The Pennsylvania State University

The Graduate School

Department of Neural and Behavioral Sciences

BLOCKADE OF THE OGF-OGFR AXIS ENHANCES REPAIR PROCESSES

IN DIABETIC SKIN AND BONE

A Dissertation in

Anatomy

by

Michelle B. Titunick

©2018 Michelle B. Titunick

Submitted in Partial Fulfillment of the Requirements

for the Degree of

Doctor of Philosophy

December 2018

ii

The dissertation of Michelle B. Titunick was reviewed and approved* by the following: Patricia J. McLaughlin Professor of Neural and Behavioral Sciences Director of the Graduate Program in Anatomy Dissertation Advisor Chair of Committee Ian S. Zagon Distinguished Professor of Neural and Behavioral Sciences Gregory S. Lewis Assistant Professor of Orthopaedics and Rehabilitation Christopher Niyibizi Associate Professor of Orthopaedics and Rehabilitation *Signatures are on file in the Graduate School.

iii

ABSTRACT

Almost 9.5% of the U.S. population had diabetes in 2015 with 5-10% having type 1

diabetes. In addition to diabetic complications such as retinopathy, nephropathy,

neuropathy, and cardiovascular disease, there is also poor wound healing and delayed

fracture healing. Preclinical studies from our laboratory have demonstrated that topical

naltrexone (NTX) is an effective and safe treatment for diabetic complications such as

corneal keratopathy, dry eye, and non-healing ulcers. Naltrexone is an opioid receptor

antagonist that, at an appropriate dosage, blocks the opioid growth factor (OGF)-opioid

growth factor receptor (OGFr) axis resulting in increased cell proliferation. [Met5]-

enkephalin, or OGF, is a highly conserved endogenous opioid peptide that tonically

regulates cell proliferation by delaying the G0/G1 phase of the cell cycle.

Both systemic and topical NTX have been shown to enhance corneal epithelial wound

healing in normal and diabetic rats, diabetic rabbits, as well as the type 2 diabetic model,

db/db. Topical NTX reverses dry eye in diabetic animal models, and increases collagen

formation and angiogenesis in cutaneous wounds of diabetic rats and mice. Phase 1

clinical trials with topical NTX administered to the eye have reported tolerability and

safety of the therapy; preclinical pathology studies confirm the lack of toxicity following

sustained topical application.

The underlying cause of delayed healing of bone fractures in diabetes is unclear. Both

preclinical and clinical studies report elevated levels of enkephalins in diabetics

suggesting that the increase in OGF, an inhibitory growth factor, may suppress cell

replication and contribute to poor wound and fracture healing. There is limited

iv

information on the presence and role of the OGF-OGFr pathway in normal bone growth

or in the process of fracture repair. Furthermore, the ability to manipulate the OGF-OGFr

axis with naltrexone during the process of bone fracture repair is unknown.

This dissertation hypothesizes that topical naltrexone to block the OGF-OGFr

regulatory pathway is an effective modulator of diabetic complications related to delays

cutaneous wound healing and bone fracture repair. The first aim tested the hypothesis

that topical naltrexone enhances closure of full-thickness cutaneous wounds in type 1

diabetic rats at a rate comparable to standard of care- Regranex®. In aims 2 and 3, studies

were conducted to investigate the role of the OGF-OGFr axis in diabetic bone. Studies

were designed to determine whether diabetes is associated with a dysregulation of the

OGF-OGFr pathway that subsequently changes bone composition in a type 1 diabetic rat

model, and whether blockade of the pathway alters the rate of repair of diabetic bone.

Lastly, studies were conducted to test the hypothesis that systemic naltrexone blockade

of the OGF-OGFr pathway in diabetic animal models may protect against the

complications related to bone fracture and repair.

Male Sprague Dawley rats (Charles River Laboratories) were used throughout all the

experiments and type 1 diabetes was induced by injections of streptozotocin. In aim 1

topical application of NTX accelerated the rate of closure of 6mm full thickness cutaneous

wounds at a rate comparable to a daily application of the standard of care Regranex®.

Analyses of the skin revealed that naltrexone treatment increased DNA synthesis, as well

as expression of platelet-derived growth factor and vascular endothelial growth factor,

required for the granulation tissue formation and angiogenesis.

v

In the second series of experiments, OGF expression was detected at a comparable

level in bones from normal rats aged 1 day to adult. OGFr expression was detected in the

femur at all ages, and expression was elevated at days 1 and 21. OGF and OGFr expression

were increased in type 1 diabetic rat bone relative to normal, non-diabetic bone. Serum

levels of OGF were also increased in diabetic rats.

Histological analyses of femurs from normal and type 1 diabetic rats revealed no

significant differences in the number of osteoclasts, but did indicate with safranin O

staining a decrease in calcified cartilage in diabetic rats. In comparison to normal bone

composition, Ki67 staining showed a decrease in proliferative cells, and VEGF staining

revealed a decrease in vascularity in diabetic bone. Seven days following fracture, the

callus was examined by radiography and histology. Calluses in normal femurs had more

cartilage than in diabetic bones, and more granulation tissue was evident in diabetic bones

in comparison to diabetic bones treated with naltrexone. Bone tissue treated with

naltrexone from diabetic rats displayed elevated levels of Ki67 staining relative to tissue

treated with vehicle suggesting that topical application of naltrexone may accelerate early

phases of bone repair. Serum OGF levels increased in DB rats with fractures treated with

vehicle compared to the serum of DB rats without a fracture, suggesting fracture or

fracture repair may increase serum enkephalin levels. NTX-treated rats had decreased

serum OGF levels relative to that of diabetic vehicle-treated rats.

Assessment of diabetic bone composition in animals treated systemically with

either saline or naltrexone revealed that naltrexone-treated bones appeared to be stronger

and absorbed more energy than vehicle-treated diabetic bones. No significant differences

vi

between the 3 groups were seen in trabecular μCT measurements. Diabetic bone treated

with either vehicle or naltrexone had significantly lower cross-sectional areas and cortical

areas. No gross morphological differences were detected after 21 days of naltrexone

therapy.

In summary, naltrexone is effective in enhancing full-thickness cutaneous wound

healing and early phases of fracture repair, and is able to change mechanical properties of

diabetic bone. Naltrexone is comparable to the current standard of care for non-healing

wounds, Regranex®, and has been proven to be safe in multiple tolerability studies. Local

NTX can decrease granulation tissue in diabetic fracture calluses and increase cartilage and

bone volume. Systemic naltrexone can increase diabetic bone strength and reduce OGFr

expression.

vii

TABLE OF CONTENTS LIST OF FIGURES ………………………………..……………………………………………………………. xi LIST OF TABLES …………………………………..………………………………………………………….. xii ABBREVIATIONS .………………………………..………………………………………………………….. xiii ACKNOWLEDGEMENTS .……………………….………………………………………………………… xvi

CHAPTER 1: INTRODUCTION …………………………………………………………………………. 1 1.1. Diabetes ……………….………………………………………………………………………… 2

1.2. Animal Models ……………………………………………………………………………….. 2 1.3. Cutaneous Wound Healing ……………………….……………………………………. 5

1.3.1. Cellular and Molecular Basis of Full-Thickness Wound Healing ……… 5 1.3.1.1. Hemostasis ....………………………………………………………………. 5 1.3.1.2. Inflammation ..……………….…………………………………………….. 6 1.3.1.3. Proliferation ....……………………………………………………………… 7 1.3.1.4. Remodeling ….……………………………………………………………… 8 1.3.2. Cutaneous Healing Complications, Amputation, and Diabetes ……… 8 1.3.3. Current Treatments in Wound Healing …………………………………………. 9 1.4. Bone Growth and Remodeling ……………………………………………………….. 12 1.4.1. Cellular and Molecular Basis of Fracture Healing …………………………… 12 1.4.1.1. Inflammation ……………………………………………………………….. 12 1.4.1.2. Soft Callus Formation …………………………………………………… 13 1.4.1.3. Hard Callus Formation …………….…………………………………… 14 1.4.1.4. Bone Remodeling ……………………………………….………………… 15 1.4.2. Bone Healing Complications and Fractures in the Diabetic

Population ………………………………………………………………………………………………. 15

1.4.3. Current Treatments in Fracture Repair ………………………………………….. 18 1.5. Opioids …………………………………………..………………………………………………. 19 1.6. The OGF-OGFr Axis ..………………………………………..……………………………… 21 1.7. Opioid Receptor Antagonists ..……………………………………………………..… 21 1.7.1. Naltrexone …………………………………………………………………………………….. 21 1.7.1.1. Function of Naltrexone Blockade Actions ……………………… 22 1.7.2. Naloxone ……………………………………………………………………………………….. 24 1.8. Current Gap in Knowledge ..……………………………………………………………. 26 1.9. Hypothesis and Specific Aims .………………………………………………………… 26 1.10. References ………..………………………..…………………………………………………… 28

viii

CHAPTER 2: TOPICAL NALTREXONE IS A SAFE AND EFFECTIVE ALTERNATIVE TO STANDARD TREATMENT OF DIABETIC WOUNDS ………………………………………

53

2.1. Abstract ……………………………………………………………………………………………….. 54 2.1.1. Objective ……………………………………………………………………………………….. 54 2.1.2. Approach ………………………………………………………………………………………. 54 2.1.3. Results …………………………………………………………………………………………… 54 2.1.4. Innovation and Conclusion ……..…………………………………………………….. 55 2.2. Introduction ………………………………………………………………..……………………… 55 2.3. Clinical Problem Addressed ……………………………………………………..…………. 57 2.4. Materials and Methods ……………………………..………………………………………… 58 2.4.1. Animals and Induction of Diabetes ………………………………………………… 58 2.4.2. Cutaneous Wound Surgery ……………………………………………………………. 59 2.4.3. Wound Treatment and Closure ……………………………………………………… 59 2.4.4. BrdU-Labeling and DNA Synthesis …………………………………………………. 60 2.4.5. Histological Analysis and Immunohistochemistry ………………………….. 60 2.4.6. Statistical Analysis …………………………………………………………………………. 61 2.5. Results …………………………………………………………………………………………………. 61 2.5.1. Body Weight and Blood Glucose Measurements …………………………… 61 2.5.2. Full-Thickness Wound Closure .……………………………………………………… 62 2.5.3. Histological Analysis of Skin .………………………………………………………….. 64 2.5.4. DNA Labeling Indexes .…………………………………………….…………………….. 64 2.5.5. Tissue Pathology and Immunohistochemistry .………………………………. 66 2.5.6. PDGF Expression ….………………………………………………………………………… 66 2.5.7. VEGF Expression .…………………………………………………………………………… 67 2.5.8. FGF-2 Staining .………………………………………………………………………………. 68 2.6. Discussion .……………………..……………………………………………………………………. 69 2.7. Innovation .………………………………………………………………………………………….. 71 2.8. Key Findings …….………………………………………………………………………………….. 72 2.9. Acknowledgements and Funding Sources …………………………………………… 72 2.10. Author Disclosure and Ghostwriting ………………………………………….……… 72 2.11. Abbreviations and Acronyms ……………………………………………………..……… 72 2.12. References ..………………………………………………………………………………………. 74 2.13. About the Authors …………………………………………………………………………….. 79

ix

CHAPTER 3: THE OPIOID GROWTH FACTOR-OPIOID GROWTH FACTOR RECEPTOR AXIS AND DIABETIC BONE COMPOSITION …………………………………….

