. Leptin treatment has been
predominantly examined in the C57BU6 strain since it is the background for
leptin deficient (ob/ob) and dysfunctional leptin receptor (db/db) genetic mouse
models. Although leptin treatment of wild type C57BU6 mice does not affect
most bone parameters, labeled bone forming surface have been reported to be
significantly decreased at 2.5 and 10 pg leptin/day doses (13). In that study, the
mice were treated for 14 days, so it may be that longer treatments would have
resulted in bone loss in control mice as we have seen witha 28 day treatment. It
is also important to note that the majority of studies using leptin as bone loss
therapeutic (ie: for ovariectomy and disuse (14, 15)) have used rats not mice,
suggesting that species differences must also be considered. Regarding gender,

key studies demonstrating Ieptin’s bone anabolic potential have used female rats

129

(unloading and ovariectomy (14, 15)) while both male and female ob/ob mouse
bones positively respond to leptin treatment (13, 55). Thus, gender is less likely
to play a role in our findings.

We hypothesized that leptin treatment of diabetic mice would prevent
bone loss based on the finding that subcutaneous infusion of leptin with osmotic
mini-pumps reduces marrow adiposity and restores bone mass in leptin deficient
ob/ob mice (13). Similarly, hypothalamic injection of leptin can also correct
skeletal abnormalities in ob/ob mice (55). Leptin injection into control (leptin-
replete) mice did not affect bone mass (13). This suggests that leptin deficiency
could be critical for determining leptin effectiveness. Our diabetic mice exhibit a
significant suppression in leptin levels although they are not completely deficient
as seen in ob/ob mice. It is possible that the low levels of endogenous leptin
affect adiposity but not bone loss. An alternative concept is based on the
comparative phenotypes between leptin deficient and T1-diabetic mice. While
leptin deficient mice lose long bone mass, they have increased vertebral
trabecular bone mass (5, 8, 9, 56-58); this is in contrast to T1-diabetic mice,
which lose bone at all sites examined to date. Thus, leptin may not be playing a
role in regulating bone mass in this disease model. However, the adiposity
phenotype, long bone but not vertebral adiposity, is identical to what we observe
in T1-diabetic mice and is restored with leptin treatment.

We did observe that leptin treatment increased serum osteocalcin levels
(although not statistically significantly) in both control and diabetic mice

suggesting a potential positive effect, however at the RNA level, the suppression

130

in osteocalcin levels by diabetes was not prevented with leptin treatment. Our
studies also demonstrate that leptin treatment significantly suppresses osteoclast
activity in diabetic mice as indicated by reduced active serum TRAP5b and bone
TRAP5 RNA levels. Previous reports suggest that leptin treatment does not
influence osteoclast activity (13) or can suppress activity possibly through
inducing OPG expression and suppressing RANK ligand (15).

Why was leptin unable to prevent bone loss in diabetes? We do not
believe there were any technical problems with leptin administration because of
its potent effect on food intake (Figure 13) and marrow adiposity (Figure 14) and
because leptin treatment was successful at raising serum leptin levels (Table 1).
Concentration could play a role based on recent studies indicating the efficacy of
leptin in restoring bone density is concentration dependent. Specifically, Martin
et al. (42) demonstrated in the tail-suspended disuse rat model that only low
dose leptin treatment (50 pglkg/day) was effective in preventing femur trabecular
bone loss. High dose leptin treatment (500 pglkg/day) reduced femur bone mass
in control rats and did not prevent bone loss in suspended groups. Either
treatment dose prevented marrow adiposity. Previous studies in rats have used
doses between ~100-350 pglkg/day with therapeutic success (14, 15). In ob/ob
mice, successful therapeutic doses have spanned 2.5 - 10 pg/day or ~100-400
pglkglday for control mice and ~66-263 pglkglday for leptin-deficient mice (13).
Our dose was intermediate (6.6 pglday; ~ 240 pglkg/day): lower than the high
dose in the Martin et al. study, but within the range to successfully treat bone loss

in ob/ob mice (13). While our results exhibited similarities to the high dose leptin

131

treatment response seen in tail-suspended rats (bone loss in all conditions and
adiposity prevention) (42), the high dose (but not the low dose) in that study
decreased serum osteocalcin, whereas we observed no significant change in
serum osteocalcin levels (Figure 17).

Interestingly, blood glucose levels were significantly reduced by leptin
treatment in both control and diabetic mice. This is consistent with a previous
report in which STZ-induced, diabetic mice overexpressing leptin became
hyperglycemic but were more sensitive to insulin and required lower doses
(compared to mice not overexpressing leptin) to maintain euglycemia (59). We
do not believe the lower blood glucose in the leptin-treated mice confounded our
results because we demonstrated bone loss to the same degree in leptin-treated
and vehicle-treated diabetic mice. We also found leptin treatment affected other
mouse parameters. For example, leptin-treated control mice exhibited lower
body weights than diabetic (vehicle- or leptin-treated) mice. Still, the diabetic
mice lost more bone (and more bone/gm body weight), supporting the notion that
diabetic weight loss cannot fully account for the bone loss. While leptin treatment
caused fat pad loss in control and diabetic mice, it prevented diabetic muscle
loss, suggesting that the leptin-treated diabetic mice are leaner compared to the
vehicle-treated diabetic mice. Similarly, the leptin-treated control mice were
leaner than vehicle-treated control mice. A trend toward reduced fat pad and
body mass was seen in wild type mice treated with 10 pg leptin/day although it

did not reach significance (13).

132

Although the reciprocal relationship of fat to bone is often observed in
diabetes, it is possible that they are not related to each other and occur through
two completely different mechanisms. It is also possible that leptin did inhibit
adipocyte differentiation (or only inhibited lipid deposition in adipocytes) and
other factors may be necessary to induce osteoblast differentiation. The latter is
likely, due to the complex pathology of T1-diabetes, which includes
hyperglycemia, hypoinsulinemia, low serum leptin, hyperlipidemia, and
inflammation. In a previous study, we demonstrated that inhibition of PPARyZ
with BADGE has a similar effect: prevention of marrow adiposity but not bone
loss in diabetes (60). In this case, BADGE prevented hyperlipidemia, suggesting
that excess lipids are not requisite for the process of bone loss in T1-diabetics.
We have also demonstrated that loss of insulin receptor signaling in bone does
not alter bone density, which suggests hypoinsulinemia cannot alone account for
bone loss in T1-diabetes (61). It is possible that other factors that have not yet
been addressed or a combination of T1-diabetic complications are important for
bone loss to occur.

In summary, we have demonstrated a link between low leptin levels in T1-
diabetes and marrow adiposity. Although leptin treatment prevented diabetes-
induced increased marrow adipocyte number and gene expression, leptin had no
effect on bone loss. We conclude, therefore, that it is unlikely that leptin
deficiency alone is responsible for diabetic bone loss and that other factors are

necessary for the pathogenesis of T1-diabetic osteoporosis.

133

3.6. ACKNOWLEGEMENTS

The authors thank Amylin for providing the leptin, Regina Irwin for
technical assistance and critically reviewing the manuscript and Lindsay Martin,
Laura Harris, Dennean Lippner and Erin Nekritz for critically reviewing the

manuscript.

134

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CHAPTER 4

4. CCAAT/ENHANCER BINDING PROTEIN BETA-DEFICIENCY ENHANCES
TYPE 1 DIABETIC BONE PHENOTYPE BY INCREASING MARROW

ADIPOSITY AND BONE RESORPTION
4.1. ABSTRACT

C/EBPB is an important regulator of adipocyte and osteoblast
differentiation from mesenchymal stem cells in bone marrow. Bone loss in type 1
diabetes is accompanied by increased marrow fat, which could directly contribute
to reduced osteoblast activity, or could result from altered lineage selection to
adipocytes rather than osteoblasts. Increased bone C/EBPB expression at the
onset of diabetes could push progenitors toward the adipocyte lineage. Here, we
induced diabetes in C/EBPfi-null (knockout, KO) mice to inhibit marrow adiposity
and prevent bone loss. Unexpectedly, we determined that ClEBPB-deficiency
actually enhanced the diabetic bone phenotype. While KO mice had reduced
peripheral fat depot mass compared to wild type, KO diabetic mice had 5-fold
more marrow adipocytes than diabetic wild type mice. Furthermore, tibia
trabecular bone volume fraction (BVF) loss was escalated, as diabetic KO mice
lost 48% of BVF compared to KO controls while littermate wild type diabetic mice
only lost 22% compared to wild type controls. The enhanced marrow adiposity in
KO diabetics could be attributed to compensation by C/EBP6, PPARyZ, and

CIEBPa. The osteoblast marker osteocalcin was reduced similarly in diabetic KO

142

and wild type mice. However, cathepsin K mRNA and osteoclast
histomorphometry indicated that resorption was increased in the KO diabetic
mice compared to KO controls, which could account for the greater decrease in
bone density in C/EBPB KOs. Taken together, ClEBPB-deficiency has
contrasting effects on diabetes-induced peripheral and bone marrow adipose
depot mass, while causing increased bone resorption, contrary to the classical

mechanism of type 1 diabetic bone loss.

4.2. INTRODUCTION

CCAAT/ enhancer binding protein beta (CIEBPB) is a member of the basic
region-leucine zipper (bZIP) class of transcription factors. A key regulator of
adipocyte lineage selection, it is transiently expressed during early adipocyte
differentiation with C/EBPa, and then followed by expression of peroxisome
proliferator-activated receptor gamma (PPARy) 2 and ClEBPot (1-3). CIEBPB
exists in three isoforms: transcriptionally active LAP-1 and LAP-2 and generally
inactive LIP (3). Overexpression of CIEBPB alone induces adipocyte
differentiation in NIH-3T3 fibroblasts, while LIP overexpression can prevent it (4,
5). Total deficiency of C/EBPB protects mice from obesity and reduces body fat
mass (6-8). Combined knockout of C/EBPB and C/EBP6 leads to an even greater
block to the adipocyte phenotype and adiposity (9).

In addition to its role in adipocyte maturation, C/EBPB has been shown to

affect osteoblast differentiation and subsequent bone density. These effects are

143

dependent on when (early vs. late differentiation), where (local vs. systemic), and
which (LAP vs. LIP, or both) C/EBPB is expressed. CIEBPB is normally
expressed during early and late stages of osteoblast differentiation, with
decreased expression during middle stages (10). Complete knockout of CIEBPB
in mice results in decreased total body mass and total bone mineral density
(BMD) (7). Similarly, targeted expression of the C/EBPB inactive form, LIP, to
preosteoblasts and osteoblasts results in osteopenia in transgenic mice due to
decreased bone formation (11).

Type 1 (T 1 -) diabetes affects approximately 1 million people in the United
States. This subtype of diabetes is characterized by a loss of insulin-secreting
pancreatic b-cells and hyperglycemia that must be controlled by insulin delivery
in humans. In addition to complications of retinopathy, neuropathy, nephropathy,
muscle atrophy and heart disease, T1-diabetes also causes osteoporosis (12-
14). Streptozotocin (STZ) is a pharmacologic agent used to induce pancreatic B-
cell toxicity and subsequent diabetes in research animals (15). Multiple low
doses of STZ (40 mg/kg injections for five consecutive days) induce
hyperglycemia in mice by five days after the first injection (DPI). In both STZ-
mice and rats, T1 -diabetes causes bone loss and impaired bone healing (16-21).
Similarly, spontaneously T1-diabetic NOD (non-obese diabetic) mice also lose
bone (22). Therefore, the observed bone changes are caused by diabetes and
not STZ alone.

Osteoporosis from T1 -diabetes is marked by decreased bone formation in

both humans and animals (16, 23, 24), while reported effects on osteoclast bone

144

resorption have been variable. Altered mesenchymal stem cell lineage selection
to adipocytes rather than osteoblasts has been implicated as a mechanism for
bone loss in diabetes (16, 22, 25), aging (26, 27), and unloading (28). In STZ-
diabetic mice, this hypothesis is supported by an increase in expression of
adipocyte-specific genes in bone (PPARy2 and fatty-acid binding protein (FABP)
4) and visible adipocytes in the marrow (16). Marrow adiposity is not a direct
result of STZ because NOD mice also have increased adipocyte accumulation
accompanying bone loss (22).

Inhibition of marrow adiposity is important for determining whether
adiposity causes and could be targeted to treat diabetic osteoporosis. Previously,
we demonstrated that treatment with the PPARyZ inhibitor, bisphenoI-A-diglycidyl
ether (BADGE), prevents STZ-diabetic marrow adiposity, but does not protect
against bone loss (29). One interpretation of this study is that mesenchymal stem
cells still differentiate toward the adipocyte lineage, but BADGE forces them to
remain in an early-adipocyte stage. Therefore, we hypothesized that inhibition of
adipocyte differentiation prior to PPARy2 induction (at the level of C/EBPB) could
be effective at enhancing or maintaining bone density in T1-diabetes.

In the present study, we demonstrate that CIEBPB expression is increased
in bone with the onset of diabetes. We hypothesize that CIEBPB may play a role
in the diabetic bone pathology by promoting adipocyte lineage selection and
increasing marrow adiposity. Therefore, deficiency of CIEBPB could inhibit T1-
diabetic marrow adiposity at the level of lineage selection and subsequently

prevent bone loss. To our surprise, we found that C/EBPB-deficiency does not

145

prevent the increased marrow adiposity or decreased osteoblast activity

classically observed in T1-diabetes. In fact, the absence of CIEBPB exacerbated

diabetic marrow adiposity, despite causing peripheral fat loss. Bone loss was

also increased with ClEBPB—deficiency; however, unlike bone loss in wild type
diabetic mice, additional bone loss in KO mice was due to enhanced bone

resorption rather than further suppression of osteoblast activity.
4.3. MATERIALS AND METHODS

4.3.1. Animals

BALB/c mice were obtained from Harlan Sprague Dawley (Indianapolis,
IN). Heterozygous C/EBPB (C/EBP/BH') mice were obtained from Peter F.

Johnson (NCI, Frederick, MD) (30) and were bred into the BALBlc strain and
genotyped at Michigan State University by Jeffrey Leipprandt in the laboratory of
Sandra 2. Haslam. All mice were maintained on a 12-hour light, 12-hour dark
cycle at 23 °C, were given standard lab chow, and had food and water ad Iibitum.
All animal procedures were approved by the Michigan State University

Institutional Animal Care and Use Committee. To induce diabetes, 14-week old
BALBIc mice, CIEBPB KO (C/EBPfi'l') mice, and wild type littermate controls

were injected with either 40 mg/kg STZ (Sigma, St. Louis, MO) or 0.1 M citrate
bUffer, pH 4.5, vehicle for 5 consecutive days. Diabetes was confirmed 12 days

Post-first STZ injection (DPI) using a drop of blood from the saphenous vein and

146

an Accu-Chek Compact glucometer (Roche Diagnostics Corporation,
Indianapolis, IN), with blood glucose greater than or equal to 300 mgldl indicating
diabetes. Body mass was monitored during diabetes induction and throughout
the experiment. BALBlc mice were euthanized at 5, 19, 28 and 40 DPI. C/EBPfi

KO mice and wild type littermate controls were euthanized at 40 DPI.