80

3.1. Abstract ……………………………………………………………………………………………….. 81 3.1.1. Background …………………………………………………………………………………… 81 3.1.2. Methods ………………………………………………………………………………………… 81 3.1.3. Results …………………………………………………………………………………………… 81 3.1.4. Conclusions …………………………………………………………………………………… 82 3.1.5. Keywords ………………………………………………………………………………………. 82 3.2. Abbreviations …………..…………………………………………………………………………. 82 3.3. Introduction ………………………………………………………………………………………… 83 3.4. Materials and Methods ……………………………..………………………………………… 85 3.4.1. Animals and induction of diabetes .……………………………………………….. 85 3.4.2. Bone fracture surgical model ………………………………………………………… 85 3.4.3. Immunohistochemistry …..…………………………………………………………….. 86 3.4.4. Histomorphometry …..…………………………………………………………………… 87 3.4.5. Measurement of serum enkephalin ..…………………………………………….. 87 3.4.6. Statistical analyses ..………………………………………………………………………. 87 3.5. Results …………………………………………………………………………………………………. 88 3.5.1. Expression levels of OGF and OGFr during bone development ………. 88 3.5.2. Levels of OGF and OGFr in bone and serum of T1D animals ..………… 89 3.5.3. Cellular composition of T1D bone …………………………………………………. 91 3.5.4. Cellular composition of T1D bone following fracture …………………….. 92 3.6. Discussion ……..……………………..……………………………………………………………… 97 3.7. Conclusion …………………………………………………………………………………………… 99 3.8. Acknowledgements .…………………………………….……………………………………… 99 3.9. Author Contributions .………………………………….……………………………………… 99 3.10. Funding …..………………..……………………………………………………………………….. 99 3.11. Competing Interests ………………………………………………………………………….. 99 3.12. References …………………………………………………………………………………………. 100

x

CHAPTER 4: THE OPIOID ANTAGONIST NALTREXONE PREVENTS BONE DEFECTS IN A DIABETIC RAT MODEL ………………………………………………………………

105

4.1. Abstract …………………………………………………………………………………………………… 106 4.1.1. Background ……………………………………………………………………………………….. 106 4.1.2. Methods ……………………………………………………………………………………………. 106 4.1.3. Results ………………………………………………………………………………………………. 106 4.1.4. Conclusions ……………………………………………………………………………………….. 106 4.1.5. Keywords ………………………………………………………………………………………….. 107 4.3. Introduction …………………………………………………………………………………………….. 108 4.4. Materials and Methods …………………………………………………………………………… 109 4.4.1. Animals, induction of diabetes, and treatment …………………………………. 109 4.4.2. Three-point bending …………………………………………………………………………. 110 4.4.3. MicroCT …………………………………………………………………………………………….. 110 4.4.4. Immunohistochemistry ……………………………………………………………………… 111 4.4.5. Histomorphometry ……………………………………………………………………………. 112 4.4.6. Data analyses ……………………………………………………………………………………. 112 4.5. Results …………………………………………………………………………………………………….. 112 4.5.1. Three-point bending …………………………………………………………………………. 112 4.5.2. MicroCT …………………………………………………………………………………………….. 113 4.5.3. Immunohistochemistry ……………………………………………………………………… 115 4.5.4. Histomorphometry ……………………………………………………………………………. 118 4.6. Discussion ……………………………………………………………………………………………….. 118 4.7. Conclusion ………………………………………………………………………………………………. 120 4.8. Acknowledgements …………………………………………………………………………………. 120 4.9. Author Contributions ………………………………………………………………………………. 120 4.10. Funding …………………………………………………………………………………………………. 120 4.11. Competing Interests ………………………………………………………………………………. 120 4.12. References …………………………………………………..………………………………………… 121 CHAPTER 5: DISCUSSION …………………………………………………..…………………………… 126 5.1. Discussion …………………………………..…………………………………………………………… 127 5.2. References …………………………………..………………………………………………………….. 140

xi

LIST OF FIGURES

Figure 1.1. Temporal overview of phases of wound healing, prominent cell

types, and actions …………………………………………………………………………

6

Figure 1.2. Temporal overview of phases of fracture repair with prominent

cell types ………………………………………………………………………………………

12

Figure 2.1. Residual wounds over time ………………………………..………………………… 63

Figure 2.2. BrdU labeling of cutaneous wounds ………………..…………………………… 65

Figure 2.3. PDGF labeling of cutaneous wounds ……………..…………………………….. 67

Figure 2.4. VEGF labeling of cutaneous wounds ……………..………………………….…. 68

Figure 3.1. OGF and OGFr staining in developing bone ..………………………………… 89

Figure 3.2. OGF and OGFr staining in normal and diabetic bone ………….………….. 90

Figure 3.3. Comparison of normal and diabetic bone .……………………..……………. 92

Figure 3.4. Fracture callus composition …………………………………………….……….….. 93

Figure 3.5. Fracture callus composition subperiosteum …………………….………….. 95

Figure 3.6. Ki67 labeling in fracture calluses .…………………………………………………. 96

Figure 4.1. Force, energy, and stiffness histograms ……………………………………….. 113

Figure 4.2. OGF and OGFr staining in bone ……………………………………………………. 116

Figure 4.3. Osteocalcin staining in bone ………………………………………………………… 117

Figure 4.4. Ki67 staining in bone ……………………………………………………………………. 118

xii

LIST OF TABLES 4.1. Trabecular μCT measurements ………………………………………………………………….. 114

4.2. Cortical μCT measurements ………………………………………………………………………. 114

xiii

ABBREVIATIONS

AAALAC, Association for Assessment & Accreditation of Laboratory Animal Care

ACTH, adrenocorticotropic hormone

AGE, advanced glycation end products

ANOVA, analysis of variance

ATP, adenosine triphosphate

BB, biobreeding

β-end, β-endorphin

BMD, bone mineral density

BMP, bone morphogenetic proteins

BrdU, 5-bromo-2’-deoxyuridine

BV/TV, bone volume/total volume

cDNA, complementary deoxyribose nucleic acid

CLIP, corticotropin-like intermediate lobe peptide

CN, Charcot neuroarthropathy

DAPI, 4’,6-diamidino-2-phenylindole

DB, type 1 diabetic

DB/R, diabetic Regranex®-treated

DB/NTX, diabetic naltrexone-treated

DFU, diabetic foot ulcer

DKK1, dickkopf-related protein 1

Dlx5, distal-less homeobox 5

xiv

DNA, deoxyribose nucleic acid

ER, endoplasmic reticulum

ELISA, enzyme-linked immunosorbent assay

FDA, Food and Drug Administration

FGF, fibroblast growth factor

FOXO, forkhead box O

GK, Goto-Kakizaki

HDN, high dose naltrexone

hIAPP, human islet amyloid polypeptide

IACUC, Institutional Animal Care and Use Committee

IGF, insulin like growth factor

IL-1, interleukin-1

INSR, insulin receptor

ip, intraperitoneal

LCMV, lymphocytic choriomeningitis virus

LDN, low dose naltrexone

M-CSF, macrophage-colony stimulating factor

MMP, matrix metallopeptidase

mRNA, messenger RNA

MSC, mesenchymal stem cells

MSH, melanocyte stimulating hormone

NAD+, nicotinamide adenine dinucleotide

xv

NOD, non-obese diabetic

NTX, naltrexone

N, non-diabetic

OGF, opioid growth factor

OGFr, opioid growth factor receptor

PARP, poly(ADP-ribose) polymerase

PDGF, platelet derived growth factor

PENK, proenkephalin

PlGF, placental growth factor

POMC, proopiomelanocortin

RANKL, receptor activator of NFκB ligand

ROS, reactive oxygen species

RUNX2, runt-related transcription factor 2

SEM, standard error of the mean

SOX9, SRY-box 9

STZ, streptozotocin

TGF, transforming growth factor

TNF-α, tumor necrosis factor alpha

TRAP, tartrate resistant alkaline phosphatase

T1D, type 1 diabetic

T2D, type 2 diabetes

VEGF, vascular endothelial growth factor

xvi

ACKNOWLEDGEMENTS I would like to express my gratitude to my dissertation advisor, Dr. Patricia McLaughlin,

for her guidance and support throughout the dissertation process. I would also like to

thank my committee for their helpful input on this project.

Lastly, I would like to thank Gary, Elaine, and Marci Titunick for their unconditional

support and encouragement throughout my studies.

1

CHAPTER 1: INTRODUCTION

2

1.1. Diabetes

In 2015 an estimated 9.4% of the U.S. population had diabetes, 5-10% of whom

had type 1 diabetes. The total estimated healthcare cost of both types of diabetes in the

U.S. in 2017 was $327 billion[1]. Type 1 diabetes is defined by a lack of insulin production.

This autoimmune disease causes the insulin-producing beta cells of the pancreas to be

degraded and individuals with diabetes cannot control their blood glucose levels. A

fasting glucose level over 126 mg/dL with symptoms of hyperglycemia such as polydipsia,

polyphagia, and polyuria is sufficient for a diagnosis of diabetes[2]. In type 2 diabetes

individuals cannot utilize the insulin made by the pancreas leading to cells lacking energy.

Often type 2 diabetes can be controlled by lifestyle changes; eating well and exercise[3].

Complications arising from both types of diabetes include retinopathy, nephropathy,

neuropathy, cardiovascular disease[4], poor wound healing[5], and delayed fracture

healing[6].

1.2 Animal Models

Animals can be induced to have either type 1 or type 2 diabetes. Type 1 diabetes

models are preferentially used to eliminate some of the complications caused by the

comorbidities of type 2 diabetes such as increased adiposity. Chemically induced type 1

diabetes can be produced by injections of either streptozotocin (STZ) or alloxan at least

5-7 days prior to detection of hyperglycemia[7]. The disadvantage of using chemically

induced diabetes is that it can be toxic at other organs including the liver, kidney, lung,

intestines, and brain[8], and does not mimic human pathology.

3

STZ, [2-deoxy-2-(3-(methy-3-nitrosoureido)-D-glucopyranose], is synthesized

from Streptomycetes achromogenes and is administered either intraperitoneally or

intravenously. It enters pancreatic beta cells by way of the Glut-2 transporter, alkylates

the DNA[9] activating PARP leading to NAD+ depletion, a reduction in cellular ATP, and

finally a decrease in insulin production[10]. STZ also creates free radicals causing DNA

damage and cell death[7]. It can be injected as either a single high dose or multiple low

doses. STZ has been used successfully to test cutaneous wound healing in numerous

studies[11–13].

Alternatively, alloxan (2,4,5,6-tetraoxypyrimidine; 5,6-dioxyuracil) is quickly taken

up by pancreatic beta cells where free radicals are formed to which the beta cells are

especially susceptible to damage[14]. It can be administered intraperitoneally,

subcutaneously, or at a lower dose intravenously. The disadvantage to alloxan is that a

slight overdose can lead to general toxicity[9].

The most common autoimmune models of type 1 diabetes are the non-obese

diabetic (NOD) mouse and the Biobreeding (BB) rat. NOD mice have a prediabetic phase

that can be used to examine preventative treatments. These mice require insulin

treatment and many drugs that work in diabetes prevention in NOD mice do not translate

to humans. Additional limitations to this model include the requirement of a specific

pathogen-free space, an unpredictable onset of diabetes, and high maintenance

expenses[7]. BB rats derived from Wistar rats also develop diabetes spontaneously. They

require insulin therapy and develop lymphopenia which is not characteristic of type 1

4

diabetes in humans. These rats have been used for transplant and interventional studies

as well as studies on diabetic neuropathy[7].

Type 1 diabetes can be genetically induced in the AKITA mouse. In this mouse

model beta cells are destroyed due to ER stress and the animals are insulin dependent.

This model is useful for testing new insulin, transplantation, and treatments involving ER

stress. Virally-induced type 1 diabetes utilizes coxsackie B, encephalomyocarditis, or

Kilham rat viruses. Lymphocytic choriomeningitis virus (LCMV) can also be used under

the rat insulin promoter. In this case, beta cells are destroyed due to viral infection or are

involved in an immune response when exposed to LCMV. This model is complex and is

used mainly to detail the extent to which viruses can cause diabetes. In large animals,

pancreatectomy is the primary technique for inducing diabetes[7].

Type 2 diabetes can be induced genetically or by controlling food intake. Obese

monogenetic models like ob/ob mice deficient in leptin or db/db mice and Zucker diabetic

fatty rats deficient in leptin receptors are commonly used. They develop obesity-induced

hyperglycemia and can be used in studies to improve insulin resistance and beta cell

function[7]. db/db mice have also been used in studying corneal complications related to

diabetes[15]. Obese polygenic models are also available and have obesity-induced

hyperglycemia. Obesity can be induced by feeding rodents high fat diets or using animals

like the Nile grass rat and desert gerbil which naturally become obese in captivity. Non-

obese models of type 2 diabetes include the Goto-Kakizaki (GK) rat where hyperglycemia

is induced by insufficient beta cell mass or function. Genetically induced models of beta

cell dysfunction include the hIAPP mouse and the AKITA mouse. hIAPP mice produce

5

human islet amyloid polypeptide which creates amyloids in pancreatic beta cells. AKITA

mice as mentioned earlier have beta cell destruction due to ER stress[7].

The STZ-induced type 1 diabetic rat model has been successfully used in studies

looking at complications resulting from type 1 diabetes. In addition, the model is less

expensive than breeding colonies of genetically induced mice or mice that are

immunocompromised.

1.3 Cutaneous Wound Healing

1.3.1 Cellular and Molecular Basis of Full-Thickness Wound Healing

1.3.1.1 Hemostasis

Understanding the cellular and molecular basis of full-thickness wound healing

allows for investigation into complications associated with diabetes. Wound healing

begins at the time of injury with hemostasis (figure 1.1.)[16]. Vascular injury during

wounding initiates the formation of a fibrin-fibronectin clot. Fibrillar collagens type I and

II promote aggregation of platelets into a clot with platelets embedded among the fibers.

The clot acts as a hemostatic plug protecting the wounded region, facilitates cell

migration, and stores growth factors and cytokines which aid in inflammatory cell

recruitment[17].

6

Figure 1.1.: Temporal overview of phases of wound healing, prominent cell types, and

actions; from Baltzis et al. 2014[16]

1.3.1.2 Inflammation

In the second phase of wound healing, neutrophils and monocytes are recruited

by various chemotactic signals (e.g. cytokines and growth factors) including tumor

necrosis factor alpha (TNF-α), interleukin (IL)-1, and platelet derived growth factor

(PDGF). Neutrophils attack bacteria while monocytes differentiate into macrophages and

phagocytose pathogenic organisms and debris[17]. Adherence to the extracellular matrix

7

promotes colony-stimulating factor 1, tumor necrosis factor α, and platelet-derived

growth factor secretion by monocytes and macrophages. Colony-stimulating factor 1 is

necessary for monocyte and macrophage survival. Tumor necrosis factor α is an

inflammatory cytokine. Platelet-derived growth factor (PDGF) is a chemoattractant and

supports proliferation of fibroblasts[18]. Macrophages secrete transforming growth

factor alpha (TGF-α), fibroblast growth factor (FGF), and vascular endothelial growth

factor (VEGF). These factors initiate the proliferation process[17].

1.3.1.3 Proliferation

Reepithelialization begins within hours of injury[18] and is stimulated by cytokines

released by activated fibroblasts and keratinocytes[17]. There is an increase in cell

proliferation and migration. Migration is made possible by the dissolution of

hemidesmosomes[18] and concludes when there is a single layer of keratinocytes

covering the wound area and the stratified epidermis is re-established[17].

Granulation tissue infiltrates the wound area around day 4 post-injury[18] and

contains capillaries which grow into the wounded region and undergo angiogenesis[17].

Angiogenesis is promoted by cytokines such as FGF-2, VEGF, PDGF, and TGF-β1,2.

Fibronectin and hyaluronan are deposited. Fibronectin allows for initiation of collagen

fibrillogenesis and acts as an anchor for wound contraction. Hyaluronan forms a matrix

that allows for cell migration. This preliminary matrix is then replaced by collagen, and

the cellularity decreases as the granulation tissue matures[17].

8

1.3.1.4 Remodeling

Fibroblasts deposit and remodel the extracellular matrix. Collagen is deposited

causing an increase in strength. Contraction involves pulling normal dermal and adipose

tissue into the wound region[17]. Remodeling may last up to two years or more[19].

1.3.2 Cutaneous Wound Healing Complications, Amputation, and Diabetes

Five out of 1,000 diabetics require a lower extremity amputation[20]. More than

half of non-traumatic limb amputations occur in patients with diabetes, usually following

a non-healing wound or ulcer[21]. Neuropathy and vascular issues contribute to the

production of non-healing wounds and ulcers. Peripheral neuropathy is a leading cause

of foot ulcers as patients do not feel the discomfort or pain associated with an injury[22].

Autonomic neuropathy results in reduced sweating causing dry skin and fissures in

diabetics[23], creating an entry point for bacterial and fungal infections.

Diabetics are at twice the risk for peripheral vascular disease compared to non-

diabetics[5]. Red blood cells in diabetics are less deformable causing complications in

capillaries[24] and high blood glucose levels increase blood viscosity[25]. These two

conditions cause blood to remain stagnant in vessels. Peripheral arterial disease is caused

by atherosclerosis where fatty deposits form plaques within arteries which eventually

occlude the lumen[22]. Calcification leading to stroke and cardiovascular disease is also

common in diabetes[26]. Resultant reduced mobility may put patients at greater risk for

ulcers. Diabetic microangiopathy exhibits thickened capillary basement membranes

leading to decreased oxygen and nutrient delivery to tissues[5]. Although hypoxia is

necessary for vascularization, when oxygen levels do not eventually elevate fibroblasts

9

cannot produce collagen[27]. Leukocytes have difficulty reaching the wound site due to

narrowed vessels[28,29] and may also be less active[28].

Diabetics have a prolonged inflammatory phase preventing granulation tissue

formation[22]. Proteases that remain can cause damage to the new extracellular

matrix[5]. Inflammatory cytokines also cause a reduction in proliferative factors[22] and

make fibroblasts less responsive to growth factors[30]. It is suggested that bacteria thrive

in wounds with high blood glucose levels and that the increase in glucose may also be

detrimental to neutrophil action[31,32]. Often the classic signs of infection are absent or

diminished in diabetics due to neuropathy and ischemia[33].

1.3.3 Current Treatments in Wound Healing

Standard care of diabetic ulcers includes debridement, dressing, pressure off

loading, and infection management. Debridement removes callus, necrotic dermal tissue,

foreign debris, and bacteria; all of which retard wound healing. Sharp debridement is the

gold standard of care and allows the wound to respond better to topical treatments[16].

Wound dressings must protect the wound from secondary infections, keep the wound

moist, remove exudates, and promote tissue regeneration. Research has not shown a

specific type of dressing to be significantly better than another[16]. Removing pressure

from the ulcer is another important aspect to adequate wound care. Casts can be used

to redistribute pressure or surgical methods such as Achilles tendon lengthening,

metatarsal-phalangeal joint arthroplasty, metatarsal head resection, and liquid silicone

injections plantar to metatarsal heads may be utilized. Research suggests there is very

little gained by surgical methods compared to casting methods[16].

10

To assist in adequate vascularization surgical therapies including angioplasty,

endarterectomy, grafting, or bypass may be considered[16]. Infection is managed by

debridement and antibiotics. Improvements of the patient’s general health will also

contribute to better outcomes. Keeping blood glucose levels under control, managing

dyslipidemia, smoking cessation, and appropriate diet will aid healing[34,35].

Adjunctive therapies include bioengineered skin substitutes, growth factors,

oxygen therapy, negative pressure wound therapy, and electrical stimulation and

shockwaves. Currently, there are two US Food and Drug Administration (FDA) approved

bioengineered skin substitutes. Apligraf® which is composed of cultured living dermis and

epidermis from neonatal foreskin with extracellular matrix, dermal fibroblasts, epidermal

keratinocytes, and a stratum corneum has an extracellular matrix lattice made of bovine

type I collagen. The dermal fibroblasts and keratinocytes produce growth factors. The

stratum corneum is a physical barrier against mechanical damage and infection.

Dermagraft® is composed of neonatal-derived dermal fibroblasts on bioabsorbable

polyglactin mesh. The fibroblasts secrete extracellular matrix and along with growth

factors such as PDGF-A, insulin-like growth factor, keratinocyte growth factor, heparin-

binding epidermal growth factor, transforming growth factors, and VEGF[16] is used to

promote healing.

Platelet-derived growth factor (PDGF), or becaplermin, has been approved by the

FDA for treatment of diabetic foot ulcers under the name Regranex®. PDGF reportedly

activates inflammatory cells, stimulates cell proliferation and migration, and enhances

11

protein and extracellular matrix synthesis[16]. However, there is a black box warning on

Regranex® indicating an increased risk of cancer mortality[36].

Another, more experimental method of wound healing utilizes hyperbaric oxygen

therapy that alters microbial balance, soft tissue infection, and angiogenesis[16].

Negative pressure wound therapy exposes the wound to subatmospheric pressure[37] to

assist in the removal of excess fluid[38,39], reduce bacterial load, reduce mechanical load,

or increase granulation tissue production[40]. The results have been inconclusive as to

its effects[16]. Pulsed electromagnetic field stimulation may decrease fibroblast and

endothelial cell doubling time in culture, have bacteriostatic and bactericidal properties,

increase migration of neutrophils and macrophages[41], and increase expression of

angiogenesis-related growth factors[42]. There are few clinical trials on electrical

stimulation[43].

Future directions in wound therapy include stem cell transplants, gene therapy,

neuropeptide-based treatments, and cytokine inhibition. Intramuscular injections of

peripheral blood mononuclear cells, bone marrow mesenchymal stem cells, or bone

marrow-derived mononuclear cells have increased wound healing[44,45]. Local

application of both bone marrow cells[46] and bone marrow mesenchymal stem cells[47]

have improved wound healing. Topical application of a replication-defective adenovirus,

encoding PDGF, accelerated wound healing in diabetic patients[48]. VEGF has been

administered using adenovirus in animal models with promising results[49].

Studies have suggested angiotensin II and angiotensin (1-7) play a role in wound

healing and have used norleu3-angiotensin (1-7), an angiotensin (1-7) analog, to enhance

12

wound healing in patients[50–52]. Topical substance P accelerated wound healing,

increased contraction, and increased levels of VEGF, TGF-β1 and TNF-α[53]. Another

treatment being explored is recruitment and activation of anti-inflammatory

macrophages. This will promote an anti-inflammatory environment conducive for growth

factors and wound healing[54].

1.4. Bone Repair and Remodeling

1.4.1 Cellular and Molecular Basis of Fracture Healing

1.4.1.1 Inflammation

Figure 1.2.: Temporal overview of phases of fracture repair with prominent cell types,

from Einhorn and Gerstenfeld, 2015[55]

Fracture repair greatly mirrors the wound repair process. Both processes begin

with an inflammatory phase and rely healivy on vascularzation. Immediately following

traumatic bone fracture, bleeding causes the formation of a hematoma (figure 1.2.) which

is invaded by platelets and macrophages. These inflammatory cells secrete factors that

13

foster clotting. Capillaries advance into the clot which is now becoming granulation

tissue. Multiple factors, including platelet-derived growth factor (PDGF), fibroblast

growth factor-2 (FGF-2), and vascular endothelial growth factor (VEGF), are released to

recruit more inflammatory cells and mesenchymal stem cells[56]. PDGF is secreted by

platelets, monocytes, macrophages, endothelial cells, and osteoblasts, and stimulates

proliferation and migration of osteoblasts and mesenchymal stem cells (MSCs). Fibroblast

growth factor is secreted by monocytes, macrophages, mesenchymal cells, osteoblasts,

and chondrocytes, and supports angiogenesis and mesenchymal cell proliferation. Acidic

FGF regulates chondrocyte proliferation[57] while basic FGF enhances proliferation of

osteoblasts and prevents their apoptosis[58]. Insulin like growth factor, secreted by bone

matrix, endothelial cells, osteoblasts, and chondrocytes, promotes bone matrix formation

and cell proliferation[57]. Bone morphogenetic proteins (BMPs) are secreted by

osteoprogenitor cells, mesenchymal cells, osteoblasts, and chondrocytes. BMPs are

chemotactic factors that also regulate mesenchymal and osteoprogenitor cell

proliferation and differentiation, enhance angiogenesis by activating endothelial cells,

promote extracellular matrix synthesis, and stimulate the synthesis and secretion of

insulin like growth factor (IGF) and VEGF to support angiogenesis[57].

1.4.1.2 Soft Callus Formation

Fractures heal through the process of endochondral ossification when there is

mechanical instability. A callus is formed first from cartilage and later by bone giving

stability to the fractured ends of the bone. The invading mesenchymal cells differentiate

into chondrocytes. The cartilage bridges the fracture increasing support. Chondrogenesis

14

is supported by bone morphogenetic proteins. Chondrocytes secrete extracellular matrix

proteins, especially collagen II. The cartilaginous callus is then invaded by vasculature

stimulated by pro-angiogenic factors[56]. Proliferating chondrocytes increase levels of

SOX9 (SRY-box9)[59]. Hypertrophic chondrocytes increase levels of runt-related

transcription factor 2 (RUNX2), alkaline phosphatase, collagen I and X, matrix

metallopeptidase 13 (MMP13), VEGF, osteocalcin, osterix, osteopontin, PDGF, BMPs, and

placental growth factor (PIGF). RUNX2 is associated with osteoblast differentiation from

MSC to preosteoblast[60]. Alkaline phosphatase correlates to osteoblast activity.

MMP13 degrades collagen II and aggrecan in the extracellular matrix[59]. Osterix is

required for differentiation from preosteoblast to mature osteoblast, activating DKK1, a

WNT antagonist[60]. Osteocalcin is pro-osteoblastic and stimulates matrix

mineralization. Osteopontin is a non-collagenous component of the extracellular

matrix[61].

1.4.1.3 Hard Callus Formation

High osteoblast activity and mineralized bone matrix formation are key

characteristics of hard callus formation[56]. MMP9 degrades collagen[59] and the soft

callus is removed and replaced by irregular bone[56]. This initial woven bone matrix is

created by mature osteoblasts. BMPs play an important role in differentiation from

osteoprogenitor cell to mature osteoblast. High levels of BMPs inhibit osteocalcin and

osteopontin[59]. Although stem cells from the periosteum and bone marrow can

produce bone, they are not necessary for bone formation. This indicates that

15

osteoprogenitors may originate from other locations such as circulation[62],

vasculature[63], and local tissue[56,64].