4.3.2. RNA Analyses

Immediately after euthanasia, tibias were cleansed of muscle and
connective tissue, snap frozen in liquid nitrogen, and stored at -80 °C. Frozen
tibias were crushed under liquid nitrogen conditions with a Bessman Tissue
Pulverizer (Spectrum Laboratories, Inc., Rancho Dominguez, CA). RNA was
isolated with Tri Reagent (Molecular Research Center, Inc., Cincinnati, OH) and
integrity was assessed by formaldehyde-agaroSe gel electrophoresis. cDNA was

synthesized by reverse transcription with Superscript II Reverse Transcriptase Kit

and oligo dT(12-13) primers (Invitrogen, Carlsbad, CA) and amplified by real time

PCR with iQ SYBR Green Supermix (Biorad, Hercules, CA) and gene-specific
primers synthesized by Integrated DNA Technologies (Coralville, IA).
Hypoxanthine guanine phosphoribosyl transferase (H PRT) mRNA levels do not
fluctuate in diabetes or deletion of C/EBPB, and were used as an internal control.
HPRT was amplified using 5’-AAG CCT AAG ATG AGC GCA AG-3' and 5’-'I'IA
CTA GGC AGA TGG CCA CA-3’ (31). C/EBPB was amplified using 5’-CAA GCT

GAG CGA CGA GTA CA—3’ and 5’-CAG CTG CTC CAC CTT CTT CT-3' (32).

147

FABP4 (aP2) was amplified using 5'-GCG TGG AAT TCG ATG AAA TCA-3' and
5'-CCC GCC ATC TAG GGT TAT GA—3' (33). C/EBP8 was amplified using 5’-
CGC AGA CAG TGG TGA GCT TG-3’ and 5’-CTT GCG CAC AGC GAT GTI'
GTT-3’ (32). PPARy2 was amplified using 5'-TGA AAC TCT GGG AGA TTC TCC
TG-3' and 5’-CCA TGG TAA TTI' C'I'l’ GTG AAG TGC-3' (34). CIEBPa was
amplified using 5’-GAA CAG CAA CGA GTA CCG GGT -3’ and 5’-GCC ATG
GCC 'I'I'G ACC AAG GAG-3’ (32). Osteocalcin was amplified using 5'-ACG GTA
TCA CTA T'I‘I' AGG ACC TGT G-3' and 5'-ACT TTA TTT TGG AGC TGC TGT
GAC-3' (35). Tartrate-resistant acid phosphatase (TRAP5) was amplified using
5'-AAT GCC TCG ACC TGG GA-3' and 5'-CGT AGT CCT CCT TGG CTG CT-3'
(36). Cathepsin K was amplified using 5'-GCA GAG GTG TGT ACT ATG-3' and
'-GCA GGC GTT GTT C'l‘l' ATT-3' (37). Amplicons were compared with a 100
bp DNA ladder (Invitrogen, Carlsbad, CA) on a 2% agarose gel in order to verify

RT—PCR results and to verify the absence of C/EBPB in the knockout animals.

4.3.3. Bone histology and histomorphometry

Femurs and tibias were fixed in 10% formalin and transferred to 70%
ethanol after 24 hours. Fixed samples were processed on an automated Thermo
Electron Excelsior tissue processor for dehydration, clearing, and infiltration using
a routine overnight processing schedule. Samples were then embedded in
Surgipath embedding paraffin on a Sakura Tissue Tek ll embedding center.

Paraffin blocks were sectioned at 5 pm on a Reichert Jung 2030 rotary

148

microtome. Slides were stained for TRAP activity and counter stained with
hematoxylin according to manufacturer protocol (387A-1 KT, Sigma, St. Louis,
MO). Osteoclast surface area was measured and expressed as a percentage of
total bone surface in the tibia trabecular region ranging from the proximal growth
plate to 2 mm distal. Visible adipocytes, greater than 30 pm, were counted in the
same region of the tibia and in the region of the femur immediately proximal to

the distal growth plate, extending 2 mm distal.

4.3.4. Micro Computed Tomography (pCT) Analyses

Fixed tibias were scanned using a GE Explore Locus pCT system at a
voxel resolution of 20 pm obtained from 720 views. Beam angle of increment was
0.5, and beam strength was set at 80 peak kV and 450 pA. Each run included
Control and diabetic, WT and CIEBPB KO bones, and a calibration phantom to
standardize grayscale values and maintain consistency. Based on autothreshold
and isosurface analyses of multiple bone samples, a fixed threshold (800) was
used to separate bone from bone marrow. Trabecular bone analyses were
performed in a region of trabecular bone defined at0.17 mm (~1% of the total
length) distal to the growth plate of the proximal tibia extending 2 mm toward the
diaphysis, and excluding the outer cortical shell. Trabecular bone mineral content
(BMC), bone mineral density (BMD), bone volume fraction (BVF), thickness
(TbTh), spacing (T bSp), and number (TbN) values were computed by a GE

Healthcare MicroView software application for visualization and analysis of

149

volumetric image data. Trabecular isosurface images were taken from a
cylindrical region in the tibia immediately distal to the proximal growth plate

measuring 1.0 mm in length and 1.0 mm in diameter.
4.3.5. Statistical Analyses

All measurements are presented as the mean :I: standard error of the
mean (SEM). Statistical significance was determined with a student’s t-test
(assuming equal variance) using Microsoft Excel (Microsoft Corporation,

Redmond, WA).
4.4. RESULTS

To determine the temporal pattern of ClEBPti expression in T1-diabetic
bone, we treated mice with STZ or vehicle to induce T1-diabetes and harvested
bone at 5, 19, 28 and 40 DPI. At 40 DPI, adipocyte numbers are clearly
increased in diabetic bone marrow (Figure 18A), consistent with previous reports
from our lab and others (16, 22, 29, 38, 39). However, at 5 DPI, the point at
which blood glucose levels become significantly elevated in diabetic mice (190 :t
12 mgldl versus 152 i 5 mgldl in controls), adiposity markers are already
beginning to increase at the RNA level (Figure 188) (40). Therefore, expression
of transient transcription factors involved in early adipogenesis should be evident

at this early time point. Consistent with adipocyte lineage selection in diabetic

150

bone, we observed an increase in CIEBPB mRNA levels at 5 DPI. This increase
was concurrent with an increase in aP2, a marker of mature adipocytes. The
latter remained elevated throughout the time course (Figure 183), as we have

previously demonstrated (40), while C/EBPB mRNA levels declined consistent

with adipocyte maturation (41).

151

 

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Figure 18. CIEBPB expression is increased in diabetic bone in conjunction
with aP2 expression and is followed by increased marrow adiposity. BALB/c
mice were injected with streptozotocin (STZ) to induce diabetes. Mice were
harvested 5, 19, 28 and 40 days post injection (DPI) with STZ. A: Fixed femurs
from diabetic mice at 5 and 40 DPI were stained with hematoxylin. Large white
circular features (denoted by arrows) are adipocytes. B: mRNA was extracted
from frozen tibias and made into cDNA, which was amplified by RT-PCR with
primers specific to C/EBPB, aP2 and HPRT (a housekeeping gene control).
Points represent mean :1: standard error of diabetic mRNA levels expressed
relative to control values (normalized to one). *p<0.05 by student’s t-test.

152

To determine whether CIEBPB plays an important role in diabetic bone
loss and marrow adiposity, we induced diabetes in C/EBPB KO mice and wild
type littermate controls. Mice were harvested at 40 DPI. Knockout of C/EBPB
was confirmed by RT-PCR in tibia (Figure 19). Consistent with previous studies,
the non-diabetic CIEBPB KO mice had lower glucose levels than non-diabetic
wild type mice (42). However, this hypoglycemia was not significant enough to
affect diabetes induction in KO mice. Blood glucose levels were increased to
greater than 500 mgldl in both wild type and CIEBPB KO diabetic mice (Figure

19), and were not significantly different from each other.

General mouse phenotype analyses indicate that C/EBPfi-l- mice are

smaller than wild type mice: they have lower total body, muscle and fat pad
mass, consistent with past studies (7, 42). Consistent with a diabetic phenotype,
both wild type and KO diabetic mice lost weight. However the KO diabetic mice
lost more weight: -16% versus -7% in diabetic wild type mice (Table 4). Body
weights of both control and diabetic KO mice were lower than treatment-matched
wild type mice. A portion of the diabetic weight loss can be accounted for by
muscle loss (-16% in wild type and -22% in KO mice) and peripheral fat loss (-
48%) in both wild type and KO mice (Table 4). Liver mass increased 28% in wild
type diabetic compared to control mice, whereas it increased only 10% in KO
diabetic compared to control mice (Table 4). Therefore, the greater body mass
loss induced by diabetes in the KO animals could be attributed to increased

muscle loss and less liver hypertrophy.

153

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Figure 19. Diabetes induction did not differ between CIEBPB knockout and
wild type mice. LEFT: To confirm genotypes, mRNA was extracted from frozen
tibias and was made into cDNA with a reverse transcriptase reaction. cDNA was
amplified by RT-PCR with primers specific to C/EBPB and values are expressed
relative to HPRT, a housekeeping gene control. Amplicons were separated on
1.5% agarose DNA gel and visualized by ethidium bromide staining. Absence of
a C/EBPB-specific amplicon was indicative of CIEBPB knockout. RIGHT: Blood
glucose was measured in control (C, white bars), diabetic (D, gray bars), wild
type (WT) and C/EBPfl' mice at 40 DPI. Bars represent mean :I: standard error.
*p<0.05 by student’s t-test. N 2 5 per condition.

154

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155

To determine whether ClEBPB-deficiency prevented adipocyte
accumulation in the diabetic marrow, we quantitated adipocytes in hematoxylin
stained tibia sections (Figure 20A and 203). Marrow adipocyte number did not
differ significantly between KO and wild type mice, although KO mice tended to
have more adipocytes (Figure 20B). Diabetes increased adipocyte numbers in
both wild type and KO mice compared to corresponding controls. Interestingly,
marrow adipocyte accumulation was enhanced by the lack of C/EBPB; KO
diabetic mice had over 5-fold more adipocytes than wild type diabetic mice.
Similar to adipocyte number, aP2 gene expression increased in wild type diabetic

mice compared to wild type control mice (Figure 20C), which is consistent with

previous findings (16, 29, 38, 40, 43). C/EBPB'I' control mice also had

significantly lower aP2 expression than wild type control mice. Despite this.
diabetes induced aP2 expression in KO mice to levelssignificantly higher than all

other groups.

156

Figure 20. Diabetic marrow adiposity was increased by CIEBPB deficiency.
A: Represenative photomicrographs of tibia sections stained with hematoxylin
from control (C) and diabetic (D) wild type and C/EBPH" tibias. Photographs
were taken 1 mm distal to the proximal growth plate. B: Visible adipocytes were
counted in the marrow portion of tibia sections in the area 2 mm distal to the
proximal growth plate. C: mRNA was extracted from frozen tibias and was made
into cDNA with a reverse transcriptase reaction. cDNA was amplified by RT-PCR
with primers specific to aP2, an adipocyte marker, and values were expressed
relative to HPRT, a housekeeping gene control. Bars represent mean 1 standard
error of control (C, white bars), diabetic (D, gray bars), wild type and ClEBPa"'
mice. *p<0.05 by student’s t-test. N 2 5 per condition.

157

 

 

 

 

 

 

 

 

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In order to understand how deletion of CIEBPB, a transcription factor
important for adipocyte differentiation, could enhance diabetic marrow adiposity,
we measured other adipogenic transcription factors in both wild type and KO,
control and diabetic mice (Figure 21). C/EBP8, which is normally expressed with
CIEBPB early in adipocyte differentiation, was significantly decreased (nearly 5-

fold) in diabetic wild type compared to diabetic control bone. This decrease was

prevented in the C/EBPfi'l' mice. PPARy2 and C/EBPa, both transcription factors

present later in adipocyte differentiation, were unchanged in diabetic wild type
mice compared to wild type controls, but were significantly elevated in KO
diabetic mice compared to KO controls. Heightened adipogenic transcription
factor expression in KO diabetic bone (Figure 21) is consistent with the higher

marrow adiposity that we observed (Figure 20).

159

 

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Figure 21. Adipogenic transcription factors are elevated in diabetic CIEBPB’
" compared to wild type diabetic bone. mRNA was extracted from frozen tibias
and was made into cDNA with a reverse transcriptase reaction. cDNA was
amplified by RT-PCR with primers specific to CIEBP6, PPARy2 and CIEBPa.
Diabetic levels were expressed relative to controls for the corresponding
genotype. Bars represent mean :I: standard error of diabetic wild type (gray bars)
and diabetic C/EBPH” (white bars) bone. *p<0.05 compared to genotype-
matched control by student’s t-test. N 2 5 per condition.

160

Next, we examined bone density parameters in the tibia of these mice to
determine if C/EBPB-deficiency had an impact on the diabetic bone phenotype.
We scanned tibias with a pCT and examined the trabecular bone immediately
distal to the proximal growth plate (Table 5 and Figure 22A). Control C/EBPB KO
mice had significantly lower BMC, BMD, BVF, and Tb.N. than control wild type
mice, similar to recent findings (7, 44). Diabetes caused trabecular bone loss in
both wild type and KO mice. Specifically, trabecular BMC, BMD, BVF and Tb.Th.

were lower (Table 5). Thus, diabetic bone loss was not prevented by CIEBPB-

deficiency. The amount of loss, however, was greater in diabetic C/EBPfi'l' mice

compared to diabetic wild type mice. Specifically, BMD was further decreased (3
reduction of 66 mg/cc versus 39 mglcc, 31% versus 15% in KO versus wild type
mice, respectively). Diabetes also caused BMC to be reduced 31% in K0 and
only 15% in wild type mice. BVF loss was 48% in diabetic KO mice, while only
22% in diabetic wild type mice (Table 5, Figure 22A and 228). Trabecular
thickness decreased 38% in diabetic KO mice versus 20% in diabetic wild type
mice. Trabecular spacing trended to be higher (p<0.1) in the diabetic KO mice,
but it did not change in the diabetic wild type mice. Similarly, although trabecular
number was unchanged in the diabetic wild type mice compared to control wild
type mice, it was significantly lower (27% decrease) in the diabetic KO mice

compared to control KO mice (Table 5).