1.4.1.4 Bone Remodeling

In the final stage of fracture repair woven bone is formed into organized lamellar

bone. The osteoclast, a large multinucleated cell originating from a hemopoietic

precursor, resorbs bone. A border is sealed off and acid and proteinases are released

allowing for demineralization and degradation of organic components. The debris is

accumulated in vesicles, and the osteoclasts are apoptosed or resume a non-resorbing

form. Osteoblasts migrate to the region and lay down new bone. Osteoblasts secrete

macrophage-colony stimulating factor (M-CSF) and receptor activator of NFκB ligand

(RANKL) to promote osteoclasts survival and activity. RANKL is vital in coordinating the

balance of bone formation and resorption[56].

1.4.2 Bone Healing Complications and Fractures in the Diabetic Population

Diabetes can decrease linear bone growth in adolescence[65], reduce bone

mineral density (BMD)[66], increase fracture risk[67], and result in poor osseous

healing[68]. Type 1 diabetes has been associated with decreased osteoblast recruitment

and activity. Studies have shown unaltered or decreased bone resorption in experimental

diabetes. Experiments have also shown “poor trabecular connectivity, increased porosity

and lower bone spicule/marrow space ratio[69].” Collagen reduction was noted within 2

weeks of diabetes induction. The fracture callus maturation time is prolonged by 87%,

with bone bridging delayed by 40%[69]. The callus showed reduced undifferentiated

mesenchymal cells at 4 days with delayed differentiation[69]. DNA content within a

16

diabetic callus was 40% less compared to normal calluses indicating decreased cell

proliferation[70].

Diabetics are at risk for developing Charcot neuroarthropathy (CN) whereby

bones, joints, and soft tissue of the foot and ankle are susceptible to inflammation and

eventual deformation. The Charcot foot is subject to fractures and dislocations as the

bone deteriorates. Loss of protective sensation in diabetes prevents patients from

noticing a plantar foot injury resulting in prolonged weight-bearing insult, perpetuating

inflammation. Motor neuropathy causes tendon contractures exacerbating deforming

forces on the foot. Autonomic neuropathy prevents regulation of peripheral circulation,

increasing blood flow, and resulting in a bounding pulse. A common feature of CN is

“rocker bottom foot,” a collapse of the midfoot joint[71].

It is possible that increased levels of tumor necrosis factor-α (TNF-α), a

proinflammatory mediator, may prevent diabetics from restricting levels of other

regulatory factors[72]. Glycation of type I collagen decreases osteoblast ability to adhere

to extracellular matrix and lowers alkaline phosphatase activity[73]. Serum alkaline

phosphatase and osteocalcin, indicators of osteoblasts and osteoblastic activity, are lower

in T1D animals than in non-diabetic animals. Both STZ-induced diabetic mice and

nonobese spontaneously diabetic mice have decreased trabecular bone in the tibia[74].

In vitro studies suggest high glucose and advanced glycation end products (AGEs)

are associated with inhibited osteoblast function and decreased mineralized matrix

formation[75]. Experimentally AGE treatment caused a dose-dependent inhibition in

bone healing amongst non-diabetic animals[76,77]. Reactive oxygen species (ROS) are

17

formed due to either high glucose levels and/or insulin insufficiency. High ROS levels

reportedly inhibit differentiation of osteoblasts[78]. Bone may also be affected by poor

expression of genes such as Cbfa1/Runx2 and Dlx5 which regulate osteoblast

differentiation[79]. High levels of Forkhead box O (FOXO), induced by oxidative stress,

antagonizes Wnt signaling and decreases bone formation[80]. A lack of insulin can also

have an effect on proliferation. Insulin acts through the insulin receptor (INSR) to increase

cell proliferation and cell growth through mitogen-activated protein kinases[81].

During fracture repair there is a decrease in growth and angiogenic factors,

reduced proliferation, and increased apoptosis[82]. Long bone fractures in STZ animal

models form smaller calluses with reduced bone composition (less bone per area of

callus) and decreased mechanical strength[70,83,84]. Biomechanical strength was

decreased in animal models by 20% in femurs and tibias[85,86]. A study[87] involving

ectopic bone formation in rats noted that cell proliferation in diabetic rats was 35% that

of non-diabetic controls. Systemic insulin raised cell proliferation to 81% of controls. It

is important to note that systemic administration of insulin may result in

hypoglycemia[81] and may not be a preferred treatment.

Often low oxygen levels are associated with diabetes. Ischemia can lead to cell

death, delayed chondrocyte and osteoblast differentiation, and poor fracture healing[88].

Oxygen is necessary for aerobic metabolism, enzymatic activities, collagen synthesis, as a

signaling molecule, and potentially for stem cell maintenance[89]. Hyperbaric oxygen

enhances cell proliferation and mineralization of osteoblasts[90]. Oxygen “decreases

18

sclerostin expression, increases Wnt signaling, and increases BMP2, IGF, and VEGF

expression[89].”

1.4.3 Current Treatments in Fracture Repair

Maintaining healthy insulin levels does not completely ameliorate bone loss[91].

At this time, standard of care for diabetic patients receiving surgical treatment for a

fracture include internal fixation and prolonged off-loading. Biologics have begun to be

applied in cases were a patient is at higher risk of non-union such as diabetics and

smokers. These include platelet-derived growth factor, platelet-rich plasma, bone

morphogenetic proteins 2 and 7, and demineralized bone matrix. As these treatments

are not fully integrated into standard practice there is still room for improvement and for

a disease modifying treatment. Experimentally, local delivery of insulin[92],

vanadium[93], and manganese chloride[94] have all shown to improve fracture healing in

a rat model. Local insulin may inhibit FOXO1 and therefore prevent Wnt signaling

antagonism. Vanadium[93] and manganese chloride[94] are insulin-mimetic and

osteogenic. Basic fibroblast growth factor applied to fractures normalized diabetic repair

and enhanced healing in non-diabetic animals[95]. In another study using STZ-induced

diabetic rats there was a dose-dependent effect on callus formation 3 weeks post-

fracture. Increases in mechanical properties mirrored the histological and radiological

changes[95]. Low intensity pulsed ultrasound treatment improved levels of growth

factors and increased callus cartilage and blood vessel density[96]. Injection of platelet-

rich plasma which contains high levels of mitogenic factors normalized fracture repair by

increasing cell proliferation in diabetic rats[97]. Recombinant human bone

19

morphogenetic protein-2[98] and PDGF-BB[99] have both been used to enhance diabetic

fracture healing in rat models.

Most of the new platforms for treatment are focused on treating the diabetic

complications only and not altering the disease. Disease modifying treatments would be

beneficial and focus on multiple complications of diabetes. It is thought that the

dysregulation of the OGF-OGFr axis may cause disruption in bone healing and that

blockade of the pathway would treat the underlying pathophysiology of diabetes.

1.5. Opioids

Opioids function in many capacities including analgesia, cardiovascular control,

behavior, and cell division and growth[100], and can be endogenous or exogenous in

nature. Endogenous opioids regulate stress responses and are found in the

hypothalamus, pituitary, and adrenal medulla. Precursors for endogenous opioid

peptides include beta-endorphin/ACTH precursor (also known as

proopiomelanocortin/POMC), proenkephalin/proenkephalin A, or dynorphin/neo-

endorphin precursor (also known as prodynorphin/proenkephalin B). The beta-

endorphin (β-END)/ACTH precursor gives rise to beta-endorphin and adrenocorticotropic

hormone (ACTH). ACTH (1-39) can be cleaved into melanocyte stimulating hormone

(MSH) and CLIP (Corticotropin-Like Intermediate Lobe Peptide) or ACTH (18-39).

Proenkephalin produces several copies of met-enkephalin and leu-enkephalin, whereas

prodynorphin gives rise to α/β-neo-endorphin, dynorphin A, and dynorphin B. The

pituitary is the major site of β-END/ACTH synthesis and secretion, but proenkephalin is

localized in a wide variety of tissues spanning all three germ layers. Dynorphin is

20

produced in the gut, posterior pituitary, and brain[101]. The endogenous opioid of

interest in this dissertation is [met5]-enkephalin, a pentapeptide with the sequence Tyr-

Gly-Gly-Phe-Met[100]. [Met5]-enkephalin expression appears to be conserved from

bacteria to humans[102,103]. It also goes by the name opioid growth factor (OGF) and

decreases cell proliferation without increasing apoptosis or necrosis and does not affect

cell differentiation[100].

Insulin and glucagon secretion is stimulated by β-END and inhibited by [met5]-

enkephalin[104]. Plasma [met5]-enkephalin levels were reported to be significantly

higher in type 1 diabetics compared to normal individuals[105].

Proenkephalin (PENK), the precursor to [met5]-enkephalin, is expressed in embryonic

mesenchymal tissues of cartilage and bone[106]. Fetal rat calvaria-derived cells and

osteoblast-like cells grown in culture also tested positive for PENK mRNA[107]. Elhassan

et al.[108] demonstrated the presence of PENK-derived peptides in bone using

immunohistochemistry. [Met5]-enkephalin appeared in both mature bone matrix and in

osteoblasts. PENK declines with age[109]. There is an inverse relationship between

osteoblastic PENK mRNA and cell differentiation. As levels of PENK decrease, alkaline

phosphatase activity increases. [Met5]-enkephalin reduced alkaline phosphatase activity

in ROS17/2.8, an osteoblastic cell line[107]. Enkephalinase is expressed in

osteoblasts[110] and may moderate the levels of enkephalin to allow for activity during

particular stages of fracture healing[106].

21

1.6. The OGF-OGFr Axis

[Met5]-enkephalin, also termed opioid growth factor (OGF), is a negative growth

regulator delaying the G0/G1 phase of the cell cycle through interactions at the opioid

growth factor receptor (OGFr), originally termed opioid receptor zeta (ζ). It has been

shown to be the most potent endogenous opioid associated with growth, with

concentrations as low as 10-10M inhibiting cell proliferation[111]. The peptide-receptor

interaction is reversible with the peptide released in an autocrine manner that is tissue

and species non-specific[111]. OGF binds to OGFr creating a complex that translocates

into the nucleus resulting in an increase of the cell cycle inhibitors p16 and p21 and a

decrease in cell proliferation[111,112].

Biochemical and molecular studies of OGFr indicate that OGFr is present on a variety

of tissues and located on the nuclear membrane where it then translocates into the

nucleus. Molecular comparison of classical opioid recpetors and OGFr indicate that there

is no homology between the classical opioid receptors and OGFr. Fluorescence in situ

hybridization determined that OGFr is at human chromosome location 20q13.3. OGFr is

found predominantly in the perinuclear region of developing and renewing cells[111].

1.7. Opioid Receptor Antagonists

1.7.1. Naltrexone

Naltrexone (NTX) is a general opioid receptor antagonist that has no intrinsic

abilities except to block ligands from opioid receptors including μ, δ, and κ receptors. NTX

is a synthetic congener of oxymorphone[113,114]. The duration of receptor blockade by

naltrexone is important for determining the biological outcome. Intermittent blockade

22

by naltrexone causes feedback mechanisms to upregulate production and/or secretion of

more endogenous opioids that subsequently interact with the receptors allowing for

enhanced decreased cell replication. This paradigm is activated following low dose

naltrexone (LDN) therapy. LDN treatment is effective for cancer and inflammatory

disorders whereby the circulating levels of OGF are diminished[115,116]. Continuous

receptor blockade, often invoked by higher dosages of naltrexone, results in blockade of

the OGF inhibition of the cell cycle, and the result is an increase in cell replication. This

paradigm has been explored for the treatment of epithelial complications associated with

diabetes[117].

1.7.1.1 Function of Naltrexone Blockade Actions

Corneal explants grown in culture were used to examine growth of epithelium in

the presence of OGF and high dose naltrexone (HDN). Exogenous OGF decreased cell

division and led to disorganized outgrowth of the epithelium while NTX accelerated cell

division without changing the normal growth pattern[118,119]. OGF decreased DNA

synthesis and NTX increased DNA synthesis[119]. Corneal epithelial wound healing in rats

was increased by both systemic and topical NTX[120,121]. Rabbit corneal wounds also

healed more rapidly with blockade of the OGF-OGFr axis[122]. In a study using a Helios®

gene gun system to deliver sense cDNA or antisense cDNA for OGFr into corneal epithelial

cells, sense cDNA increased OGFr production and antisense decreased OGFr production.

Sense cDNA caused a delay in wound healing of rat corneas while those corneas treated

with antisense cDNA resulted in increased wound healing[123,124]. Animals were

treated with NTX for 1 week to determine its safety. Cells having undergone DNA

23

synthesis were increased by 69-85% with NTX treatment. Epithelial thickness and cellular

packing density also increased. There were no negative effects on the tissue. Organ

cultured human corneas with epithelial wounds were exposed to OGF and NTX resulting

in decreased cell proliferation in those treated with OGF and increased proliferation in

those treated with NTX[125].

Humans with both type 1 and type 2 diabetes often have complications associated

with the corneal surface include keratopathy and delayed corneal healing[126–128],

decreased corneal sensitivity[127–131], and dry eye[129,130,132,133]. Diabetic animal

models for type 1 diabetes were established, and rats received standardized corneal

epithelial wounds and a subset were treated intraperitoneally with NTX twice daily. There

was a significant increase in corneal reepithelialization in type 1 diabetic rats receiving

NTX compared to diabetic rats receiving saline. NTX normalized the rate of repair in

diabetic rats to the level of normal non-diabetic rats[134]. A similar study was used to

show that intensive insulin therapy would also be able to enhance the repair

process[135]. In another study, diabetic animals treated with topical insulin had wounds

19-60% smaller than those treated with vehicle only[136]. Topical NTX also increased the

rate of reepithelialization in diabetic rats with corneal wounds[137]. Investigations

combining topical NTX and topical insulin demonstrated that combined treatment was

not more effective than either insulin or NTX used alone[138] suggesting a similar

pathway or maximum potential to increase wound healing[100]. Investigations

examining use of NTX for treatment of corneal surface wound in the type 2 diabetic

model, db/db, revealed that the OGF-OGFr axis was present in this model and if

24

completely blocked by NTX, enhanced reepithelialization[15]. No pathology was

associated with NTX treatment[139].

Diabetes is often accompanied by peripheral neuropathy[140–145] and dry

eye[129,131,146]. Using von Frey hairs to test sensitivity, corneal sensitivity was

normalized in diabetic rats by treatment with topical NTX[147]. Treatment with NTX

normalized tear production for up to 48h. When treating one eye only there is no effect

seen in the contralateral eye, suggesting there is no cross over[147]. NTX did not affect

tear production in normal rats with normal tear volume.

Diabetes is also associated with slow cutaneous wound healing. Both normal and

type 1 diabetic rats treated three times daily with NTX had increased DNA synthesis[148].

Normal wounded rats treated with NTX had smaller residual wounds than control

animals. Diabetic rats treated with NTX also had smaller wounds than diabetic controls.

Moreover, there was no difference in skin histology between those animals treated with

NTX and control animals[148]. NTX helps improve cutaneous wound healing by

enhancing angiogenesis. There is a delay in FGF-2 and VEGF expression in wounded

diabetic animals[100]. Naltrexone increases formation and maturation of collagen and is

able to normalize tensile strength of healed skin[13]. The changes imparted by NTX in

type 1 diabetes have been seen in the db/db mouse model of type 2 diabetes as well[149].

1.7.2. Naloxone

Naloxone, also known as Narcan®, is most often used to treat opioid overdose and

improve respiratory depression[150]. Naloxone is shorter acting[117] and less

potent[151–153] relative to naltrexone. Naloxone, like naltrexone, has duration

25

dependent actions[154]. Studies determined that developing rats receiving naloxone for

less than 12 hours per day had decreased body and organ weight compared to

controls[154], suggesting a relationship between naloxone and organ growth.

Chick embryos received naloxone hydrochloride on multiple days[155]. Chicks

were sacrificed and femora were collected. Injection of naloxone resulted in a 2-fold

increase in thickness of the perichondrial bone cuff compared to controls. There was no

change in the density of osteoblasts in the diaphysis or mitotic activity in the growth plate.

There was a decrease in diaphyseal osteocyte density. Chicks treated during week 3 of

embryogenesis had an increase in the number of dividing cells in the growth plate and

the density of cells in the proliferating cartilage. Findings suggest that osteogenesis is

stimulated by naloxone[155].

A study utilizing a “drill hole” model examined the use of naloxone to enhance

mineralization and callus remodeling in sheep[156]. Treatment was administered

intramuscularly over 24h for four weeks. Naloxone enhanced mineralization and

remodeling especially when combined with calcium gluconate. Sheep receiving naloxone

had more radiodense drill holes than controls[156].

Mesenchymal stem cells (MSCs) and osteoblasts from human bone marrow were

cultured[157]. MSCs and osteoblasts did not express mRNA for μ- or κ-opioid receptor

nor POMC. They did express δ-opioid receptor, OGFr, and PENK. MTT assay was used to

determine proliferation in culture. Addition of [met5]-enk had no effect on cell

proliferation compared to control. Treatment with 1mM naloxone decreased the rate of

MSC proliferation and increased mineralization. There were no signs of apoptosis.

26

Osteocalcin gene expression increased after MSC culture treatment with naloxone. When

utilizing a unicortical defect model in mice researchers discovered treatment with

naloxone increased bone within the defect. There was an increase in relative bone

volume (BV/TV) corresponding to a 1.2-fold increase in trabecular number. Defect

diameter decreased 20.31% in naloxone treated defects compared to control. Treatment

with [met5]-enk resulted in no change compared to controls[157]. While studies have

shown how naloxone increases bone growth, testing has not been done using naltrexone.

1.8. Current Gap in Knowledge

The underlying causes of delayed fracture healing in diabetes are not known. Studies

in humans and animals, have reported that enkephalin levels are elevated in subjects with

diabetes[105,158]. Given that OGF (i.e. methionine enkephalin) is an inhibitory growth

factor known to depress cell replication, the dysregulation of the OGF-OGFr axis in

diabetics may be a factor in the delayed repair processes. Moreover, there is a potential

to modulate the axis to accelerate cell proliferation. Studies in animal models of type 1

and type 2 diabetes have reported that blockade of the OGF-OGFr axis with naltrexone

accelerates repair mechanisms in corneal epithelial wounds and full-thickness cutaneous

wounds. However, there is no information on the activity of this pathway in bone, or

whether the OGF-OGFr regulatory axis can be manipulated by naltrexone in bone growth

and repair.

1.9. Hypothesis and Specific Aims

The hypothesis of this research is that the OGF-OGFr axis is dysregulated in type 1

diabetes and that sustained blockade of the axis using naltrexone will reverse diabetic

27

complications including delayed repair of full-thickness wounds and bone fractures.

Specific Aim 1 will test the hypothesis that naltrexone administered topically to a full-

thickness cutaneous wound in type 1 diabetic rats will enhance the rate of wound closure

in a manner comparable to standard of care- Regranex®. Specific Aim 2 will test the

hypothesis that the dysregulation of the OGF-OGFr pathway alters markers within bone

in diabetic animal models, and that blockade of the OGF-OGFr pathway modulates the

repair of fractured femurs. Specific Aim 3 will test the hypothesis that systemic treatment

with naltrexone to block the OGF-OGFr pathway with the antagonist naltrexone may

prevent or change bone composition in diabetes that ultimately results in complications

related to fracture and repair.

28

1.10. References

[1] Yang, W.; Dall, T.M.; Beronjia, K.; Lin, J.; Semilla, A.P.; Chakrabarti, R.; Hogan, P.F.

Economic Costs of Diabetes in the U.S. in 2017; Diabetes Care; American Diabetes

Association, 2018.

[2] Atkinson, M.A.; Eisenbarth, G.S.; Michels, A.W. Type 1 Diabetes. The Lancet, 2014,

383, 69–82.

[3] Facts About Type 2 http://www.diabetes.org/diabetes-basics/type-2/facts-about-

type-2.html (accessed May 2, 2018).

[4] Nathan, D.M. Long-Term Complications of Diabetes Mellitus. N. Engl. J. Med.,

1993, 328, 1676–1685.

[5] Falanga, V. Wound Healing and Its Impairment in the Diabetic Foot. The Lancet,

2005, 366, 1736–1743.

[6] Hofbauer, L.C.; Brueck, C.C.; Singh, S.K.; Dobnig, H. Osteoporosis in Patients With

Diabetes Mellitus. J. Bone Miner. Res., 2007, 22, 1317–1328.

[7] King, A.J. The Use of Animal Models in Diabetes Research. Br. J. Pharmacol., 2012,

166, 877–894.

[8] Lee, J.; Yang, S.; Oh, J.; Lee, M. Pharmacokinetics of Drugs in Rats with Diabetes

Mellitus Induced by Alloxan or Streptozocin: Comparison with Those in Patients

with Type I Diabetes Mellitus. J. Pharm. Pharmacol., 2010, 62, 1–23.

[9] Szkudelski, T. The Mechanism of Alloxan and Streptozotocin Action in B Cells of

the Rat Pancreas. Physiol. Res., 2001, 50, 537–546.

29

[10] Sandler, S.; Swenne, I. Streptozotocin, but Not Alloxan, Induces DNA Repair

Synthesis in Mouse Pancreatic Islets in Vitro. Diabetologia, 1983, 25, 444–447.

[11] McLaughlin, P.J.; Cain, J.D.; Titunick, M.B.; Sassani, J.W.; Zagon, I.S. Topical

Naltrexone Is a Safe and Effective Alternative to Standard Treatment of Diabetic

Wounds. Adv. Wound Care, 2017, 6, 279–288.

[12] McLaughlin, P.J.; Immonen, J.A.; Zagon, I.S. Topical Naltrexone Accelerates Full-

Thickness Wound Closure in Type 1 Diabetic Rats by Stimulating Angiogenesis.

Exp. Biol. Med., 2013, 238, 733–743.

[13] Immonen, J.A.; Zagon, I.S.; Lewis, G.S.; McLaughlin, P.J. Topical Treatment with

the Opioid Antagonist Naltrexone Accelerates the Remodeling Phase of Full-

Thickness Wound Healing in Type 1 Diabetic Rats. Exp. Biol. Med., 2013, 238,

1127–1135.

[14] Nerup, J.; Mandrap-Poulsen, T.; Helqvist, S.; Andersen, H.U.; Pociot, F.; Reimers,

J.I.; Cuartero, B.G.; Karlsen, A.E.; Bjerre, U.; Lorenzen, T. On the Pathogenesis of

IDDM. Diabetologia, 1994, 37, S82–S89.

[15] Zagon, I.S.; Sassani, J.W.; Immonen, J.A.; McLaughlin, P.J. Ocular Surface

Abnormalities Related to Type 2 Diabetes Are Reversed by the Opioid Antagonist

Naltrexone: NTX Repairs Corneal Surface Epithelium. Clin. Experiment.

Ophthalmol., 2014, 42, 159–168.

[16] Baltzis, D.; Eleftheriadou, I.; Veves, A. Pathogenesis and Treatment of Impaired

Wound Healing in Diabetes Mellitus: New Insights. Adv. Ther., 2014, 31, 817–836.

30

[17] Standring, S. Gray’s Anatomy: The Anatomical Basis of Clinical Practice. In Skin

and its appendages; Elsevier Limited, 2016.

[18] Singer, A.J.; Clark, R.A.F. Cutaneous Wound Healing. N. Engl. J. Med., 1999, 738–

746.

[19] Ramasastry, S.S. Acute Wounds. Clin. Plast. Surg., 2005, 32, 195.

[20] National Diabetes Statistics Report, 2017; Centers for Disease Control and

Prevention: Atlanta, GA, 2017.

[21] Novak, M. Diabetes Mellitus. In Manual of Nursing Practice; Nettina, S., Ed.;

Lippincott Williams & Wilkins, Wolters Kluwer: London, 2010; pp. 944–969.

[22] Sharp, A.; Clark, J. Diabetes and Its Effects on Wound Healing. Nurs. Stand., 2010,

25, 41–47.

[23] Meeking, D.; Holland, E.; Land, D. Diabetes and Foot Disease. In Diabetes: Chronic

Complications; Shaw, K.; Cummings, M., Eds.; Wiley: Chichester, 2006; pp. 47–78.

[24] BROWN, C.D.; GHALI, H.S.; ZHAO, Z.; THOMAS, L.L.; FRIEDMAN, E.A. Association of

Reduced Red Blood Cell Deformability and Diabetic Nephropathy. Kidney Int.,

2005, 67, 295–300.

[25] Morain, W.; Colen, L. Wound Healing in Diabetes Mellitus. Clin. Plast. Surg., 1990,

17, 493–501.

[26] McIntosh, C. Diabetic Foot Ulcers: An Educational Booklet; Wounds; UK

Publishing: Aberdeen, 2006.

[27] Hunt, T.; Pai, M. The Effect of Variant Ambient Oxygen Tensions on Wound

Metabolism and Collagen Synthesis. Surg. Gynaecol. Obstet., 1972, 135, 561–567.