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Figure 22. Absence of CIEBPfi exacerbated T1-diabetic bone loss without
altering osteocalcin expression. A: Representative 3-dimensional isosurface
pCT images were taken in a cylindrical region of interest (1 mm diameter x 1 mm
length) immediately distal to the proximal growth plate of the tibia. B: Bone
volume fraction was measured by pCT in fixed tibias immediately distal to the
proximal growth plate. C: mRNA was extracted from frozen tibias and was made
into cDNA with a reverse transcriptase reaction. cDNA was amplified by RT-PCR
with primers specific to osteocalcin (an osteoblast marker) and values expressed
relative to HPRT, a housekeeping gene control. Bars represent mean :l: standard
error of control (C, white bars), diabetic (D, gray bars) wild type and C/EBPfl”
mice. *p<0.05 by student’s t-test. N 2 5 per condition.

163

 

 

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164

To determine if C/EBPB deficiency affects diabetes-induced changes in
osteoblast and osteoclast gene markers, we examined expression of osteocalcin
and TRAP5, respectively. Tibia expression of osteocalcin was decreased 26% in
diabetic wild type mice compared to control wild type mice, as we have

demonstrated previously (16, 38, 40), indicating decreased osteoblast activity

with diabetes (Figure 22C). Although control C/EBPfi'l' mice had less bone than
control wild type mice, osteocalcin expression was unchanged. Diabetic C/EBPfl'
/' mice also had a decrease in osteocalcin expression (43%) compared to control

C/EBPfi-fi mice, however diabetic KO mice levels were not significantly lower

than those of diabetic wild type mice. Since knockout of CIEBPB did not alter
diabetic changes in osteocalcin expression, we examined osteoclast parameters
to determine if resorption was increased. Tibia RNA levels of TRAP5 and
cathepsin K, markers of osteoclast activity, were unchanged in diabetic wild type

mice compared to wild type controls (Figure 23A), which is similar to our previous
findings (16, 40)' TRAP5 expression was 1.5-fold higher in control C/EBPfl'l'
mice compared to wild type controls (while cathepsin K expression was
unchanged), indicating a potential role for increased resorption in C/EBPfl'l'

bone. Similar to wild type mice, TRAP5 expression was unchanged but trended
to decrease (p = 0.07) in diabetic KO bone. In contrast, cathepsin K mRNA was
elevated 2-fold in diabetic KO mice compared to control KO mice. In order to

account for the enhanced bone loss and clarify the status of bone resorption in

165

the diabetic KO mice, acid phosphatase-positive osteoclasts were measured
(Figure 238). We determined that osteoclast surface area (when expressed as a
percent of total surface area) was 73% higher in diabetic KO mice than control

KO mice. This is consistent with elevated cathepsin K mRNA levels and thus we

concluded that the enhanced bone loss in diabetic C/EBPfiJ' mice is likely due to

increased resorption.

166

Figure 23. CIEBPB knockout causes increased bone resorption in type 1
diabetes. A: mRNA was extracted from frozen tibias and was made into cDNA
with a reverse transcriptase reaction. cDNA was amplified by RT—PCR with
primers specific to TRAP5 and cathepsin K (osteoclast markers) and expressed
relative to HPRT, a housekeeping gene control. B: Decalcified tibia sections were
stained for TRAP activity in order to identify osteoclasts. Trabecular bone surface
in contact with osteoclasts was measured and expressed relative to the total
bone surface examined. Bars represent mean 1: standard error of control (C,
white bars), diabetic (D, gray bars) wild type and C/EBPfi" mice. *p<0.05 by
student’s t-test. N 2 5 per condition.

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4.5. DISCUSSION

T1-diabetes results in decreased bone formation and increased marrow
adiposity, which could be caused by mesenchymal stem cell lineage selection
preference for the adipocyte rather than the osteoblast lineage (16, 45). The
initial aim of this study was to determine if absence of CIEBPB could prevent the

altered lineage selection, thus preventing T1-diabetic bone marrow adipocyte

accumulation and bone loss. Rather, we found the opposite effect: C/EBPfi'/'

mice have enhanced bone marrow adiposity and bone loss in response to T1-
diabetes.
Although many studies have demonstrated the importance of CIEBPB for

adipocyte differentiation in vitro (1, 2, 4, 5), and have demonstrated reduced in

vivo peripheral adiposity in C/EBPfi'l' mice (6-8), bone marrow adiposity has not

been extensively examined. Here, we find no significant change in marrow

adipocyte numbers between wild type and C/EBPfi'l' mice (Figure 20), which is

consistent with a recent study (44). Expression of the adipocyte marker, aP2, is
slightly, but significantly, decreased in KO control mice compared to control wild
type mice (Figure 20C). Contrary to our hypothesis, the loss of CIEBPB
enhanced T1-diabetic bonemarrow adiposity (Figure 20). This could be
explained by altered expression of other adipogenic transcription factors. We

observed the maintenance of CIEBP6 expression in diabetic CIEBPB KO mice

169

(Figure 21) and this may be sufficient for the modest increase in CIEBPa and
PPARyZ expression that we observed (Figure 21). CIEBP?» expression has
previously been reported to be lFN-y/LPS inducible in CIEBPfi KO, but not wild
type, macrophages (46). We have previously reported inflammatory effects of T1-
diabetes in bone (40). While most inflammatory effects were observed at diabetic
onset, elevated IL-1a expression was observed at later times. lL-1 is known to
stimulate C/EBP6 expression (47-51), as well as enhance its activity (52). As
both ClEBPa (53) and PPARy2 (54) transcription can be directly regulated
through CIEBP transcription factors, a chain of events where lL-1 stimulates
CIEBP6 expression and activity, which in turn stimulates C/EBPa and PPARyZ
expression to promote adipogenesis is plausible in the diabetic CIEBPB KO
animals.

Despite the enhanced marrow adiposity, C/EBPB diabetic KO mice lost
the same percentage of peripheral fat as diabetic wild types (Table 4). Bone ‘
marrow adipocytes and peripheral adipose depots often have reciprocal
phenotypes. T1 -diabetes generally induces a loss of peripheral and visceral fat,
but this is accompanied by an increase of fat in the bone marrow (16). This
reciprocal relationship between peripheral/visceral fat and marrow adiposity is
also present in models of alcohol consumption (55) and aging (56). Here,
however, the enhanced adiposity of the bone marrow in the diabetic knockout
mice was not accompanied by enhanced peripheral fat loss, suggesting
independent regulation of the two depots in the C/EBPB KO mice. This idea is

also supported by the fact that control CIEBPB KO mice have less peripheral fat

170

with no significant change in bone marrow adiposity compared to their wild type
counterparts. Perhaps under normal conditions, ClEBPfi plays an important role
in adipose deposition in the periphery, but not in the marrow. In contrast, our
study indicates that C/EBPB is important in regulating diabetic marrow fat
accumulation, while having no effect on diabetic peripheral fat loss.

It is important to address the fact that we do not observe elevated PPARyZ
expression in diabetic wild type bone compared to control wild type bone (Figure
21), as we have seen in our past studies (16). Despite this discrepancy in
PPARy2 expression, we do observe increased marrow adiposity and aP2 mRNA
levels. It is possible that adiposity may be dependent upon the severity of
diabetes-induction, which can vary between experiments using streptozotocin
(43), and that more severe diabetes would produce detectable increases in
PPARyZ expression concurrent with aP2 and marrow adiposity.

Upon examination of the bone phenotype of C/EBPB KO mice, we found
that KO mice had significantly lower bone density. However, absence of CIEBPB
did not alter osteoblast parameters in control mice. This is consistent with a

recent study indicating bone loss, but unchanged bone formation parameters

(osteoblast surface and osteoblast number) in femurs of 12-week old C/EBP/3'/'

mice compared to wild type littermates (44). However, another study found

reduced mineral apposition rate (MAR) and bone formation rate/bone surface

(BFR/BS) in 8-week old C/EBPHl' mice suggesting CIEBPB-deficiency

suppresses osteoblast function (57). As our study was started in mice that were

14-weeks old, this apparent discrepancy may be due to the age of the mice, with

171

C/EBPB being an essential regulator of osteoblasts during development (57, 58),
but not during adult remodeling (44). Our finding that diabetic suppression of
osteocalcin expression is not altered in the knockout mice indicates that factors
other than CIEBPfi alone are necessary for reduced osteoblast activity in T1-
diabetes.

Because bone formation markers were unchanged by ClEBPB-deficiency,
we examined bone resorption parameters (TRAP5, cathepsin K and osteoclast
surface). Recent studies have demonstrated that loss of CIEBP?) enhances
osteoclast differentiation, likely through absence of LAP-induced suppression of

MafB (57). That study demonstrated increased expression of TRAP5 and

cathepsin K in C/EBPfi'l- osteoclasts in vitro, and increased TRAP staining in

vivo (specifically, larger osteoclasts were observed, with no change in osteoclast
number in 8-week old knockout mice) (57). Here, we demonstrate increased .
TRAP5 expression with C/EBPB-deficiency in vivo (Figure 23A), however we did
not observe any change in cathepsin K (Figure 23A), percentage osteoclast
surface (Figure 238), or in osteoclast number (not shown) in control KO bone
compared to control wild type bone. Another study using 12-week old wild type
and knockout mice had a similar result (no change in osteoclast parameters), but
did not examine serum or RNA markers of resorption (44). Upon induction of
diabetes, we observed no change in osteoclast parameters (TRAP5 and
cathepsin K mRNA, percentage 0C surface) in wild type mice (consistent with
previous studies), but found increases in cathepsin K expression and in

percentage 0C surface in diabetic KO mice compared to control KO mice

172

(Figure 23). TRAP5 expression, however, tended to decrease, but this was not
statistically significant. Taken together, our osteoclast data indicates that C/EBPB
may be preventing increased resorption from occurring in T1-diabetic wild type
bone.

Because C/EBPB-deficiency was unable to prevent diabetic adipocyte
accumulation, we cannot draw conclusions from the present study as to whether
altered lineage selection is indeed a mechanism of diabetic bone loss. Our
previous studies that have prevented diabetic marrow adiposity, but not bone
loss (with BADGE and leptin treatment), suggest altered lineage selection is not
the only contributing factor to decreased osteoblast activity in diabetes (29, 38).
Additionally, we found that more severe diabetes (induced by increased STZ
dose) increases marrow adiposity, while leaving bone volume fraction unchanged
(compared to normal STZ doses), which also suggests that the two may not be
linked in T1-diabetes (43). Certainly, in this study, we did observe that more

marrow adiposity was associated with more bone loss. However, our data

suggest that the bone loss in the C/EBPfi'l' mice occurred through increased

osteoclast activity, rather than suppressed bone formation. It is possible that the

enhanced osteoclast activity is a direct result of the absence of CIEBPB in
osteoclasts, or that it is secondary to the increased marrow fat because
adipocytes can secrete factors like TN F-a that induce osteoclastogenesis (59,
60).

In summary, we demonstrated that the absence of CIEBPB enhanced the

diabetic bone phenotype. Contrary to our original hypothesis, absence of CIEBPB

173

actually enhanced diabetes-induced expression of PPARyZ and CIEBPa, and
subsequent marrow adiposity. However, unlike marrow fat, we have shown that
diabetic peripheral fat changes are not dependent on C/EBPB, which speaks to
the complexity and location dependence of fat depots. Increased marrow
adiposity was concurrent with reduced bone density, but osteoblast activity
markers were not further suppressed. In fact, osteoclast activity was increased in
diabetic CIEBPB KO mice, which is contrary to the classic mechanism of bone
loss from T1-diabetes. Our findings add to the work of others that recently have
suggested a role for CIEBPB in the inhibition of osteoclast differentiation. Finally,
we conclude that C/EBPB alone is not responsible for the bone versus fat
phenotype switch observed in T1-diabetes, and that therapeutic treatments

suppressing its levels may further bone loss by increasing bone resorption.

4.6. ACKNOWLEDGEMENTS

The authors thank Pete Johnson (NCI) for providing breeder C/EBPfiJ'
mice, Sandra Haslam and Jeffery Leipprandt (MSU) for breeding, genotyping and
backcrossing the C/EBPfiJ- mice into the BALB/c strain, Regina Irwin for

technical expertise and critical review of the manuscript, and Lindsay Martin for
critical review of the manuscript. This work was funded by grants from the
National Institutes of Health (RO1DK061184) and the American Diabetes

Association (7-07-RA-105) to LRM. The authors have no financial conflicts.

174

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CHAPTER 5

5. MICE WITH OVEREXPRESSION OF WNT1OB ARE RESISTANT TO BONE
LOSS BUT NOT MARROW ADIPOSITY FROM TYPE 1 DIABETES

5.1 . ABSTRACT

Osteoporosis is a severe complication of type 1 diabetes, leaving patients
and at risk for fracture and impaired bone healing. Bone loss from diabetes is
primarily due to an osteoblast defect with no change or reduced osteoclast
activity, and is accompanied by increased marrow fat. Increased osteoblast
apoptosis and/or reduced osteoblast differentiation and maturation are likely
involved in the diabetic bone pathology. Reduced osteoblast differentiation may
be coupled to increased marrow adiposity because osteoblasts and adipocytes
arise from the same precursor: mesenchymal stem cells. Wnt/B—catenin signaling
is a potent regulator of lineage selection: it induces osteoblast differentiation and
prevents adipocyte differentiation. Here, we examined bone expression levels of
Wnt10b, which causes increased bone mass when overexpressed and reduced
bone mass when deleted. We found that in diabetes and leptin treatment (which
also causes bone loss itself and does not prevent diabetic bone loss) Wnt10b
levels were suppressed. Alternately, PTH treatment, which increases bone
formation (even in diabetic animals) promoted Wnt10b expression. We therefore

examined the ability of overexpression of Wnt10b itself to prevent diabetic bone

182

changes. Interestingly, overexpression of Wnt10b in osteoblasts (from the
osteocalcin promoter, OC-Wnt10b) prevented femur trabecular bone density
changes induced by diabetes, suggesting lower Wnt10b in diabetic bone levels
may be responsible for diabetic osteoporosis. However, diabetic OC-Wnt10b
mice were not protected from diabetic marrow adiposity, which could be due to
the localization of Wnt10b even though it is a secreted protein. Despite the
prevention of bone loss, we still observed suppressed osteoblast markers (serum
and mRNA) in the diabetic OC-Wnt10b mice compared to untreated OC-Wnt10b
mice. This suggests that bone loss may still occur in the OC-Wnt10b mice but at
a much slower rate that was not detectable after 40 days. Nonetheless, we
advocate that agonizing the Wnt10blB-catenin signaling pathway is a worthwhile

therapeutic target for osteoporosis from type 1 diabetes.

5.2. INTRODUCTION

Type 1 diabetes (T1-diabetes) is a chronic condition in which the pancreas
ceases to produce insulin, and the resulting hyperglycemia must be controlled

with insulin injections. Despite well-controlled blood glucose (as determined by

glycated hemoglobin levels, HbA1c), many patients do have complications from
T1-diabetes, including osteoporosis. Young people diagnosed with T1-diabetes
have reduced growth, which correlates with the level of glycemic control (1, 2). In

addition to affecting adolescents, diabetes can cause bone loss at all stages of

life, leaving patients at greater risk for bone loss, fracture and impaired fracture

183

healing (3-5). Serum osteocalcin, a marker of bone formation (osteoblast activity)
is significantly reduced in diabetic patients of all ages (6). Additionally, diabetic
patients have unchanged or decreased markers of resorption, suggesting that
increased osteoclast activity is not responsible for bone loss (7).

Rodent models of T1 -diabetes have a bone phenotype similar to that of
humans. One pharmacologic agent used to induce diabetes in mice is
streptozotocin (STZ). STZ is a glucose mimetic that diffuses into insulin secreting
pancreatic B-cells through the non-insulin responsive glucose transporter GLUT2
(8). The effects of STZ include DNA damage and sUbsequent B-cell death (8). In
both STZ-mice and rats, T1-diabetes causes bone loss and impaired bone
healing (9-15). Non-obese diabetic (NOD) mice are susceptible to development
of autoimmune T1-diabetes and display a similar bone phenotype to that of STZ-
diabetic mice (16). Spontaneously diabetic BioBreeding (BB) rats also have bone
loss due to decreased bone turnover (17, 18).

In addition to reduced bone formation and unchanged or reduced
resorption in type 1 diabetic bone, STZ and NOD mice have an obvious increase
in bone marrow adipocyte number (9, 16, 19-23). STZ mice have increased
expression of peroxisome proliferator-activated receptor gamma (PPARy) 2 in
bone, a transcription factor important for adipocyte maturation (9). Because
adipocytes are derived from the same stem cells that give rise to osteoblasts, the
increased marrow adiposity suggests that differentiation of MSCs may be
shunted away from the osteoblast lineage and toward the adipocyte lineage

under diabetic conditions (9). It is also possible that adipocytes secrete factors

184

(i.e. tumor necrosis factor alpha (TNFa)) into the marrow microenvironment that
reduce osteoblast differentiation or increase osteoblast apoptosis. Similarly, high
marrow fat has been demonstrated in bone loss models of aging (24, 25) and
unloading (26).

Several factors regulate both osteoblast and adipocyte lineage selection
and could be involved in diabetic bone loss. We have demonstrated, however,
that inhibition of PPARy with bisphenol-A-diglycidyl ether (BADGE) in control and
diabetic mice reduced diabetic marrow adiposity but did not prevent reduced
bone formation (osteocalcin and runx2 expression) or reduced bone density from
T1-diabetes (19). We have also prevented diabetic marrow adiposity with leptin
treatment, but did not prevent bone loss (22). Both of these studies suggest that
mature adipocytes in the bone marrow are not responsible for diabetic bone loss.
It remains plausible, however, that lineage selection still favors the adipocyte
lineage in these experiments, but that BADGE and leptin prevent maturation.
Therefore, examining the effect of other osteoblast/adipocyte lineage regulators
may provide insight into the mechanism of the diabetic bone phenotype.

Wnt/B-catenin signaling through TCFILEF-induced transcription is a potent
regulator of both bone formation and adipocyte differentiation (27, 28). Wnts are
secreted ligands that bind low density Iipoprotein receptor-related protein 5/6
(LRP5/6) and frizzled (de) membrane receptors. Dimerization of the LRP and
de receptors in response to wnts initializes a signaling cascade that inhibits
glycogen synthase kinase 3 beta (GSK3I3). Without the wnt signal (or in the

presence of endogenous pathway inhibitors such as dickkopf (Dkk) proteins),

185

GSK3B actively phosphorylates cytoplasmic B-catenin, targeting it for
degradation. When GSK3B is inactive (as in the presence of a wnt ligand),
transcriptionally active B-catenin (dephosphorylated on serine (Ser) 37 or
threonine (T hr) 41) accumulates in the cytoplasm and translocates to the nucleus
where it initiates transcription of genes with TCF/LEF binding sites, such as
Runx2 (29-31).

Mice null for LRP5 have low bone density and blindness, consistent with
the rare recessive disorder, osteoporosis-pseudoglioma syndrome (OPPG), while
gain of function of LRP5 results in high bone mass due to increased osteoblast
activity (32-37). Although complete loss of function of LRP6 is fatal, loss of one
allele causes more bone loss in mice deficient in LRP5 by exacerbating reduced
bone formation (38). Similarly, modulation of Wnt10b affects only bone formation,
not resorption (39). Wnt10b knockout (-KO) mice have low bone density while
mice with overexpression of Wnt10b from the aP2 promoter have high bone
density and low marrow adiposity (39). These mice also have attenuated bone
loss from aging and ovariectomy (39). In vitro activation and inhibition of wnt
signaling results in inhibition and activation of adipogenesis, respectively (40,
41), through regulation of expression of PPARy and CIEBPo (42, 43). Wnt1, 5a
and 7b have also been shown to promote osteoblast and/or suppress adipocyte
differentiation (38). Endogenous agonists and antagonists can further regulate
wnt signaling. Dally protein enhances the interaction between wnts and des,
while Dkks and secreted frizzled-related proteins (sFRP) antagonize it.

Additionally, sclerostin, prevents the de-LRP interaction. Targeted inhibition of

186

wnt agonists and antagonists is a relatively new and promising area of interest
for treatment of osteosclerotic and osteoporotic diseases, respectively (28).

We therefore examined wnt pathway family member changes in diabetic
bone and found that Wnt10b mRNA levels were significantly decreased. Because
Wnt10b strongly promotes bone formation and inhibits adipogenesis (39, 44, 45),
we also examined its levels with leptin treatment (which causes bone loss and
prevents diabetic marrow adiposity) and PTH treatment (which promotes bone
formation, but does not alter marrow adiposity in control or diabetic mice)
(22)(Motyl, et al., in preparation, See Chapter 6). Consistent with the known
effects of Wnt10b on bone density, we found mRNA levels decreased with leptin
treatment and increased with PTH treatment (46). Therefore, we examined the
effect of overexpression of Wnt10b itself on diabetic bone loss and marrow
adiposity. Briefly, we determined that Wnt10b prevented trabecular bone
phenotype changes in diabetes, but did not prevent the reduction cf serum or
mRNA osteoblast markers, suggesting bone loss may still occur, albeit at a
slower rate. Consistent with previous reports, we found no changes in osteoclast
surface measurements in any of the treatment groups. Interestingly, Wnt10b did
not prevent adipocyte accumulation in the diabetic bone marrow. We concluded
that because of the ability of Wnt10b to retard diabetic bone loss, Wnt10b/B-
catenin signaling should be further explored as a therapeutic target for patients

with T1-diabetic osteoporosis.

5.3. MATERIALS AND METHODS

187

5.3.1. Leptin and PTH treatment

All animal procedures were performed in accordance with Michigan State
University Institutional Animal Care and Use Committee. Leptin treatment was
performed as described previously (22). Briefly, 14-week old BALB/c mice
(Harlan Sprague Dawley,lndianapolis, IN) were anesthetized with isofluorane and
implanted subcutaneously with Alzet mini-osmotic pumps (model 2004, Durect
Corporation, Cupertino, CA) that contained either 0.9% sterile saline vehicle or
1.3 mg/ml leptin (Amylin, San Diego, CA). Osmotic pumps delivered 6.6 mg
leptin per mouse per day. Wounds were closed with staples and mice were given
a one-time injection of 0.15 mg carprofen (Pfizer, New York, NY). Mice were
harvested after 28 days.

Parathyroid hormone (PTH(1-34)) (Bachem, Torrance, CA) was stored in

glass vials topped with argon gas (to prevent oxidation) at -80 °C as a 10'4 M

stock in 4 mM HCI supplemented with 0.1% bovine serum albumin. PTH did not
go through more than one freeze/thaw cycle. Immediately before injection, PTH
was made up to 100 pl per mouse with ice cold 0.9% saline. At 14 weeks of age,
BALB/c mice (Harlan Sprague Dawley,lndianapolis, IN) were subjected to daily
subcutaneous injections of either 40 pig/kg PTH or an equivalent volume of saline

vehicle. Mice were harvested after 40 days.

5.3.2. OC-Wnt10b mice and diabetes induction

188

CS7BU6 breeder mice with transgenic overexpression of Wnt10b from the
osteocalcin promoter (OC-Wnt10b) were obtained from Ormond A. MacDougald
(University of Michigan, Ann Arbor, MI) (44). Heterozygous transgenic mice were
bred at Michigan State University with wild type CS7BL/6J mice (Jackson
Laboratories, Bar Harbor, ME). Mice were genotyped with genomic DNA isolated
from an ear punch (DNAeasy Kit, Qiagen, Valencia, CA). DNA was amplified by
real-time PCR with iQ SYBR Green Supermix (Biorad, Hercules, CA) and
primers specific to the transgene synthesized by Integrated DNA Technologies
(Coralville, IA) (44). All mice were maintained on a 12-hour light, 12-hour dark
cycle at 23 °C, were given standard lab chow, and had food and water ad Iibitum.
Diabetes was induced in female mice only, which have a diabetic bone
phenotype similar to males (23) and at the age we are interested in, OC-Wnt10b
mice have higher bone density (not shown). At 14 weeks of age, female wild type
and transgenic mice were treated with either 50 mglkg streptozotocin (to induce
diabetes) or 0.1 M citrate buffer pH 4.5 vehicle (control) for five consecutive days.
Diabetes was confirmed 12 days after the first injection (dpi, days post injection)
with an AccuChek compact glucometer (Roche, Nutley, NJ) and a drop of blood
collected from the saphenous vein. Blood glucose over 300 mgldl was
considered diabetic. Mice were harvested 4 weeks after diabetes was confirmed

(at 40 dpi). Soft tissues were immediately weighed.

5.3.3. Micro-computed tomography (pCT) analyses

189

Bones were fixed in 10% formalin and transferred to 70% ethanol after 24

. hours. Fixed femurs and tibias were scanned using a GE Explore Locus pCT
system at a voxel resolution of 20 um obtained from 720 views. Beam angle of
increment was 0.5 and beam strength was set at 80 peak kV and 450 HA. Each
run included bones from each treatment group and a calibration phantom to
standardize grayscale values and maintain consistency. Based on autothreshold
and isosurface analyses of multiple bone samples, a fixed threshold (800) was
used to separate bone from bone marrow. Femur trabecular bone analyses were
performed in a region of trabecular bone defined at 0.17 mm proximal to the
growth plate of the distal femur extending 2 mm toward the diaphysis, and
excluding the outer cortical shell. Tibia trabecular bone analyses were performed
in a region of trabecular bone defined at 0.17 mm distal to the growth plate of the
proximal tibia extending 2 mm toward the diaphysis, and excluding the outer
cortical shell. Trabecular bone mineral content (BMC), bone mineral density
(BMD), bone volume fraction (BVF), thickness (Tb Th), spacing (Tb Sp) and
number (Tb N) values were computed by a GE Healthcare MicroView software
application for visualization and analysis of volumetric image data. Trabecular
isosurface images were taken from a cylindrical region in the tibia or femur where

analyses were performed measuring 1.0 mm in length and 1.0 mm in diameter.

5.3.4. RNA analyses

190

Tibias were cleaned of muscle and connective tissue, snap frozen in liquid
nitrogen and stored at -80°C. Frozen tibias were crushed under liquid nitrogen
conditions with a Bessman Tissue Pulverizer (Spectrum Laboratories, Inc.,
Rancho Dominguez, CA). RNA was isolated with Tri Reagent (Molecular
Research Center, Inc., Cincinnati, OH) and integrity was assessed by
formaldehyde-agarose gel electrophoresis. cDNA was synthesized by reverse
transcription with Superscript II Reverse Transcriptase Kit and oligo dT(12-13)
primers (Invitrogen, Carlsbad, CA) and amplified by real-time PCR with iQ SYBR
Green Supermix (Biorad, Hercules, CA) and gene-specific primers synthesized
by Integrated DNA Technologies (Coralville, IA). Hypoxanthine guanine
phosphoribosyl transferase (HPRT) mRNA levels do not fluctuate in diabetes,
PTH treatment, leptin treatment, or overexpression of Wnt10b and were used as
an internal control. HPRT was amplified using 5’-AAG CCT AAG ATG AGC GCA
AG-3’ and 5’-TTA CTA GGC AGA TGG CCA CA-3’ (47). Wnt10b was amplified
using 5’-TCT CTT TCA GCC CTT TGC TCG GAT-3’ and 5’-ACA ACT GAA CGG
AAG GAG AAG CCT-3’ (48). Osteocalcin was amplified using 5’-ACG GTA TCA
CTA TTI' AGG ACC TGT G-3’ and 5’-ACT TTA TI'T TGG AGC TGC TGT GAC-
3’ (49). Runx2 was amplified using 5’-GAC AGA AGC 'I'I'G ATG ACT CTA AAC
C-3’ and 5’-TCT GTA ATC TGA CTC TGT CCT TGT G-3’ (50). aP2 was
amplified using 5’-GCG TGG AAT TCG ATG AAA TCA-3’ and 5’-CCC GCC ATC
TAG GGT TAT GA-3’ (51). PPARyZ was amplified using 5’-TGA AAC TCT GGG
AGA 'I'I'C TCC TG-3’ and 5’-CCA TGG TAA 'I'I'T CTT GTG AAG TGC-3’ (52).

Real time PCR was carried out for 40 cycles using the iCycler (Bio-Rad) and data

191

were evaluated using the iCycler software. Each cycle consisted of 95°C for 15 s,
60°C for 30 5 (except for osteocalcin and runx2 which had an annealing

temperature of 65°C), and 72°Cfor 30 s. cDNA-free samples, a negative control,
did not produce amplicons. Melting curve and gel analyses (sizing, isolation, and

sequencing) were used to verify single products of the appropriate base pair size.
5.3.5. Serum measurements

Blood was obtained from mice at the time of euthanasia and blood serum
prepared from each sample by centrifugation for 10 min at 4,000 rpm. Serum
was aliquoted and stored frozen at -20°C and did not go through more than one
freeze/thaw cycle. Serum osteocalcin levels were measured using Mouse
Osteocalcin EIA Kit (Biomedical Technologies, Inc., Stoughton, MA) aCcording to

manufacturer instructions.
5.3.6. Bone histology and histomorphometry

Fixed femur samples were processed on an automated Thermo Electron
Excelsior tissue processor for dehydration, clearing and infiltration using a routine
overnight processing schedule. Samples were then embedded in Surgipath
embedding paraffin on a Sakura Tissue Tek lI embedding center. Paraffin blocks
were sectioned at 5 pm on a Reichert Jung 2030 rotary microtome. Slides were

stained for tartrate-resistant acid phosphatase (TRAP) activity and counter

192

stained with hematoxylin according to manufacturer protocol (387A-1KT, Sigma,
St. Louis, MO). Osteoclast surface was measured and expressed as a
percentage of total bone surface in the femur trabecular region ranging from the
distal growth plate to 2 mm proximal. Visible adipocytes, greater than 30 um were

counted in the same area.