31

[28] Lioupis, C. Effects of Diabetes Mellitus on Wound Healing: An Update. J. Wound

Care, 2005, 14, 84–86.

[29] Kidman, K. Tissue Repair and Regeneration: The Effects of Diabetes on Wound

Healing. Diabet. Foot J., 2008, 11, 73–79.

[30] Loots, M. Fibroblasts Derived from Chronic Diabetic Ulcers Differ in Their

Response to Stimulation with EGF, IGF-I, BFGF and PDGF-AB Compared to

Controls. Eur. J. Cell Biol., 2002, 81, 153–160.

[31] McIntosh, C. Managing Diabetic Foot Ulceration: Review of Best Practice. Wound

Essent., 2009, 4, 102–111.

[32] Lipsky, B.; Berendt, A. Infection of the Foot in Persons with Diabetes:

Epidemiology, Pathophysiology, Microbiology, Clinical Presentation and Approach

to Therapy. In The Foot in Diabetes; Wiley & Sons: Chichester, 2006; pp. 159–168.

[33] Edmonds, M.; Foster, A. ABC of Wound Healing. Diabetic Foot Ulcers. Br. Med. J.,

2006, 332, 407–410.

[34] Uchi, H.; Igarashi, A.; Urabe, K.; Koga, T.; Nakayama, J.; Kawamori, R.; Tamaki, K.

Clinical Efficacy of Basic Fibroblast Growth Factor (BFGF) for Diabetic Ulcer.

Dermatology, 2009, 19, 461–468.

[35] Maier, H.; Ilich, J.; Kim, J.; Spicer, M. Nutrition Supplementation for Diabetic

Wound Healing: A Systematic Review of Current Literature. Skinmed, 2013, 11,

217–224.

[36] Tecilazich, F.; Dinh, T.; Veves, A. Emerging Drugs for the Treatment of Diabetic

Ulcers. Expert Opin. Emerg. Drugs, 2013, 18, 207–217.

32

[37] Isaac, A.; Armstrong, D. Negative Pressure Wound Therapy and Other New

Therapies for Diabetic Foot Ulceration: The Current State of Play. Med. Clin.

North Am., 2013, 97, 899–909.

[38] Argenta, L.; Morykwas, M. Vacuum-Assisted Closure: A New Method for Wound

Control and Treatment: Clinical Experience. Ann. Plast. Surg., 1997, 38, 563–576.

[39] Plikaitis, C.; Molnar, J. Subatmospheric Pressure Wound Therapy and the Vacuum-

Assisted Closure Device: Basic Science and Current Clinical Successes. Expert Rev.

Med. Devices, 2006, 3, 175–184.

[40] Seo, S.; Yeo, J.; Kim, J.; Kim, J.; Cho, T.; Lee, D. Negative-Pressure Wound Therapy

Induces Endothelial Progenitor Cell Mobilization in Diabetic Patients with Foot

Infection or Skin Defects. Exp. Mol. Med., 2013, 45, e62.

[41] Thakral, G.; Lafontaine, J.; Najafi, B.; Talal, T.; Kim, P.; Lavery, L. Electrical

Stimulation to Accelerate Wound Healing. Diabet. Foot Ankle, 2013, 4.

[42] Moretti, B.; Notarnicola, A.; Maggio, G.; Moretti, L.; Pascone, M.; Tafuri, S.;

Patella, V. The Management of Neuropathic Ulcers on the Foot in Diabetes by

Shock Wave Therapy. BMC Musculoskelet. Disord., 2009, 10.

[43] Game, F.L.; Hinchliffe, R.J.; Apelqvist, J.; Armstrong, D.G.; Bakker, K.; Hartemann,

A.; Löndahl, M.; Price, P.E.; Jeffcoate, W.J.; Diabetes and Endocrinology; Cellular

Autoimmunity; Diabetes och endokrinologi; Lund University; Lunds universitet;

Cellulär autoimmunitet. A Systematic Review of Interventions to Enhance the

Healing of Chronic Ulcers of the Foot in Diabetes. Diabetes Metab. Res. Rev.,

2012, 28, 119–141.

33

[44] Huang, P.; Li, S.; Han, M.; Xiao, Z.; Yang, R.; Han, Z.C. Autologous Transplantation

of Granulocyte Colony–Stimulating Factor–Mobilized Peripheral Blood

Mononuclear Cells Improves Critical Limb Ischemia in Diabetes. Diabetes Care,

2005, 28, 2155–2160.

[45] Dubsky, M.; Jirkovska, A.; Bem, R.; Fejfarova, V.; Pagacova, L.; Sixta, B.; Varga, M.;

Langkramer, S.; Sykova, E.; Jude, E.B. Both Autologous Bone Marrow

Mononuclear Cell and Peripheral Blood Progenitor Cell Therapies Similarly

Improve Ischaemia in Patients with Diabetic Foot in Comparison with Control

Treatment. Diabetes Metab. Res. Rev., 2013, 29, 369–376.

[46] Humpert, P.; Bartsch, U.; Konrade, I.; Hammes, H.; Morcos, M.; Kasper, M.;

Bierhaus, A.; Nawroth, P. Locally Applied Mononuclear Bone Marrow Cells

Restore Angiogenesis and Promote Wound Healing in a Type 2 Diabetic Patient.

Exp. Clin. Endocrinol. DIABETES, 2005, 113, 538–540.

[47] Falanga, V.; Iwamoto, S.; Chartier, M.; Yufit, T.; Butmarc, J.; Kouttab, N.; Shrayer,

D.; Carson, P. Autologous Bone Marrow-Derived Cultured Mesenchymal Stem

Cells Delivered in a Fibrin Spray Accelerate Healing in Murine and Human

Cutaneous Wounds. TISSUE Eng., 2007, 13, 1299–1312.

[48] Mulder, G.; Tallis, A.; Marshall, V.; Mozingo, D.; Phillips, L.; Pierce, G.; Chandler, L.;

Sosnowski, B. Treatment of Nonhealing Diabetic Foot Ulcers with a Platelet-

Derived Growth Factor Gene-Activated Matrix (GAM501): Results of a Phase 1/2

Trial. WOUND REPAIR Regen., 2009, 17, 772–779.

34

[49] Brem, H.; Kodra, A.; Golinko, M.; Entero, H.; Stojadinovic, O.; Wang, V.; Sheahan,

C.; Weinberg, A.; Woo, S.; Ehrlich, H.; Tomic-Canic, M. Mechanism of Sustained

Release of Vascular Endothelial Growth Factor in Accelerating Experimental

Diabetic Healing. J. Invest. Dermatol., 2009, 129, 2275–2287.

[50] Rodgers, K.; Verco, S.; Bolton, L.; diZerega, G. Accelerated Healing of Diabetic

Wounds by NorLeu(3)-Angiotensin (1-7). EXPERT Opin. Investig. DRUGS, 2011, 20,

1575–1581.

[51] Rodgers, K.; Espinoza, T.; Felix, J.; Roda, N.; Maldonado, S.; diZerega, G.

Acceleration of Healing, Reduction of Fibrotic Scar, and Normalization of Tissue

Architecture by an Angiotensin Analogue, NorLeu(3)-A(1-7). Plast. Reconstr. Surg.,

2003, 111, 1195–1206.

[52] Balingit, P.P.; Armstrong, D.G.; Reyzelman, A.M.; Bolton, L.; Verco, S.J.; Rodgers,

K.E.; Nigh, K.A.; diZerega, G.S. NorLeu3-A(1–7) Stimulation of Diabetic Foot Ulcer

Healing: Results of a Randomized, Parallel-group, Double-blind, Placebo-

controlled Phase 2 Clinical Trial. Wound Repair Regen., 2012, 20, 482–490.

[53] Kant, V.; Gopal, A.; Kumar, D.; Bag, S.; Kurade, N.; Kumar, A.; Tandan, S. Topically

Applied Substance P Enhanced Healing of Open Excision Wound in Rats. Eur. J.

Pharmacol., 2013, 715, 345–353.

[54] Mirza, R.; Fang, M.; Ennis, W.; Koh, T. Blocking Interleukin-1 Beta Induces a

Healing-Associated Wound Macrophage Phenotype and Improves Healing in Type

2 Diabetes. DIABETES, 2013, 62, 2579–2587.

35

[55] Einhorn, T.; Gerstenfeld, L. Fracture Healing: Mechanisms and Interventions. Nat.

Rev. Rheumatol., 2015, 11, 45–54.

[56] Schindeler, A.; McDonald, M.M.; Bokko, P.; Little, D.G. Bone Remodeling during

Fracture Repair: The Cellular Picture. Semin. Cell Dev. Biol., 2008, 19, 459–466.

[57] Tsiridis, E.; Upadhyay, N.; Giannoudis, P. Molecular Aspects of Fracture

Healing:Which Are the Important Molecules? Injury, 2007, 38, S11–S25.

[58] Kousteni, S.; Bilezikian, J.P. Cellular Actions of Parathyroid Hormone. In Principles

of Bone Biology; Elsevier Inc., 2008.

[59] Bahney, C.; Hu, D.; Miclau, T.; Marcucio, R. The Multifaceted Role of the

Vasculature in Endochondral Fracture Repair. Front. Endocrinol., 2015, 6, 4.

[60] Zhang, C. Transcriptional Regulation of Bone Formation by the Osteoblast-Specific

Transcription Factor Osx. J. Orthop. Surg., 2010, 5, 37–37.

[61] Noda, M.; Denhardt, D.T. Osteopontin. In Principles of Bone Biology; Elsevier Inc.,

2008.

[62] Eghbali-Fatourechi, G.Z.; Lamsam, J.; Fraser, D.; Nagel, D.; Riggs, B.L.; Khosla, S.

Circulating Osteoblast-Lineage Cells in Humans. N. Engl. J. Med., 2005, 352, 1959–

1966.

[63] Collett, G.D.M. Angiogenesis and Pericytes in the Initiation of Ectopic Calcification.

Circ. Res., 2005, 96, 930–938.

[64] Rumi, M.N.; Deol, G.S.; Singapuri, K.P.; Pellegrini Jr, V.D. The Origin of

Osteoprogenitor Cells Responsible for Heterotopic Ossification Following Hip

Surgery: An Animal Model in the Rabbit. J. Orthop. Res., 2005, 23, 34–40.

36

[65] Salerno, M.; Argenziano, A.; DiMaio, S.; Gasparini, N.; Formicola, S.; DeFilippo, G.;

Tenore, A. Pubertal Growth, Sexual Maturation, and Final Height in Children With

IDDM: Effects of Age at Onset and Metabolic Control. Diabetes Care, 1997, 20,

721–724.

[66] Vestergaard, P. Discrepancies in Bone Mineral Density and Fracture Risk in

Patients with Type 1 and Type 2 Diabetes—a Meta-Analysis. Osteoporos. Int.,

2007, 18, 427–444.

[67] Janghorbani, M.; Van Dam, R.M.; Willett, W.C.; Hu, F.B. Systematic Review of

Type 1 and Type 2 Diabetes Mellitus and Risk of Fracture. Am. J. Epidemiol., 2007,

166, 495–505.

[68] Cozen, L. Does Diabetes Delay Fracture Healing? Clin. Orthop., 1972, 82, 134–140.

[69] Retzepi, M.; Donos, N. The Effect of Diabetes Mellitus on Osseous Healing. Clin.

Oral Implants Res., 21, 673–681.

[70] Macey, L.R.; Kana, S.M.; Jingushi, S.; Terek, R.M.; Borretos, J.; Bolander, M.E.

Defects of Early Fracture-Healing in Experimental Diabetes. J Bone Jt. Surg Am,

1989, 71, 722–733.

[71] Rogers, L.C., DPM; Frykberg, R.G., DPM, MPH. The Charcot Foot. Med. Clin. North

Am., 2013, 97, 847–856.

[72] Jiao, H.; Xiao, E.; Graves, D.T. Diabetes and Its Effect on Bone and Fracture

Healing. Curr. Osteoporos. Rep., 2015, 13, 327–335.

37

[73] Yee, C.S.; Xie, L.; Hatsell, S.; Hum, N.; Murugesh, D.; Economides, A.N.; Loots,

G.G.; Collette, N.M. Sclerostin Antibody Treatment Improves Fracture Outcomes

in a Type I Diabetic Mouse Model. Bone, 2015, 82, 122–134.

[74] Botolin, S.; McCabe, L.R. Bone Loss and Increased Bone Adiposity in Spontaneous

and Pharmacologically Induced Diabetic Mice. Endocrinology, 2007, 148, 198–

205.

[75] Ogawa, N.; Yamaguchi, T.; Yano, S.; Yamauchi, M.; Yamamoto, M.; Sugimoto, T.