5.3.7. Statistical Analyses

All measurements are presented as mean :1: standard error. Statistically

significant (or = 0.05) effects were determined with a student’s t-test.

5.4. RESULTS

T1-diabetic bone loss is caused by reduced bone formation with
unchanged or reduced resorption. Reduced osteoblast differentiation is
accompanied by increased marrow adiposity, suggesting a role for altered
lineage selection. We measured wnt signaling family members and found that
Wnt10b, a known regulator of osteoblast and adipocyte differentiation, was
significantly suppressed in diabetic bone (Figure 24A). As we have recently
demonstrated, PTH treatment promotes bone formation in diabetes, but does not
alter marrow adiposity. Consistent with recent findings (46), we found that PTH
induced Wnt10b mRNA levels (Figure 248). Additionally, we have shown that

leptin treated mice have bone loss, but are protected from diabetic marrow

193

adiposity (22). Here, for the first time, we show that leptin treatment reduced

bone expression of Wnt10b (Figure 24C).

194

Figure 24. Wnt10b mimics bone volume fraction changes in diabetes and in
PTH and leptin treatments. (A) Mice were treated with either 50 mglkg
streptozotocin daily for 5 days to induce diabetes (D) or vehicle (control, C). N =
6-10. (B) Mice were treated with either daily subcutaneous injections of 40 jig/kg
PTH (P) or vehicle (control, C) for 40 days. N = 8-14. (C) Mice were implanted
with osmotic minipumps containing either leptin (L) or saline vehicle control (C)
such that leptin-treated mice received 6.6 pg leptin per day. N = 6-8. Fixed
femurs (A) or tibias (8.0) were analyzed by uCT in order to determine bone
volume fraction (BVF) of the trabecular bone immediately proximal to the distal
growth plate of the femur, or immediately distal to the proximal growth plate of
the tibia. Representative three-dimensional images were taken from the same
area as BVF measurements. mRNA from frozen tibias was converted to cDNA by
reverse transcriptase reaction and amplified with primers specific for Wnt10b and
HPRT, a non-modulated housekeeping gene control. Bars represent mean :1:
standard error. *p < 0.05 by student’s t-test.

195

 

 

 

 

 

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Because we have found decreased Wnt10b levels in diabetes and in leptin
treatment (which does not avert diabetic bone loss), and increased in PTH
treatment (which promotes bone formation in diabetes) we wanted to determine
whether overexpression of Wnt10b itself was sufficient to prevent diabetic bone
loss and/or marrow adiposity. We bred mice with overexpression of Wnt10b from
the osteocalcin promoter (OC-Wnt10b) (44), and determined bone density and
Wnt10b expression levels in male and female, wild type and transgenic mice at
21 weeks of age, when we normally examine effects of diabetes one bone. At
this age, the female OC-Wnt10b mice had higher trabecular bone parameters
than males (not shown). We have previously shown that gender does not affect
the diabetic bone phenotype (23), so we induced diabetes in only female OC-
Wnt10b mice and wild type littermates.

Euglycemic wild type and OC-Wnt10b mice had similar blood glucose
levels (Figure 25). Streptozotocin successfully induced diabetes in both
genotypes, and glucose was not significantly different in diabetic OC-Wnt10b
mice compared to diabetic wild type mice. Control OC-Wnt10b mice did not have
significantly different body mass from wild type mice (Figure 25). We normally
see significant weight loss in wild type mice after diabetes is induced, but in this
case we did not. However, diabetes did induce significant weight loss in OC-
Wnt10b transgenic mice. Consistent with no difference in body mass compared
to wild type mice, OC-Wnt10b control mice did not have altered femoral fat pad
or tibialis anterior muscle mass. Diabetic wild type mice did have significant fat

and muscle mass loss compared to control wild type mice, as we and others

197

have demonstrated previously (9). OC-Wnt10b mice also had significant fat and
muscle mass loss when made diabetic, and their levels did not differ from
diabetic wild type mice. Mass changes in the visceral perirenal fat pad (not
shown) mimicked those of the subcutaneous femoral fat pad.

We performed uCT analysis of trabecular bone immediately proximal to
the distal growth plate of the femur (Figure 26, Table 6). As previously reported,
control OC-Wnt10b mice had significantly elevated BMC, BMD, BVF, trabecular
thickness, and trabecular number, and reduced trabecular spacing compared to
control wild type mice (44). Consistent with the literature, diabetes induced a
significant reduction in trabecular BMC, BMD, BVF, thickness and number, but
had no statistically significant effect on trabecular spacing in wild type mice (9,
15). Interestingly, diabetes did not induce significant changes in any of the
trabecular bone density parameters in OC-Wnt10b diabetic mice compared to
OC-Wnt10b controls. Correspondingly, diabetic OC-Wnt10b mice had
significantly higher BMC, BMD, BVF, trabecular thickness, and number, and

lower trabecular spacing than diabetic wild type mice.

198

Figure 25. Overexpression of Wnt10b did not prevent diabetes induction by
streptozotocin. Wild type (WT) and heterozygous OC-Wnt10b (T G) mice were
treated with either streptozotocin to induce diabetes (D) or vehicle (control, C)
and harvested at 40 dpi. Nonfasting blood glucose, body mass, femoral fat pad
mass and tibialis anterior muscle mass were measured immediately. Bars
represent mean 1 standard error. N = 5-8 per group. *p < 0.05 by student’s t-test.

199

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Figure 26. Mice with Wnt10b overexpression were protected from diabetes-
induced trabecular bone loss. Fixed femurs from control (C) and diabetic (D),
wild type (WT) and OC-Wnt10b (TG) mice were analyzed by pCT.
Representative three-dimensional images were taken from the volume of
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fraction (BVF) and bone mineral density (BMD) measurements were determined
in similar region of interest, excluding the cortical bone. Bars represent mean :t
standard error. N = 5-8 per group. *p < 0.05 by student’s t-test.

201

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In order to determine how bone formation was affected by Wnt10b
overexpression, we examined serum and mRNA marl<ers of osteoblast activity.
Serum osteocalcin was elevated in control OC-Wnt10b mice compared to wild
type controls, which is consistent with higher bone formation (Figure 27A).
Similar to previous findings, and consistent with low bone formation in diabetes,
wild type diabetic mice had significantly lower serum 00 levels than wild type
controls. Despite protection from diabetic bone loss, OC-Wnt10b mice had
reduced serum 00 compared to control OC-Wnt10b mice, suggesting that bone
formation was reduced. This finding was unusual because diabetes did not
induce bone loss in these mice, so we examined mRNA markers of bone
formation as well. The pattern of OC mRNA expression in bone was similar to
that of serum 0C levels. Runx2 levels, however, only tended toward higher levels
in OC-Wnt10b control mice compared to wild type controls. Although Runx2 was
significantly decreased in wild type diabetic mice compared to wild type controls,
it only tended toward lower levels (p = 0.07) in diabetic OC-Wnt10b mice
compared to OC-Wnt10b controls.

Despite the apparent reduced bone formation in the OC-Wnt10b mice,
they did not have significantly decreased trabecular bone density, so we
therefore measured bone resorption to gain an understanding of overall bone
remodeling. TRAP5 positive osteoclast surface (% total) was not significantly
altered by Wnt10b overexpression or diabetes (Figure 27C), which is consistent
with previous reports (9, 44). Because osteoclast surface is dependent on total

bone Surface (which is markedly altered with both, diabetes and overexpression

204

of Wnt10b), unchanged % osteoclast surface is indicative of resorption changing
proportionately to bone formation.

In order to address whether diabetic marrow adiposity was altered by
overexpression of Wnt10b, we examined adipocytes in the marrow of femurs. and
measured tibia mRNA expression levels of adipocyte markers (Figure 28).
Overexpression of Wnt10b from the 0C promoter did not alter marrow adiposity
in control mice (Figure 28A), while it did in mice with overexpression of Wnt10b
from the aP2 promoter (39). As we have demonstrated in the past, T1 -diabetes
increased bone marrow adipocyte number in wild type mice. This increase in
marrow adiposity was similar in diabetic OC-Wnt1 Ob mice, suggesting that
overexpression of Wnt10b in osteoblasts was not sufficient to alter adipogenesis
in the marrow. We also measured mRNA markers of adiposity and found that
aP2 expression coincided with adipocyte numbers (Figure 283). PPARyZ
expression level were also unchanged in control OC-Wnt10b mice compared to
control wild type mice. PPARyZ, however, only tended toward increases in both
diabetic groups compared to their genotype matched controls, and diabetic wild

type PPARyZ levels were not different from those of diabetic OC-Wnt10b mice.

205

Figure 27. Diabetes reduces bone formation in both wild type and OC-
Wnt10b mice, but does not alter resorption. (A) Total osteocalcin, a marker of
bone formation, was measured with an ELISA in serum from control (C) and
diabetic (D), wild type (WT) and OC-Wnt10b (T G) mice. (B) mRNA from frozen
tibias was converted to cDNA by reverse transcriptase reaction and amplified
with primers specific for osteocalcin, runx2 and HPRT, a non-modulated
housekeeping gene control. (C) Fixed femurs sections were stained for TRAP
activity counterstained with hematoxylin. Surface of bone in contact with TRAP5
positive osteoclasts was measured and expressed as a percent of the total bone

surface. Bars represent mean :t standard error. N = 5—8 per group. *p < 0.05 by
student’s t-test.

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adiposity. (A) Fixed femurs sections were stained for TRAP activity
counterstained with hematoxylin. Visible adipocytes were counted in the femur
trabecular area immediately proximal to the distal growth plate and extending 2
mm proximal. (B) mRNA from frozen tibias was converted to cDNA by reverse
transcriptase reaction and amplified with primers specific for the adipocyte
markers aP2 and PPARy2, and for HPRT, a non—modulated housekeeping gene
control. Bars represent mean 1 standard error. N = 5-8 per group. *p < 0.05 by
student’s t-test.

208

5.5. DISCUSSION

To address the role of Wnt10b in diabetic bone loss we induced diabetes
in wild type and OC-Wnt10b transgenic mice. Vlflld type diabetic mice did lose
trabecular bone compared to controls, but interestingly, OC-Wnt10b mice did not
lose bone. However, serum and RNA markers of bone formation were
significantly reduced in diabetic mice from both genotypes. Consistent with past
studies, osteoclast surface was not different in any of the treatment groups (9,
44). Additionally, despite the fact that Wnt10b is known to inhibit adipogenesis,
OC-Wnt10b mice still accumulated marrow fat after diabetes was induced. We
concluded that OC-Wnt10b mice are resistant to bone loss from diabetes and
that reduced Wnt10b levels in diabetes may be a significant contributor to bone
loss. Although diabetic OC-Wnt10b mice did have reduced bone formation
markers, we believe that the positive effects of agonizing Wnt10b/B-catenin
signaling could outweigh the negative effects of diabetes if this pathway was
further explored as a treatment target for diabetic bone loss.

Here, we overexpressed Wnt10b in osteoblasts, which may not be the
source of the reduced Wnt10b in diabetes. Since wnts are secreted, there are
several cell types in bone that could be responsible for the reduction of Wnt10b
levels. A very recent report indicates that the anabolic effects of intermittent PTH
are dependent on T-Iymphocyte derived expression of Wnt10b (46). Consistent
with this finding, we found elevated Wnt1 Ob expression in bone of PTH-treated

mice (Figure 248), and we determined that PTH is capable of promoting bone

209

formation in diabetes (Motyl, et al., in preparation, See Chapter 6). Additionally,
recent work from our lab demonstrates that the cellular composition of the bone
marrow is dramatically altered in response to diabetes (Martin, et al. 2010,
submitted), and it will be incumbent upon us to determine how lymphocyte sub-
populations are affected and how marrow derived Wnt10b is altered in diabetes.

Overexpression of Wnt10b from the osteocalcin promoter results in high
expression levels in mature osteoblasts and osteocytes, but not preosteoblasts
(which do not yet express 0C). Therefore, if it is most important for Wnt10b to
work in a paracrine/autocrine dependent manner in preosteoblasts to promote
maturation, then overexpression of Wnt10b in only mature osteoblasts may not
be able to fully prevent the diabetic bone phenotype of reduced osteoblast
maturation (9, 15, 53).

In addition to reduced maturation, diabetic bone loss may likely result from
increased osteoblast apoptosis as well. We have recently demonstrated elevated
osteoblast apoptosis during diabetes onset in mice (Martin, et al., 2010,
submitted). Wnt1 and Wnt3a have been demonstrated to prevent osteoblast
death from serum starvation in vitro in a B-catenin dependent manner (54).
Additionally, oxidative stress has been hypothesized to be involved in diabetic
bone pathology (10, 55). Increased reactive oxygen species (ROS) induce FoxO
transcription factors to bind B-catenin and activate FoxO target genes. This pulls
B-catenin away from TCF/LEF-responsive genes and therefore antagonizes the
end point of wnt signaling (56). Since wnt signaling regulates many of the wnt

pathway members, increased oxidative stress could potentially be responsible for

210

reduced Wnt10b levels we have found in diabetes (57). Agonizing the wnt
pathway with overexpression of Wnt10b may be sufficient to inhibit early
osteoblast apoptosis from diabetes and is a topic of current examination in our
laboratory.

We also found that Wnt10b expression was suppressed by chronic
subcutaneous infusion of leptin (Figure 24C). To our knowledge, this is the first
demonstration of leptin affecting Wnt10b levels in bone. The effect of leptin on
bone density and Wnt10b in this case could be two-fold: through reduced body
mass and mechanical load on bone, or through BZ-adrenergic inhibition of bone
formation. Leptin treatment reduces food intake and body mass, and changes in
bone volume fraction are proportionate to body mass changes (21). Sclerostin,
an endogenous wnt signaling antagonist, is secreted by osteocytes in response
to unloading, then subsequently reduces [El-catenin activity and bone formation in
osteoblasts (58). Alternately, sclerostin levels are reduced in response to
increased load (59). Interestingly, wnt/B-catenin signaling pathway members, and
in particular Wnt1 Ob levels, are elevated in response to increased mechanical
load in vivo and in vitro in microstrained osteoblast cultures (60). It is unclear
whether Wnt10b itself is reduced in response to unloading situations like hind
limb suspension or simulated microgravity and whether this response would be
maintained long term as in our experiment (22). Independent of body mass,
leptin itself can reduce bone density through hypothalamic relay activation of 62-
adrenergic receptor signaling (61). A previous study demonstrated that the B-

blocker porpranolol was able to enhance osteogenic effects of intermittent PTH

211

therapy (62), which we now know is dependent on Wnt10b (46), suggesting that
regulation of these pathways may not be entirely independent.

The fact that overexpression of Wnt10b from the osteocalcin promoter did
not prevent diabetic marrow adiposity (Figure 28) is very interesting because
aP2-Wnt10b mice have significantly reduced marrow and peripheral fat (39, 45).
We did not detect differences in adipocyte numbers or gene expression between
control wild type and OC-Wnt10b mice. This difference could be due to different
levels of Wnt10b protein expression in aP2-Wnt10b versus OC-Wnt10b mice, or
it is possible that as we hypothesized above with osteoblasts, the effect of
Wnt10b in adipocytes is strongest when it is expressed locally.

Despite the fact that we observed reduced markers of mature osteoblasts
in diabetic OC-Wnt10b mice, bone density loss induced by diabetes was
prevented by overexpression of Wnt10b. Diabetic OC-Wnt10b mice had
increased bone density compared to wild type mice (control and diabetic). This
finding suggests that reduced Wnt10b levels in diabetic bone may be in part
responsible for diabetic bone loss in wild type mice. Similarly, our findings
warrant further exploration into agonizing the wnt/B-catenin signaling pathway as

a treatment for diabetic osteoporosis.

5.6. ACKNOWLEDGEMENTS

The authors thank Amylin for providing the leptin and Laurie McCauley for

assistance with the protocol for PTH treatment. We also thank Ormond

212

MacDougald for the OC-Wnt10b breeder mice. We thank Regina Irwin and
Lindsay Martin for technical assistance and critical review of the manuscript. This
work was funded by grants from the National Institutes of Health
(RO1DK061184) and the American Diabetes Association (7-07-RA—105) to LRM.

The authors have no financial conflicts.

213

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CHAPTER 6

6. OSTEOPOROSIS FROM TYPE 1 DIABETES IS REVERSED BY
INTERMITI'ENT PARATHYROID HORMONE STIMULATION OF BONE

REMODELING AND REDUCTION OF OSTEOBLAST APOPTOSIS

6.1 . ABSTRACT

Type 1 diabetic osteoporosis results from impaired osteoblast activity and
osteoblast death. Because diabetes does not affect or reduces resorption,
antiresorptive bisphosphonates may not be the best therapy. Anabolic
intermittent parathyroid hormone (PTH) stimulates bone remodeling and
increases bone density. Here, we examined the ability of 8 uglkg and 40 pg/kg
intermittent PTH to counteract diabetic bone loss. We found significant elevation
of trabecular bone density parameters with the 40 uglkg dose in control mice and
diabetic mice (compared to their metabolic-state matched vehicle treated
controls), and elevation of trabecular bone parameters in diabetic 8 ug/kg treated
mice. Increased bone density in diabetic PTH-treated mice was due to increased
bone formation, mineral apposition, and osteoblast surface, all of which are
defective in type 1 diabetes. Reduction of diabetes-induced osteoblast apoptosis
could account for some of the bone-forming effects of PTH. We also determined
that 40 uglkg PTH treatment reversed preexisting bone loss from diabetes, which

will be significant for clinical application. We conclude that intermittent PTH will

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likely be an effective therapeutic for patients with type 1 diabetes and that its use

in these subjects should be explored.

6.2. INTRODUCTION

Bone is a highly dynamic tissue that is constantly remodeling to maintain
blood calcium homeostasis and to respond to altered demand for structural
support. Osteoblasts (bone forming cells) and osteoclasts (bone resorbing cells)
work simultaneously to repair microcracks and maintain bone density and
strength. Certain diseases and conditions can alter the balance of bone formation
and resorption and can often lead to osteoporosis in men and women, which can
heighten bone loss from aging and menopause (1). One such disease is type 1
(T 1-) diabetes, in which patients are hypoglycemic and hyperinsulinemic and
have bone loss and increased fracture risk (2-6). As with secondary osteoporosis
resulting from any disease, understanding the mechanism of T1-diabetic bone
loss is necessary for choosing the best therapies. Osteoporosis from T1-diabetes
is marked by decreased bone formation (and unchanged or decreased
resorption) in both humans and animals (1, 7-13). Reduced bone formation
markers in diabetes include reduced mRNA levels of osteocalcin (OC), runt-
related transcription factor 2 (runx2), osterix and Dlx5 in bone, and reduced
serum OC, suggesting impaired differentiation and maturation of osteoblasts. It
has been suggested that reduced differentiation to the osteoblast lineage could

be related to increased differentiation of adipocytes in the bone marrow (both

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osteoblasts and adipocytes arise from mesenchymal stem cells (MSCs)) (8, 12-
15). However, recent studies have demonstrated a disconnect between
osteoblast and adipocyte differentiation in T1-diabetes. For example, we have
demonstrated that inhibition of marrow adiposity with neither leptin nor the
PPARy inhibitor bisphenol-A—diglycidyl ether (BADGE) prevents bone loss from
T1-diabetes (16, 17). In addition to reduced maturation, we have found that
osteoblasts undergo apoptosis as early as 2 days after high blood glucose is

detectable in the streptozotocin mouse model of T1-diabetes and at comparably

early time points in the spontaneously diabetic [mil-Ame mice (Martin et al.,

2010, submitted). This osteoblast apoptosis remains detectable for several
weeks. Thus, a treatment that could target osteoblasts (by promoting
differentiation and/or preventing apoptosis) would be ideal for diabetic patients.
Most of the treatments for osteoporosis, however, are antiresorptive,
meaning they work by inhibiting osteoclast activity. Bisphosphonates (the most
widely used antiresorptive therapy) have a similar structure to inorganic
pyrophosphate and are incorporated into bone (18). However, bisphosphonates
are more stable than inorganic phosphate and can withstand exposure to bone
resorbing acids secreted by osteoclasts (19). Therefore, when osteoclasts
encounter bisphosphonates embedded in bone, resorption is halted, and
osteoclasts often undergo apoptosis (20). This could present a problem for
fracture repair and everyday remodeling of microcracks. Intact, recent clinical
evidence suggests long-term bisphosphonate therapy could increase incidence

of fracture (21-25). In diabetes, bone resorption is already suppressed (1); further

224

suppression of resorption with antiresorptive therapies may worsen impaired
fracture healing in these patients. Additionally, one of the Well-characterized side
effects of bisphosphonates is osteonecrosis of the jaw (bisphosphonate-related
osteonecrosis, BON) (26, 27). Diabetes appears to be a risk factor for BON:
compared to the total patient population receiving bisphosphonates for
osteoporosis, diabetic patients receiving bisphosphonates were nearly 5 times
more likely to be diagnosed with BON (28). Other antiresorptive treatments for
osteoporosis exist (including hormone replacement therapy and selective
estrogen receptor modulators (SERMs), and calcitonin). However, because bone
loss from T1-diabetes is due to reduced osteoblast bone formation, and not
increased resorption, we maintain that anabolic therapies that target bone
formation directly may be most appropriate.

Currently, the only anabolic treatment in use is truncated parathyroid
hormone (PTH), also called teriparatide (29, 30). The mechanism of PTH action
in bone is complex and dependent on dosing. Chronic PTH stimulation causes
bone resorption and calcium release. However, when administered intermittently
(daily subcutaneous injections) PTH causes a net increase in bone formation and
reduction of fracture risk in humans and laboratory animals (31-35). PTH
receptors are present on cells in the osteoblast lineage including preosteoblasts,
osteoblasts, osteocytes and bone lining cells. lnterrnittent PTH promotes
osteoblast differentiation and inhibits osteoblast and osteocyte apoptosis,
perhaps through upregulation of runx2 (36). Additionally, in vitro PTH treatment

activates the TCF/LEF-dependent transcription, which is activated by Wnt

225

signaling (37), a potent regulator of bone formation (38, 39). PTH may also inhibit
production of sclerostin, which inhibits bone formation by inhibiting Wnt and bone
morphogenic protein (BMP) signaling (40). A very recent report indicates that the
anabolic effects of intermittent PTH are dependent on T-lymphocyte expression
of Wnt1 0b (41). Because no PTH receptors are present on cells of the osteoclast
lineage, net bone resorption from chronic PTH treatment is likely due to secreted
osteoblast factors (RAN KL and M-CSF) activating osteoclast activity.

There are no reports examining the efficacy of PTH treatment for T1-
diabetic bone loss in humans. In laboratory animals, intermittent PTH treatment
has proven to be anabolic (32), and is effective at increasing bone formation in
models of unloading, ovariectomy, and alcohol consumption (34, 42-46).
Additionally, 4-week intermittent PTH treatment of STZ-diabetic rats improves
bone density parameters 4, 6 and 8 weeks after diabetes induction (47).
Similarly, PTH related protein (PTHrP) treatment of streptozotocin-diabetic mice
also enhances bone formation markers and bone density (48). However, the
effects of the already commercially available intermittent PTH therapy on mouse
models of T1-diabetes induced bone loss have not been determined.

While understanding exactly what diabetic characteristics (hyperglycemia,
inflammation, oxidative stress) cause these bone changes, it is also important to
utilize the therapeutic resources available to treat patients on an individualized
basis. Therefore, here we examined the efficacy of intermittent PTH therapy on
several aspects (bone formation, osteoblast apoptosis, and bone resorption) of

bone loss in T1-diabetic mice. Briefly, we determined that daily subcutaneous

226

PTH (at both 8 ug/kg and 40 ug/kg doses) is sufficient to restore trabecular bone
density of T1 -diabetic mice back to untreated control levels. We demonstrated
that this effect was primarily due to increased osteoblast maturity, viability and
mineralization. We also found that PTH was capable of restoring diabetic bone
density to normal levels even when initiated after bone loss had already
occurred, which is crucial for PTH to be an effective clinical treatment of
osteoporosis in T1-diabetic patients. Our findings are clinically relevant and

warrant further exploration of the use of PTH in human diabetic subjects.

6.3. MATERIALS AND METHODS

6.3.1. Diabetes induction

BALB/c mice were obtained from Harlan Sprague Dawley (Indianapolis,
IN). All mice were maintained on a 12-hour light, 12-hour dark cycle at 23 °C,
were given standard lab chow, and had food and water ad Iibitum. At 14 weeks of
age, mice were treated with either 50 mglkg streptozotocin (to induce diabetes)
or 0.1 M citrate buffer pH 4.5 vehicle (control) for five consecutive days. Diabetes
was confirmed 12 days after the first injection (dpi, days post injection) with an
AccuChek compact glucometer (Roche, Nutley, NJ) and a drop of blood collected
from the saphenous vein. Blood glucose over 300 mgldl was considered diabetic.

Mice were harvested at 5, 20 or 40 dpi.

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6.3.2. PTH Treatment

PTH (Bachem, Torrance, CA) was stored in glass vials topped with argon

gas at -80 °C as a 10'4 M stock in 4 mM HCI supplemented with 0.1% bovine

serum albumin. PTH did not go through more than one freeze/thaw cycle.
Immediately before injection, PTH was made up to 100 pl per mouse with ice
cold 0.9% saline. Control and diabetic mice were subjected to daily
subcutaneous injections of each PTH dosing regimen: 1) daily vehicle treatment
from 0 dpi until harvest, 2) daily 8 uglkg PTH from 0 dpi until harvest, 3) daily 40
uglkg PTH from 0 dpi until harvest, or 4) daily vehicle treatment from 0-19 dpi
followed by daily 40 ug/kg PTH from 20-40 dpi. As stated above, mice were
harvested at 5, 20 or 40 dpi, at which time serum was collected, tissues were
weighed, and tissues and bones were either fixed in formalin or frozen in liquid
nitrogen and stored at -80°C. Blood glucose was measured at the time of harvest
with an AccuChek compact glucometer (Roche, Nutley, NJ). All animal
procedures were performed in accordance with Michigan State University

Institutional Animal Care and Use Committee.

6.3.3. Micro-computed tomography (pCT) analyses

Bones were fixed in 10% formalin and transferred to 70% ethanol after 24
hours. Fixed tibias were scanned using a GE Explore Locus pCT system at a

voxel resolution of 20 um obtained from 720 views. Beam angle of increment was

228

0.5 and beam strength was set at 80 peak kV and 450 pA. Each run included
bones from each treatment group and a calibration phantom to standardize
grayscale values and maintain consistency. Based on autothreshold and
isosurface analyses of multiple bone samples, a fixed threshold (800) was used
to separate bone from bone marrow. Cortical bone analyses were made in a
defined 2 mm x 2 mm x 2 mm cube in the mid-diaphysis immediately proximal to
the distal tibial-fibular junction, with the exception of cortical bone mineral density
(BMD), which were made in a 0.1 mm x 0.1 mm x 0.1 mm cube. Trabecular bone
analyses were performed in a region of trabecular bone defined at 0.17 mm (1%
of the total length) distal to the growth plate of the proximal tibia extending 2 mm
toward the diaphysis, and excluding the outer cortical shell. Trabecular bone
mineral content (BMC), BMD, bone volume fraction (BVF), thickness (Tb Th),
spacing (Tb Sp) and number (Tb N) and cortical BMD, moment of inertia (MOI),
thickness, inner and outer perimeter, and marrow, cortical and total area values
were computed by a GE Healthcare MicroView software application for
visualization and analysis of volumetric image data. Trabecular isosurface
images were taken from a cylindrical region in the tibia immediately distal to the

proximal growth plate measuring 0.8 mm in length and 0.8 mm in diameter.

6.3.4. Bone histology and histomorphometry

Fixed femur samples were processed on an automated Thermo Electron

Excelsior tissue processor for dehydration, clearing and infiltration using a routine

229

overnight processing schedule. Samples were then embedded in Surgipath
embedding paraffin on a Sakura Tissue Tek ll embedding center. Paraffin blocks
were sectioned at 5 um on a Reichert Jung 2030 rotary microtome.

Slides were stained for tartrate-resistant acid phosphatase (TRAP) activity
and counter stained with hematoxylin according to manufacturer protocol (387A-
1KT, Sigma, St. Louis, MO). Osteoclast surface area was measured and
expressed as a percentage of total bone surface in the femur trabecular region
ranging from the distal growth plate to 2 mm proximal. Osteoblasts were counted
and expressed relative to total trabecular bone surface. Visible adipocytes,
greater than 30 pm were counted and number was expressed relative to marrow
area.

To detect cell death in vivo, the TAcs-XL® Basic In Situ Apoptosis
Detection Kit was used according to manufacturer protocol (T revigen lnc.,
Gaithersburg, MD). Positive controls included slides incubated with nuclease.
Five trabecular regions were examined for each femur slide. Total osteoblast
number counted ranged between 45 and 150 per mouse.

For mineral apposition rate (MAR), mice were injected intraperitoneally
with 200 pl of 10 mg/ml calcein (Sigma, St. Louis, MO, USA) dissolved in saline 9
and 2 days before harvest. Fixed L5 vertebrae were then embedded and
sectioned at 5 pm on a Reichert Jung 2030 rotary microtome. Sections were
photographed under fluorescent light and the distance between lines of calcein
was measured. All photomicrograph measurements were performed with Image

Pro Plus software (Media Cybernetics, lnc., Bethesda, MD).