The Combination of High Glucose and Advanced Glycation End-Products (AGEs)

Inhibits the Mineralization of Osteoblastic MC3T3-E1 Cells through Glucose-

Induced Increase in the Receptor for AGEs. Horm. Metab. Res., 2007, 39, 871–

875.

[76] Cortizo, A.M.; Lettieri, M.G.; Barrio, D.A.; Mercer, N.; Etcheverry, S.B.; McCarthy,

A.D. Advanced Glycation End-Products (AGEs) Induce Concerted Changes in the

Osteoblastic Expression of Their Receptor RAGE and in the Activation of

Extracellular Signal-Regulated Kinases (ERK). Mol. Cell. Biochem., 2003, 250, 1–10.

[77] Santana, R.B.; Xu, L.; Chase, H.B.; Amar, S.; Graves, D.T.; Trackman, P.C. A Role for

Advanced Glycation End Products in Diminished Bone Healing in Type 1 Diabetes.

Diabetes, 2003, 52, 1502–1510.

[78] Bai, X.; Lu, D.; Bai, J.; Zheng, H.; Ke, Z.; Li, X.; Luo, S. Oxidative Stress Inhibits

Osteoblastic Differentiation of Bone Cells by ERK and NF-KappaB. Biochem.

Biophys. Res. Commun., 2004, 314, 197.

38

[79] Lu, H.; Kraut, D.; Gerstenfeld, L.C.; Graves, D.T. Diabetes Interferes with the Bone

Formation by Affecting the Expression of Transcription Factors That Regulate

Osteoblast Differentiation. Endocrinology, 2003, 144, 346–352.

[80] Almeida, M.; Han, L.; Martin-Millan, M.; O’Brien, C.; Manolagas, S. Oxidative

Stress Antagonizes Wnt Signaling in Osteoblast Precursors by Diverting Beta-

Catenin from T Cell Factor- to Forkhead Box O-Mediated Transcription. J. Biol.

Chem., 2007, 282, 27298–27305.

[81] Graves, D.T.; Alblowi, J.; Paglia, D.N.; O’Connor, J.P.; Lin, S. Impact of Diabetes on

Fracture Healing. J. Exp. Clin. Med., 2011, 3, 3–8.

[82] Graves, D.; Kayal, R. Diabetic Complications and Dysregulated Innate Immunity.

Front. Biosci.-LANDMARK, 2008, 13, 1227–1239.

[83] Gooch, H.; Hale, J.; Fujioka, H.; Balian, G.; Hurwitz, S. Alterations of Cartilage and

Collagen Expression during Fracture Healing in Experimental Diabetes. Connect.

TISSUE Res., 2000, 41, 81–81.

[84] Herbsman, H.; Powers, J.C.; Hirschman, A.; Shaftan, G.W. Retardation of Fracture

Healing in Experimental Diabetes. J. Surg. Res., 1968, 8, 424–431.

[85] HOU, J.; ZERNICKE, R.; BARNARD, R. EXPERIMENTAL DIABETES, INSULIN-

TREATMENT, AND FEMORAL-NECK MORPHOLOGY AND BIOMECHANICS IN RATS.

Clin. Orthop., 1991, 278–285.

[86] Reddy, G.K.; Stehno-Bittel, L.; Hamade, S.; Enwemeka, C.S. The Biomechanical

Integrity of Bone in Experimental Diabetes. Diabetes Res. Clin. Pract., 2001, 54, 1–

8.

39

[87] Weiss, R.E.; Reddi, A.H. Influence of Experimental Diabetes and Insulin on Matrix-

Induced Cartilage and Bone Differentiation. Am. J. Physiol. - Endocrinol. Metab.,

1980, 238, 200–207.

[88] Lu, C.; Miclau, T.; Hu, D.; Marcucio, R.S. Ischemia Leads to Delayed Union during

Fracture Healing: A Mouse Model. J. Orthop. Res., 2007, 25, 51–61.

[89] Lu, C.; Saless, N.; Wang, X.; Sinha, A.; Decker, S.; Kazakia, G.; Hou, H.; Williams, B.;

Swartz, H.M.; Hunt, T.K.; Miclau, T.; Marcucio, R.S. The Role of Oxygen during

Fracture Healing. Bone, 2012, 52, 220–229.

[90] Wu, D.; Malda, J.; Crawford, R.; Xiao, Y. Effects of Hyperbaric Oxygen on

Proliferation and Differentiation of Osteoblasts from Human Alveolar Bone.

Connect. TISSUE Res., 2007, 48, 206–213.

[91] Campos Pastor, M.M.; López-Ibarra, P.J.; Escobar-Jiménez, F.; Serrano Pardo,

M.D.; García-Cervigón, A. Intensive Insulin Therapy and Bone Mineral Density in

Type 1 Diabetes Mellitus: A Prospective Study. Osteoporos. Int., 2000, 11, 455–

459.

[92] Gandhi, A.; Beam, H.A.; O’Connor, J.P.; Parsons, J.R.; Lin, S.S. The Effects of Local

Insulin Delivery on Diabetic Fracture Healing. Bone, 2005, 37, 482–490.

[93] Ippolito, J.A.; Krell, E.S.; Cottrell, J.; Meyer, R.; Clark, D.; Nguyen, D.; Sudah, S.;

Muñoz, M.; Lim, E.; Lin, A.; Lee, T.J.H.; O’Connor, J.P.; Benevenia, J.; Lin, S.S.

Effects of Local Vanadium Delivery on Diabetic Fracture Healing. J. Orthop. Res.,

2017, 35, 2174–2180.

40

[94] Hreha, J.; Wey, A.; Cunningham, C.; Krell, E.S.; Brietbart, E.A.; Paglia, D.N.;

Montemurro, N.J.; Nguyen, D.A.; Lee, Y.; Komlos, D.; Lim, E.; Benevenia, J.;

O’Connor, J.P.; Lin, S.S. Local Manganese Chloride Treatment Accelerates Fracture

Healing in a Rat Model. J. Orthop. Res., 2015, 33, 122–130.

[95] KAWAGUCHI, H.; KUROKAWA, T.; HANADA, K.; HIYAMA, Y.; TAMURA, M.; OGATA,

E.; MATSUMOTO, T. STIMULATION OF FRACTURE REPAIR BY RECOMBINANT

HUMAN BASIC FIBROBLAST GROWTH-FACTOR IN NORMAL AND

STREPTOZOTOCIN-DIABETIC RATS. ENDOCRINOLOGY, 1994, 135, 774–781.

[96] Coords, M.; Breitbart, E.; Paglia, D.; Kappy, N.; Gandhi, A.; Cottrell, J.; Cedeno, N.;

Pounder, N.; O’Connor, J.P.; Lin, S.S. The Effects of Low-intensity Pulsed

Ultrasound upon Diabetic Fracture Healing. J. Orthop. Res., 2011, 29, 181–188.

[97] Gandhi, A.; Doumas, C.; O’Connor, J.P.; Parsons, J.R.; Lin, S.S. The Effects of Local

Platelet Rich Plasma Delivery on Diabetic Fracture Healing. Bone, 2006, 38, 540–

546.

[98] Azad, V.; Breitbart, E.; Al-Zube, L.; Yeh, S.; OʼConnor, J.P.; Lin, S.S. RhBMP-2

Enhances the Bone Healing Response in a Diabetic Rat Segmental Defect Model. J.

Orthop. Trauma, 2009, 23, 267–276.

[99] Tyndall, W.; Beam, H.; Zarro, C.; O’Connor, J.; Lin, S. Decreased Platelet Derived

Growth Factor Expression during Fracture Healing in Diabetic Animals. Clin.

Orthop., 2003, 319–330.

41

[100] Sassani, J.W.; Mc Laughlin, P.J.; Zagon, I.S. The Yin and Yang of the Opioid Growth

Regulatory System: Focus on Diabetes—The Lorenz E. Zimmerman Tribute

Lecture. J. Diabetes Res., 2016, 2016, 1–23.

[101] Akil, H.; Watson, S.J.; Young, E.; Lewis, M.E.; Khachaturian, H.; Walker, J.M.

Endogenous Opioids: Biology and Function. Annu. Rev. Neurosci., 1984, 7, 223–

255.

[102] Zagon, I.S.; McLaughlin, P.J.; Goodman, S.R.; Rhodes, R.E. Opioid Receptors and

Endogenous Opioids in Diverse Human and Animal Cancers. J. Natl. Cancer Inst.,

1987, 79, 1059.

[103] Zagon, I.S.; McLaughlin, P.J. An Opioid Growth Factor Regulates the Replication of

Microorganisms. Life Sci., 1992, 50, 1179–1187.

[104] Vermes, I.; Steinmetz, E.; Schoorl, J.; van der Veen, E.A.; Tilders, F.J. Increased

Plasma Levels of Immunoreactive Beta-Endorphin and Corticotropin in Non-

Insulin-Dependent Diabetes. Lancet Lond. Engl., 1985, 2, 725.

[105] Negri, M.; Tonnarini, G.; D’Alessandro, M.; Fallucca, F. Plasma Met-Enkephalin in

Type I Diabetes. Metabolism., 1992, 41, 460–461.

[106] Rosen, H.; Krichevsky, A.; Bar-Shavit, Z. The Enkephalinergic Osteoblast. J. Bone

Miner. Res., 1998, 13, 1515–1520.

[107] Rosen, H.; Polakiewicz, R.D.; Benzakine, S.; Bar-Shavit, Z. Proenkephalin A in Bone-

Derived Cells. Proc. Natl. Acad. Sci., 1991, 88, 3705–3709.

[108] Elhassan, A.M.; Lindgren, J.U.; Hultenby, K.; Bergstrom, J.; Adem, A. Methionine-

Enkephalin in Bone and Joint Tissues. J. Bone Miner. Res., 1998, 13, 88–95.

42

[109] Rosen, H.; Krichevsky, A.; Polakiewicz, R.D.; Benzakine, S.; Bar-Shavit, Z.

Developmental Regulation of Proenkephalin Gene Expression in Osteoblasts. Mol.

Endocrinol., 1995, 9, 1621–1631.

[110] Indig, F.E.; Benayahu, D.; Fried, A.; Wientroub, S.; Blumberg, S. Neutral

Endopeptidase (EC 3.4.24.11) Is Highly Expressed on Osteoblastic Cells and Other

Marrow Stromal Cell Types. Biochem. Biophys. Res. Commun., 1990, 172, 620–

626.

[111] Zagon, I.S.; Verderame, M.F.; McLaughlin, P.J. The Biology of the Opioid Growth

Factor Receptor (OGFr). Brain Res. Rev., 2002, 38, 351–376.

[112] Cheng, F.; McLaughlin, P.J.; Banks, W.A.; Zagon, I.S. Passive Diffusion of

Naltrexone into Human and Animal Cells and Upregulation of Cell Proliferation.

Am. J. Physiol.-Regul. Integr. Comp. Physiol., 2009, 297, R844–R852.

[113] Blumberg, H.; Dayton, H.B.; George, M.; Rapaport, D.N. N-Allylnoroxymorphone:

A Potent Narcotic Antagonist; 1961.

[114] Pert, C.B.; Pasternak, G.; Snyder, S.H. Opiate Agonists and Antagonists

Discriminated by Receptor Binding in Brain. Science, 1973, 182, 1359–1361.

[115] Zagon, I.S.; Donahue, R.N.; McLaughlin, P.J. Opioid Growth Factor-Opioid Growth

Factor Receptor Axis Is a Physiological Determinant of Cell Proliferation in Diverse

Human Cancers. Am. J. Physiol. - Regul. Integr. Comp. Physiol., 2009, 297, 1154–

1161.

43

[116] Ludwig, M.; Zagon, I.; McLaughlin, P. Serum [Met(5)]-Enkephalin Levels Are

Reduced in Multiple Sclerosis and Restored by Low-Dose Naltrexone. Exp. Biol.

Med., 2017, 242, 1524–1533.

[117] McLaughlin, P.J.; Zagon, I.S. Duration of Opioid Receptor Blockade Determines

Biotherapeutic Response. Biochem. Pharmacol., 2015, 97, 236–246.

[118] Zagon, I.S.; Sassani, J.W.; McLaughlin, P.J. Opioid Growth Factor Modulates

Corneal Epithelial Outgrowth in Tissue Culture. Am. J. Physiol. - Regul. Integr.

Comp. Physiol., 1995, 268, 942–950.

[119] Zagon, I.S.; Sassani, J.W.; Kane, E.R.; McLaughlin, P.J. Homeostasis of Ocular

Surface Epithelium in the Rat Is Regulated by Opioid Growth Factor. Brain Res.,

1997, 759, 92–102.