230

6.3.5. RNA analyses

Tibias were cleaned of muscle and connective tissue, snap frozen in liquid
nitrogen and stored at -80°C. Frozen tibias were crushed under liquid nitrogen
conditions with a Bessman Tissue Pulverizer (Spectrum Laboratories, Inc.,
Rancho Dominguez, CA). RNA was isolated with Tri Reagent (Molecular
Research Center, Inc., Cincinnati, OH) and integrity was assessed by
formaldehyde-agarose gel electrophoresis. cDNA was synthesized by reverse
transcription with Superscript II Reverse Transcriptase Kit and oligo dT(12-13)
primers (Invitrogen, Carlsbad, CA) and amplified by real-time PCR with iQ SYBR
Green Supermix (Biorad, Hercules, CA) and gene-specific primers synthesized
by Integrated DNA Technologies (Coralville, IA). Hypoxanthine guanine
phosphoribosyl transferase (HPRT) mRNA levels do not fluctuate in diabetes or
with PTH treatment and were used as an internal control. HPRT was amplified
using 5'—AAG CCT AAG ATG AGC GCA AG-3' and 5’-TTA CTA GGC AGA TGG
CCA CA-3’ (49). Osteocalcin was amplified using 5’-ACG GTA TCA CTA TIT
AGG ACC TGT G-3’ and 5’-ACT T'I'A TTT TGG AGC TGC TGT GAC-3’ (50).
TRAP5 was amplified using 5’-AAT GCC TCG ACC TGG GA-3’ and 5’-CGT AGT
CCT CCT TGG CTG CT-3’ (51). Receptor activator of nuclear factor kappa-B
ligand (RANKL) was amplified using 5'-TT'I' GCA GGA CTC GAC TCT GGA G-3'
and 5'-TCC CTC C'l'l' TCA TCA GGT TAT GAG-3' (52). OPG was amplified
using 5'-GAA GAA GAT CAT CCA AGA CAT TGA C-3' and 5'-TCC ATA AAC

231

TGA GTA GCT TCA GGA G-3' (53). Real time PCR was carried out for 40 cycles
using the iCycler (Bio-Rad) and data were evaluated using the iCycler software.
Each cycle consisted of 95°C for 15 s, 60°C for 30 s (except for osteocalcin
which had an annealing temperature of 65°C), and 72°C for 30 s. cDNA-free
samples, a negative control, did not produce amplicons. Melting curve and gel
analyses (sizing, isolation, and sequencing) were used to verify single products

of the appropriate base pair size.

6.3.6. Statistical Analyses

All measurements are presented as mean a: standard error. Statistically
significant ((1 = 0.05) main effects (of PTH dose or diabetes) as well as PTH x
diabetes interaction (which would indicate diabetes altering PTH effects or visa
versa) were determined using factorial analysis of variance (ANOVA) and one-
way ANOVA with Tukey HSD post hoc test (where necessary) with SPSS
statistical software (Chicago, IL). Student’s t-test was also used to determine

significance where necessary.

6.4. RESULTS

6.4.1. Diabetes induction and body composition

232

In order to assess the efficacy of anabolic PTH therapy for osteoporosis
from T1-diabetes, we induced diabetes with streptozotocin in BALB/c mice while
simultaneously beginning a PTH treatment regimen. Control and diabetic mice
were injected daily with either vehicle for PTH, 8 uglkg PTH, or 40 pig/kg PTH.
Mice were harvested 40 days after the first injection (dpi). Statsitical analysis with
ANOVA indicated a significant (p < 0.05) effect from diabetes on blood glucose
(as we would expect) but no significant effect of PTH treatment on blood glucose
of either non-diabetic (euglycemic) or diabetic mice. Similarly, ANOVA did not
detect a significant interaction between PTH and diabetes, indicating that it did
not interfere with diabetes induction (Table 7). Similar to previous findings,
diabetic mice weighed 9% less than controls at the end of the study (16). Weight
loss was due in part to muscle and fat loss: diabetic mice had 16% lower tibialis
anterior mass, 41% lower femoral fat pad mass and 57% lower perirenal fat pad
mass (Table 7). When analyzed by ANOVA, neither 8 nor 40 ug/kg PTH altered
the diabetes-induced loss of total body, fat or muscle mass (Table 7). As others
and we have previously demonstrated, loss of peripheral and visceral fat depots
in diabetes was accompanied by increased bone marrow adiposity (Table 7).
Diabetes induced a 3.3-fold increase in marrow adipocyte number in the distal
femur of vehicle treated mice. Interestingly, 8 pg/kg PTH treatment doubled
marrow adipocyte number in control mice, although this was not significant by
ANOVA, and neither was the 1.5-fold increase in 40 ug/kg PTH treated controls.
The level of adiposity in diabetic mice did not differ between PTH treatment

groups.

233

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234

6.4.2. PTH Counteracted Diabetic Bone Loss

When analyzed by ANOVA, significant diabetes effects (p < 0.05) were
found with BMC, BMD, BVF, trabecular thickness and trabecular number (Figure
29, Table 8). Significant PTH effects were found with all of the above parameters,
in addition to trabecular spacing. No significant interaction between diabetes and
PTH was found in any of the trabecular bone parameters. Significance between
groups was determined with a post-hoc test (only after factorial ANOVA
determined significance). Only high dose, 40 pglkg, PTH significantly increased
trabecular BMC, BMC, BVF, trabecular thickness, and trabecular number in
euglycemic mice (Figure 29, Table 8). The increase in trabecular bone
parameters observed in euglycemic mice treated with 8 uglkg was not significant
by ANOVA. As we have demonstrated in the past, diabetes significantly reduced
tibia trabecular BMC, BMD, BVF, trabecular thickness, and trabecular number
(Figure 29, Table 8) (8). Although diabetes still reduced all of these parameters in
PTH treated mice, bone density did not decline to the level of vehicle treated
diabetics, and, in the 40 (.9le group, were more closely aligned with those of
healthy untreated controls. Similar to the non-diabetic mice, all of the trabecular.
bone parameters were significantly elevated (with the exception of trabecular
spacing, which was significantly decreased) in the diabetic 40 (1ng PTH group
compared to the vehicle treated diabetics. The increase in bone density in
diabetic PTH treated mice (compare to vehicle treated diabetics) was higher with

the 40 ug/kg dose of PTH than with the 8 uglkg close, but 8 ug/kg PTH still

235

produced a significant increase in BVF and trabecular number. For example,
trabecular number increased 25% in 8 ug/kg PTH treated diabetics and 39% in
40 ug/kg PTH treated diabetics.

Cortical bone thickness was significantly affected by diabetes and PTH
treatment, according to ANOVA, but there was no significant interaction between
diabetes and PTH. Thickness was increased in 40 pig/kg PTH treated euglycemic
controls (compared to vehicle treated controls), but not 8 09le treated controls
(Table 8). This can be attributed to an increase outer perimeter in the 40 itng
group, which consequently increased both cortical and total area, while 8 uglkg
PTH did not. Similar to our previous findings, diabetes did not induce any
significant changes in cortical bone parameters in vehicle treated mice. However,
diabetes did reduce cortical thickness and cortical area in 40 ug/kg PTH treated
mice compared to 40 pglkg PTH treated controls, but these parameters were not

significantly different from healthy untreated controls. No significant changes

were found in cortical BMD, MOI, inner perimeter or marrow area.

236

Figure 29. PTH treatment counteracted trabecular bone loss from T1-
diabetes. Diabetes was induced with STZ at 14 weeks of age. At the same time,
control (citrate buffer only) and diabetic mice were started on a daily regimen of
subcutaneous injections of PTH (8 or 40 pig/kg) or saline vehicle. PTH treatment
was continued for the remainder of the study. Mice were harvested at 40 dpi and
tibias were analyzed by uCT. (A) Representative three-dimensional isosurface
images of the trabecular bone of the proximal tibia of control and diabetic, vehicle
and PTH treated mice. (B) BVF of control (white bars) and diabetic (gray bars),
vehicle and PTH treated mouse tibias. Bars represent mean 0 standard err0r. N

z 12 per group. Significance determined with ANOVA. *p < 0.05 compared to
treatment (vehicle, 8 or 40 pg/kg PTH) matched control. "p < 0.05 compared to

vehicle-treated control. #p < 0.05 compared to vehicle-treated diabetic.

237

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239

6.4.3. PTH increased bone formation in a dose-dependent manner

PTH treatment of euglycemic control mice significantly (by factorial
ANOVA) affected osteocalcin, such that there was a stepwise increase in
expression that reached statistical significance in the 40 (1ng PTH treated group
(Figure 30A). Diabetes also had a significant effect on 00 levels: diabetes
reduced 00 levels in untreated mice, consistent with previous findings, and that
reduction remained evident in both PTH treated diabetic groups compared to
PTH treated controls. However, as with the controls, PTH treatment increased
osteocalcin in the diabetic mice compared to untreated diabetics, and this was
significant with the 40 pig/kg dose. Furthermore, 00 mRNA levels were not
different in the diabetic 40 ug/kg mice compared to untreated controls, which
suggests equivalent levels of bone formation in these two groups.

In order to address whether bone formation changes were due to altered
osteoblast surface or rate of mineralization, we counted osteoblasts on the
trabecular surface and examined the distance between calcein labels
incorporated in to the bone. When expressed per mm bone surface, osteoblast
number was not significantly elevated in either 8 or 40 pig/kg PTH treated control
groups compared to untreated controls (Figure 308). However, mineral
apposition rate (MAR) was significantly increased in the 40 ug/kg PTH control
group compared to untreated and 8 ug/kg PTH treated controls (Figure 300),
suggesting the increased bone formation was due to faster mineralization, rather

than more bone forming surface. When made diabetic, untreated mice had

240

significantly less osteoblasts and reduced MAR, which is consistent with diabetic
osteoporosis being primarily the result of an osteoblast defect. Interestingly, PTH
treatment prevented any significant reduction in osteoblast number in diabetics
compared to PTH treated controls (Figure 308). Additionally, osteoblast number
was significantly higher in PTH treated diabetic mice compared to untreated
diabetic mice. MAR trended (p = 0.08) to be lower in the 8 uglkg PTH treated
diabetic mice compared to PTH treated controls (Figure 300), which could still
account for bone loss without altered osteoblast number. Similarly, MAR was
significantly reduced in diabetic 40 pglkg PTH treated mice compared to
treatment-matched controls, but the value of MAR in 40 pg/kg PTH treated mice
was not different from untreated control mice and was significantly higher than
untreated diabetic mice. Taken together, high dose PTH appears capable of
promoting bone formation in diabetic mice by preventing diabetes-induced loss of
osteoblasts and by increasing MAR. Minor protection from bone loss in diabetic 8
pig/kg PTH treated mice was likely also due to improved osteoblast number and

MAR.

241

Figure 30. PTH promotes bone formation in diabetes. (A) mRNA from frozen
tibiae was converted to cDNA and amplified with primers specific for osteocalcin
(0C) and the housekeeping gene HPRT. (B) Osteoblasts lining the surface of
trabeculi in the distal femur were identified in hematoxylin stained slides based
on morphology, counted, and expressed per mm bone surface. (C) The distance
between calcein double labels was measured in undecalcified, unstained L5
vertebrae sections and expressed per day. Bars represent mean :1: standard error
of control (C, white bars) and diabetic (D, gray bars), vehicle, 8 ug/kg and 40
pg/kg PTH treated mice at 40 dpi. PTH treatment was daily from 0 to 40 dpi. N =

4-8 per group. Significance determined with ANOVA. *p < 0.05 compared to
treatment (vehicle, 8 or 40 pg/kg PTH) matched control. "p < 0.05 compared to
vehicle treated control. Mp < 0.05 compared to vehicle treated control and 8

pglkg PTH treated control. #p < 0.05 compared to vehicle treated diabetic.

242

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6.4.4. Effect of PTH on Bone Resorption

Total bone remodeling (as indicated by TRAP5 mRNA expression) was
significantly elevated in the non-diabetic 40 pg/kg PTH treated group, but not in
the 8 pglkg group, compared to untreated controls (Figure 31A), which together
with increased bone formation, is indicative of increased remodeling overall in the
40 pg/kg group. As we have demonstrated previously, diabetes did not alter total
bone resorption in untreated mice, and this effect was consistent in the PTH
treated groups. In agreement with overall increases in remodeling, the 40 ug/kg
PTH treated diabetic mice had elevated TRAP5 expression compared to vehicle
treated diabetic mice. We also examined osteoclast surface and did not find any
significant diabetes or PTH effects, despite the increase in TRAP5 expression
with the 40 pglkg dose (Figure 31 B). it is possible that there is more total
resorption in these mice (since they have more bone surface), but when
expressed as a function of total surface, it is unchanged. The fact that there is no
change in osteoclast surface in diabetic vehicle treated mice compared to

controls is consistent with our previous findings (8).

244

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Figure 31 . High dose PTH promotes bone resorption in diabetic mice. A)
mRNA from frozen tibiae was converted to cDNA and amplified with primers
specific for tartrate resistant acid phosphatase (TRAP5) and the housekeeping
gene HPRT. (B) Osteoclasts were identified with TRAP5 staining in the distal
femur. Length of trabecular bone surface covered by osteoclasts was measured
and expressed as a percent of the total bone surface. Bars represent mean 2
standard error of control (C, white bars) and diabetic (D, gray bars), vehicle, 8
pg/kg and 40 (Lg/kg PTH treated mice at 40 dpi. PTH treatment was daily from 0

to 40 dpi. N = 4-8 per group. Significance determined with ANOVA. *p < 0.05
compared to treatment (vehicle, 8 or 40 ug/kg PTH) matched control. "p < 0.05

compared to vehicle—treated control. p < 0.05 compared to vehicle—treated
diabetic.

245

6.4.5. Osteoblast viability is improved by PTH treatment

We have previously demonstrated significantly elevated osteoblast death
in diabetic mice throughout the time course of diabetes (Martin et al. 2010,
submitted), which could account for the reduced osteoblast surface and MAR. In
order to address the ability of PTH treatment to reduce diabetic osteoblast death,
we induced diabetes as before, and simultaneously began treating control and
diabetic mice with daily injections of 40 ptng PTH. We harvested the mice at 5
dpi, a time point shortly after blood glucose rises and osteoblast death is
detectable (12), even though mice are not technically considered diabetic (>300
mgldl) at this time. PTH did not alter blood glucose levels at this early time point
(Figure 328). No significant changes in RANKLIOPG ratio were detectable,
which is consistent with no alterations in osteoclast activation in diabetes (Figure
320). In order to address osteoblast activity, we measured tibia mRNA levels of
osteocalcin (Figure 320). PTl-l treatment trended to increase osteocalcin
expression in non-diabetic mice (p = 0.1). As we have demonstrated in the past,
osteocalcin was reduced nearly 5-fold in untreated diabetic mice compared to
untreated controls (12). PTH-treated diabetic mice had significantly higher
osteocalcin expression that untreated diabetics, which is consistent with our
previous findings at 40 dpi (Figure 30). These results suggest that the early
diabetes-induced reduction of bone formation can also be treated with PTH.

Because early bone changes have been demonstrated to be due to

increased osteoblast apoptosis as well as reduced maturation, we examined

246

TUNEL-stained femur sections to quantify osteoblast death (Figure 32A and
32E). Consistent with our previous finding, TUNEL positive osteoblasts increased
in untreated diabetic mice compared to controls. PTH-treatment of euglycemic
mice significantly lowered baseline TUNEL staining. Although diabetes still
induced osteoblast death in PTH-treated mice, the level of TUNEL staining was
not different from untreated controls and trended to be lower than that in
untreated diabetics. These results suggest that the mechanism of improved bone
formation in PTH-treated diabetic mice could be reduced basal osteoblast

apoptosis.

247

Figure 32. PTH ameliorates diabetes-induced osteoblast death. Mice were
injected with either citrate buffer (control) or STZ to induce hyperglycemia. At the
same time, control and diabetic mice were started on a daily regimen of
subcutaneous injections of PTH (4O pig/kg) or saline vehicle. PTH treatment was
continued until harvest at 5 dpi. (A) TUNEL stained trabecular bone in the distal
femur. Dark brown nuclear stain (indicated by arrows) is positive for TUNEL
label. (B) Blood glucose was measured at the time of harvest and was not
significantly altered by PTH-treatment. mRNA from frozen tibiae was converted
to cDNA and amplified with primers specific for RANKL or OPG (C) or with
primers specific for DC and the housekeeping gene HPRT (D). (E) TUNEL
positive osteoblasts were identified based on dark brown nuclear stain, counted
and expressed relative to total osteoblasts. Bars represent mean a; standard error
of control (C, white bars) and diabetic (D, gray bars), vehicle and 40 (1.9le PTH
treated mice at 5 dpi. PTH treatment was daily from 0 to 5 dpi. N = 5-6 per group.

Significance determined with student’s t-test. *p < 0.05 between bracketed bars.

248

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249

6.4.6. Diabetic bone loss can be reversed by PTH

Although PTH treatment was able to counteract bone loss when initiated
simultaneously with diabetes induction (Figure 29 and Table 8), we recognize
that diabetic patients will most likely not start treatment for bone loss when
diagnosed with diabetes. Even if this were the case, diagnosis does not occur
until after significant changes in blood glucose levels are evident, and based on
our findings, after bone changes are initiated (12)(Martin et al., 2010, submitted).
Therefore, we wanted to address whether PTH could improve bone density even
after bone loss had already occurred.

We induced diabetes with streptozotocin as before. At 20 dpi, we either
harvested control and diabetic mice, or treated control and diabetic mice with 40
pig/kg PTH or vehicle. At 20 dpi, diabetic mice had significantly lower BMD and
showed a trend toward lower (p = 0.07) BVF (Figure 33A and 338). Similarly,
untreated diabetic mice at 40 dpi had lower BMD and BVF compared to
untreated controls. Treating non-diabetic mice with 40 pglkg PTH days 20-40
was not enough to induce a significant increase in BMD or BVF compared to
untreated controls. However, treating diabetic mice with PTH days 2040
increased BMD and BVF to levels not different from PTH-treated euglycemic
controls, indicating short-term PTH imparts a stronger affect in diabetic mice than
in controls.

To address the osteoblast/osteoclast phenotype in these mice we

measured osteocalcin and TRAP5 mRNA levels (Figure 33C). We significantly

250

reduced osteocalcin levels at 20 dpi and 40 dpi in untreated diabetic bone
compared to untreated controls. PTH treatment days 20-40. promoted osteocalcin
expression in both control and diabetic mice, although the increase was not
significant. However, osteocalcin levels in PTH-treated diabetic mice were not
different from untreated controls. Similar to our previous findings, the resorption
marker TRAP5 was unchanged or trended to decrease in diabetic mice at days
20 and 40, respectively. PTH treatment for the last 20 days of the experiment
was not enough to increase TRAP5 levels in euglycemic mice. Similarly, TRAP5
expression tended toward increased levels in PTH-treated diabetic mice
compared to untreated diabetic mice at 40 dpi, but this was not statistically
significant. Fortunately, however, PTH is capable of restoring trabecular bone

density to normal levels even after bone loss has already occurred.

251

Figure 33. PTH promotes bone formation in diabetic mice when initiated
after diabetic bone loss is detectable. Mice were treated with streptozotocin to
induce diabetes or citrate buffer (control). At 20 dpi, mice were either harvested,
or treated with vehicle or 40 pg/kg PTH daily until harvest at 40 dpi. (A)
Representative pCT isosurface images of trabecular bone immediately distal to
the proximal growth plate. (B) Tibia bone mineral density (BMD) and bone
volume fraction (BVF) from control (C, white bars) and diabetic (D, dark gray
bars) untreated and PTH treated mice. (C) mRNA from frozen tibiae was
converted to cDNA and amplified with primers specific for the bone formation
marker osteocalcin or resorption marker TRAP5 and expressed relative to the
housekeeping gene, HPRT. Bars represent mean :r: standard error. N 2 6 per

group. Significance determined with student’s t-test. *p < 0.05 between

bracketed bars.

252

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253

6.5. DISCUSSION

Osteoporosis is a severe complication of T1-diabetes that results from
reduced bone formation, and unchanged or reduced resorption. Therefore, we
wanted to determine whether anabolic, intermittent PTH therapy was capable of
enhancing bone formation in T1-diabetes, and apropos, whether it would be an
appropriate therapy for diabetic patients. Briefly, we found that 40 ug/kg PTH
(and to a lesser extent 8 pg/kg PTH) counteracted trabecular bone changes
when diabetic mice were treated daily for 40 days, starting at the same time as
the diabetes-inducing streptozotocin injections. The higher dose of PTH was able
to promote overall bone remodeling by increasing osteocalcin expression, MAR,
osteoblast number and TRAP5 expression in diabetic mice. In addition, PTH
treatment lowered basal osteoblast apoptosis, such that diabetes-induced
osteoblast apoptosis levels were not different from apoptosis levels in control,
untreated mice. Finally, we also determined that PTH treatment was capable of
reversing diabetic bone loss after significant trabecular bone changes had
already occurred, which makes PTH treatment in diabetic patients clinically

relevant.
6.5.1. Both low and high dose PTH treats trabecular bone loss from diabetes

Although diabetes significantly reduced MAR and osteocalcin expression
in 40 pg/kg PTH-treated mice, PTH treatment increased baseline levels of bone

formation such that the values in diabetic PTH-treated mice did not differ from

254

those of untreated controls. This is consistent with the known PTH regulation of
osteoblast proliferation and differentiation, and stimulation of Runx2
transcriptional activity (by which osteocalcin is regulated) (54, 55). Similarly, tail-
suspended mice treated with 40 pg/kg PTH (5 days/wk) had increased bone
formation rate, albeit to a lesser extent that control mice (42). Our finding that
diabetic PTH treated bone parameters do not differ from untreated controls is
extremely significant. Treating human diabetic patients with PTH may promote
bone formation, and overall remodeling, to levels that are comparable to healthy
age-matched subjects, which would be the ultimate goal of any therapy.

Low dose 8 ug/kg PTH did not significantly increase trabecular bone
density in control mice, but there was a clear trend toward increased bone
formation based on bone density measurements and tibia osteocalcin expression
(Figure 29, Table 8, Figure 30). Interestingly, PTH appeared to be more capable
of increasing bone density in the diabetic mice: 8 pig/kg PTH treated diabetic
mice had significantly higher BVF, trabecular number and osteoblast number
compared to untreated diabetic mice and did not have reduced MAR compared
to 8 pg/kg PTH treated controls. In younger 9.5-week-old CS7BL16 mice, 10
(Lg/kg PTH modestly increased trabecular bone density parameters (56),
suggesting that even a slightly higher dose might have had a significant effect in
our euglycemic mice. In rats and after bone perturbations, effective doses of PTH
appear to be much lower: 1 ug/kg PTH was capable of increasing bone density in
hind-limb unloaded rats (57), although it was unclear whether this dose would

have had an effect in healthy, load bearing rats in this study. Similarly, rats had

255

increased BMC, BMD, and fracture healing by day 21 of 30 mglkg PTH treatment
and by day 35 of 5 jig/kg PTh treatment (58). In another study 10 pig/kg PTH
increased ultimate load to failure, BMD, BMC of rats by 28 days after fracture
(59). Thus, it is possible that if our experiment had extended longer, we would
have seen a significant effect with the 8 pig/kg dose in the control mice, and a
stronger effect in the diabetic mice. Pettway, et al. demonstrated that the greatest
increases in osteoblast proliferation occur after the first week of treatment (54).
This idea, in combination with the fact that lower dose (8 pg/kg) PTH is capable
of improving bone density (Table 8), suggests that an ideal treatment regimen
might consist of an initial time period of high dose PTH to stimulate bone
formation, followed by lower dose PTH to maintain bone density at a healthy

level.

6.5.2. PTH restores bone density after it has occurred

We determined that PTH could restore bone density to normal levels even
after bone loss had already occurred from diabetes (Figure 33), which is
consistent with other bone loss conditions, but has not been demonstrated with
diabetes. Sibonga, et al. demonstrated that 80 jig/kg PTH could restore
preexisting bone density in rats fed alcohol, which, like diabetes reduces bone
formation (46). Additionally, the same dose of PTH reverses bone loss from
ovariectomy in rats (44). Lozano, et al. demonstrated increased bone formation

in diabetic mice treated with PTHrP, which signals through the shared

256

PTH/PTHrP receptor, two weeks after diabetes was confirmed, a time point when
bone loss should have been detectable (17, 48, 53). It was important to address
this issue because pathways responsible for bone loss from T1-diabetes (which
are not completely understood) could overlap with the pathways for PTH induced
bone formation, and if this were the case, then PTH might not have been an

effective therapeutic.

6.5.3. Diabetic response to PTH is stronger

Interestingly, we determined that PTH-treatment of diabetic mice during
the 20-day period had a greater effect than PTH-treatment of control mice for 20
days (Figure 33). It is possible that the diabetes-induced suppression of
resorption actually helped bone density increase faster in the diabetic group,
although we did not detect a significant suppression of TRAP5 expression from
diabetes (p = 0.2) in this case. Along the same lines, PTH did not increase
osteoblast number per bone surface (Figure 30B) in control mice, but it did
prevent a decrease after diabetes induction, suggesting a stronger response of
osteoblasts to PTH in a diabetic environment.

Martin, et al. 2010 (submitted) demonstrated that bone marrow cell
composition is altered at early time points during diabetes-induction, and could
potentially be altered throughout the disease. This could account for increased
expression of inflammatory cytokines we have observed in diabetic bone (12).

Recent evidence suggests that intermittent PTH requires T-cell derived

257

expression of the potent osteogenic protein Wnt10b, and that its action is through
canonical wnt signaling (41). However, it is unclear how important canconical wnt
signaling is for PTH induced bone formation. The absense of LRP5 (a canonical
wnt coreceptor) does not alter PTH-induced increases in bone density in male or
female mice (60, 61). Similarly, LRP5 is not required for high bone remodeling
caused by overexpression of PTH/PTHrP receptor in osteocytes, but it is
required for the high bone mass phenotype in these mice (62). However,
canonical wnt signaling can also work through LRP6. Interestingly, PTH is
capable of activating TCF-dependent transcription by
phosphorylating/inactivating GSK3B through the PKA/cAMP pathway (63). This
could account for some of the observed effects of PTH on wnt pathway family
members (37). There is also evidence that PTH works through Wnt4, a non-
canonical wnt ligand that does not require LRP5/6 (64-66). Whether or not
marrow alterations affect the potency of PTH treatment, the ability of PTH to
stimulate bone formation faster in diabetic mice supports the notion that
osteoporosis treatments should be tailored to etiology of the disease and
warrants further investigation into the effects of PTH therapy in human subjects

with T1-diabetes.
6.5.4. Reduction of osteoblast death: basal and diabetes induced

In addition to PTH-induced osteoblast differentiation and activity, there is

ever-increasing evidence that PTH prevents osteoblast apoptosis. Bellido, et al.

258

demonstrated a significant reduction of osteoblast apoptosis with 10—300 pglkg
after 28 days in female Swiss-Webster mice (36). Here, we demonstrated that 40
ug/kg PTH reduced basal osteoblast apoptosis and reduced diabetes-induced
increases in osteoblast apoptosis to a level comparable to that of untreated
control mice (Figure 32). The mechanism of PTH protection against apoptosis
may be dependent on Smad3 (67, 68). Recent evidence indicates that PTH

promotes repair of DNA damage by increasing PCNA and Foxo3a (69).

6.5.5. Cortical bone effects

Although it appeared that trabecular effects of PTH were stronger in
diabetes, we have some evidence that cortical effects of PTH were weakened by
diabetes. We found that 40 (Lg/kg PTH treatment increased cortical bone
thickness and outer perimeter in euglycemic mice. This is consistent with other
studies in ovariectomized mice and rats (70, 71), however in our case, PTH could
not increase cortical thickness in diabetic mice compared to diabetic controls
(Table 8). This suggests that the mechanism of cortical bone accrual in PTH
treated mice is affected by diabetes whereas the mechanism of trabecular bone
accrual is not. Calvi, et al. demonstrated a similar phenomenon: when the PTH
receptor was constitutively active, trabecular bone density increased while
periosteal MAR and cortical thickness decreases (72), suggesting the
mechanism of PTH action is location-dependent. Recently, Jilka, et al. found that

although PTH has a profound anti-apoptotic effect in trabecular bone, cortical

259

bone accrual from PTH is more likely due to pro-differentiation effects on
preosteoblasts because of the comparatively low levels of osteoblast apoptosis in
the periosteum, coupled with fast (2 day) increases in periosteal osteoblast
number after PTH treatment (73, 74). Levels of hyperglycemia equivalent to
those in diabetes are known to have antiodifferentiation as well as pro-apoptotic
effects on osteoblasts (75)(Martin, et al. 2010) and it is likely that both

differentiation and apoptosis play a role in the diabetic osteoblast phenotype.

6.6. SUMMARY

Our data indicate that PTH is an effective treatment for T1—diabetic bone
loss in mice because it promotes remodeling and reduces diabetes-induced
osteoblast apoptosis. Because of its ability to restore bone density to normal
levels, even after bone loss has already occurred, it is likely to be an effective
therapeutic in human patients. Intermittent PTH therapy might be a better option
than the commonly used antiresorptive bisphosphonates because of its ability to
promote bone formation and resorption, which are both depressed in diabetic

patients.

6.7. ACKNOWLEDGEMENTS

The authors thank Laurie McCauley for her advice on PTH treatment. The

authors thank Regina Irwin and Lindsay Martin for technical assistance and

260

critical review of the manuscript, and the MSU Investigative Histology Laboratory
for technical assistance. This work was funded by grants from the National
Institutes of Health (RO1DK061184) and the American Diabetes Association (7-

07-RA-105) to LRM. The authors have no financial conflicts.

261

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