[120] Zagon, I.S.; Sassani, J.W.; McLaughlin, P.J. Re-Epithelialization of the Rat Cornea Is

Accelerated by Blockade of Opioid Receptors. Brain Res., 1998, 798, 254–260.

[121] Zagon, I.; Sassani, J.; McLaughlin, P. Cellular Dynamics of Corneal Wound Re-

Epithelialization in the Rat - II. DNA Synthesis of the Ocular Surface Epithelium

Following Wounding. BRAIN Res., 1999, 839, 243–252.

[122] Zagon, I.S.; Sassani, J.W.; McLaughlin, P.J. Re-Epithelialization of the Rabbit

Cornea Is Regulated by Opioid Growth Factor. Brain Res., 1998, 803, 61–68.

[123] Zagon, I.S.; Sassani, J.W.; Verderame, M.F.; McLaughlin, P.J. Particle-Mediated

Gene Transfer of Opioid Growth Factor Receptor CDNA Regulates Cell

Proliferation of the Corneal Epithelium. Cornea, 2005, 24, 614–619.

44

[124] Zagon, I.S.; Sassani, J.W.; Malefyt, K.J.; McLaughlin, P.J. Regulation of Corneal

Repair by Particle-Mediated Gene Transfer of Opioid Growth Factor Receptor

Complementary DNA. Arch. Ophthalmol., 2006, 124, 1620–1624.

[125] Zagon, I.S.; Sassani, J.W.; McLaughlin, P.J. Reepithelialization of the Human

Cornea Is Regulated by Endogenous Opioids. Invest. Ophthalmol. Vis. Sci., 2000,

41, 73–81.

[126] Abdelkader, H.; Patel, D.V.; McGhee, C.N.; Alany, R.G. New Therapeutic

Approaches in the Treatment of Diabetic Keratopathy: A Review. Clin. Experiment.

Ophthalmol., 2011, 39, 259–270.

[127] Rosenberg, M.E.; Tervo, T.M.T.; Immonen, I.J.; Muller, L.J.; Gronhagen-Riska, C.;

Vesaluoma, M.H. Corneal Structure and Sensitivity in Type 1 Diabetes Mellitus.

Invest. Ophthalmol. Vis. Sci., 2000, 41, 2915–2921.

[128] Ozdemir, M.; Ozdemir, G.; Buyukbese, M.A.; Cetinkaya, A. Risk Factors for Ocular

Surface Disorders in Patients with Diabetes Mellitus. Diabetes Res. Clin. Pract.,

2003, 59, 195–199.

[129] Cousen, P.; Cackett, P.; Bennett, H.; Swa, K.; Dhillon, B. Tear Production and

Corneal Sensitivity in Diabetes. J. Diabetes Complications, 2007, 21, 371–373.

[130] Dogru, M.; Katakami, C.; Inoue, M. Tear Function and Ocular Surface Changes in

Noninsulin-Dependent Diabetes Mellitus. Ophthalmology, 2001, 108, 586–592.

[131] Neira-Zalentein, W.; Holopainen, J.; Tervo, T.; Borras, F.; Acosta, M.; Belmonte, C.;

Gallar, J. Corneal Sensitivity in Diabetic Patients Subjected to Retinal Laser

Photocoagulation. Invest. Ophthalmol. Vis. Sci., 2011, 52, 6043–6049.

45

[132] Kaiserman, I.; Kaiserman, N.; Nakar, S.; Vinker, S. Dry Eye in Diabetic Patients. Am.

J. Ophthalmol., 2005, 139, 498–503.

[133] Grus, F.H.; Sabuncuo, P.; Dick, H.B.; Augustin, A.J.; Pfeiffer, N. Changes in the Tear

Proteins of Diabetic Patients. BMC Ophthalmol., 2002, 2, 4–4.

[134] Zagon, I.S.; Jenkins, J.B.; Sassani, J.W.; Wylie, J.D.; Ruth, T.B.; Fry, J.L.; Lang, C.M.;

McLaughlin, P.J. Naltrexone, an Opioid Antagonist, Facilitates Reepithelialization

of the Cornea in Diabetic Rat. Diabetes, 2002, 51, 3055–3062.

[135] Zagon, I.S.; Sassani, J.W.; McLaughlin, P.J. Insulin Treatment Ameliorates Impaired

Corneal Reepithelialization in Diabetic Rats. Diabetes, 2006, 55, 1141–1147.

[136] Zagon, I.S.; Klocek, M.S.; Sassani, J.W.; McLaughlin, P.J. Use of Topical Insulin to

Normalize Corneal Epithelial Healing in Diabetes Mellitus. Arch. Ophthalmol.,

2007, 125, 1082–1088.

[137] Zagon, I.S.; Sassani, J.W.; Myers, R.L.; McLaughlin, P.J. Naltrexone Accelerates

Healing without Compromise of Adhesion Complexes in Normal and Diabetic

Corneal Epithelium. Brain Res. Bull., 2007, 72, 18–24.

[138] Klocek, M.S.; Sassani, J.W.; McLaughlin, P.J.; Zagon, I.S. Naltrexone and Insulin Are

Independently Effective but Not Additive in Accelerating Corneal Epithelial

Healing in Type I Diabetic Rats. Exp. Eye Res., 2009, 89, 686–692.

[139] Klocek, M.S.; Sassani, J.W.; McLaughlin, P.J.; Zagon, I.S. Topically Applied

Naltrexone Restores Corneal Reepithelialization in Diabetic Rats. J. Ocul.

Pharmacol. Ther., 2007, 23, 89–102.

46

[140] Tavakoli, M.; Quattrini, C.; Abbott, C.; Kallinikos, P.; Marshall, A.; Finnigan, J.;

Morgan, P.; Efron, N.; Boulton, A.; Malik, R. Corneal Confocal Microscopy A Novel

Noninvasive Test to Diagnose and Stratify the Severity of Human Diabetic

Neuropathy. DIABETES CARE, 2010, 33, 1792–1797.

[141] Tavakoli, M.; Kallinikos, P.; Iqbal, A.; Herbert, A.; Fadavi, H.; Efron, N.; Boulton,

A.J.M.; A Malik, R. Corneal Confocal Microscopy Detects Improvement in Corneal

Nerve Morphology with an Improvement in Risk Factors for Diabetic Neuropathy.

Diabet. Med., 2011, 28, 1261–1267.

[142] Tavakoli, M.; Boulton, A.J.M.; Efron, N.; Malik, R.A. Increased Langerhan Cell

Density and Corneal Nerve Damage in Diabetic Patients: Role of Immune

Mechanisms in Human Diabetic Neuropathy. Contact Lens Anterior Eye, 2010, 34,

7–11.

[143] Tavakoli, M.; Petropoulos, I.N.; Malik, R.A. Assessing Corneal Nerve Structure and

Function in Diabetic Neuropathy. Clin. Exp. Optom., 2012, 95, 338–347.

[144] Tavakoli, M.; Petropoulos, I.N.; Malik, R.A. Corneal Confocal Microscopy to Assess

Diabetic Neuropathy: An Eye on the Foot. J. Diabetes Sci. Technol., 2013, 7, 1179–

1189.

[145] Papanas, N.; Ziegler, D. Corneal Confocal Microscopy: A New Technique for Early

Detection of Diabetic Neuropathy. Curr. Diab. Rep., 2013, 13, 488–499.

[146] Alves, M.; Carvalheira, J.; Modulo, C.; Rocha, E. Tear Film and Ocular Surface

Changes in Diabetes Mellitus. Arq. Bras. Oftalmol., 2008, 71, 96–103.

47

[147] Zagon, I.S.; Klocek, M.S.; Sassani, J.W.; McLaughlin, P.J. Dry Eye Reversal and

Corneal Sensation Restoration With Topical Naltrexone in Diabetes Mellitus. Arch.

Ophthalmol., 2009, 127, 1468–1473.

[148] McLaughlin, P.J.; Pothering, C.A.; Immonen, J.A.; Zagon, I.S. Topical Treatment

with the Opioid Antagonist Naltrexone Facilitates Closure of Full-Thickness

Wounds in Diabetic Rats. Exp. Biol. Med., 2011, 236, 1122–1132.

[149] Immonen, J.; Zagon, I.; McLaughlin, P. Topical Naltrexone as Treatment for Type 2

Diabetic Cutaneous Wounds. Adv. Wound Care, 2014, 3, 419–427.

[150] Naloxone Hydrochloride https://www.drugs.com/monograph/naloxone-

hydrochloride.html (accessed May 9, 2018).

[151] Blumberg, H.; Dayton, H.B. Naloxone and Related Compounds. In Antagonist and

Antagonists Actions of Narcotic Analgesic Drugs; Kosterlitz, H.W.; Collier, H.O.J.;

Villareal, V.E., Eds.; University Park Press: Baltimore, 1973; p. 110.

[152] Blumberg, H.; Dayton, H.B.; Wolf, P.S. Analgesic and Narcotic Antagonist

Properties of Noroxymorphone Derivatives. Toxicol. Appl. Pharmacol., 1967, 10.

[153] Jiang, J.B.; Hanson, R.N.; Portoghese, P.S.; Takemori, A.E. Stereochemical Studies

on Medicinal Agents. 23. Synthesis and Biological Evaluation of 6-Amino

Derivatives of Naloxone and Naltrexone. J. Med. Chem., 1977, 20, 1100–1102.

[154] Zagon, I.S.; McLaughlin, P.J. Naloxone Modulates Body and Organ Growth of Rats:

Dependency on the Duration of Opioid Receptor Blockade and Stereospecificity.

Pharmacol. Biochem. Behav., 1989, 33, 325–328.

48

[155] Liskov, A.V.; Solnyshkova, T.G.; Frolov, B.A.; Pavlovichev, S.A. Effect of Naloxone

Hydrochloride on Osteogenesis in Chick Embryos. Bull. Exp. Biol. Med., 2005, 139,

331–333.

[156] Petrizzi, L.; Mariscoli, M.; Valbonetti, L.; Varasano, V.; Langhoff, J.; Von

Rechenberg, B. Preliminary Study on the Effect of Parenteral Naloxone, Alone and

in Association with Calcium Gluconate, on Bone Healing in an Ovine “Drill Hole”

Model System. BMC Musculoskelet. Disord., 2007, 8, 43–43.

[157] Thakur, N.; DeBoyace, S.; Margulies, B. Antagonism of the Met5-Enkephalin-

Opioid Growth Factor Receptor-Signaling Axis Promotes MSC to Differentiate into

Osteoblasts: Antagonism of OGFR by Naloxone Increases Bone Formation. J.

Orthop. Res., 2015, n/a-n/a.

[158] KOLTA, M.; PIERZCHALA, K.; HOUDI, A.; VANLOON, G. EFFECT OF DIABETES ON

THE LEVELS OF 2 FORMS OF MET-ENKEPHALIN IN PLASMA AND PERIPHERAL-

TISSUES OF THE RAT. NEUROPEPTIDES, 1992, 21, 55–63.

[159] Motyl, K.; McCabe, L.R. Streptozotocin, Type I Diabetes Severity and Bone. Biol.

Proced. Online, 2009, 11, 296–315.

[160] McNair, P.; Madsbad, S.; Christiansen, C.; Faber, O.K.; Transbøl, I.; Binder, C.

Osteopenia in Insulin Treated Diabetes Mellitus: Its Relation to Age at Onset, Sex

and Duration of Disease. Diabetologia, 1978, 15, 87–90.

[161] Levin, M.E.; Boisseau, V.C.; Avioli, L.V. Effects of Diabetes Mellitus on Bone Mass

in Juvenile and Adult-Onset Diabetes. N. Engl. J. Med., 1976, 294, 241–245.

49

[162] Guo, J.; Dong, W.; Jin, L.; Wang, P.; Hou, Z.; Zhang, Y. Hydrogen-Rich Saline

Prevents Bone Loss in Diabetic Rats Induced by Streptozotocin. Int. Orthop., 2017,

41, 2119–2128.

[163] Erdal, N.; Gürgül, S.; Demirel, C.; Yildiz, A. The Effect of Insulin Therapy on

Biomechanical Deterioration of Bone in Streptozotocin (STZ)-Induced Type 1

Diabetes Mellitus in Rats. Diabetes Res. Clin. Pract., 2012, 97, 461–467.

[164] Lloyd, S.A.; Lang, C.H.; Zhang, Y.; Paul, E.M.; Laufenberg, L.J.; Lewis, G.S.;

Donahue, H.J. Interdependence of Muscle Atrophy and Bone Loss Induced by

Mechanical Unloading. J. Bone Miner. Res., 2014, 29, 1118–1130.

[165] Bouxsein, M.; Boyd, S.; Christiansen, B.; Guldberg, R.; Jepsen, K.; Muller, R.

Guidelines for Assessment of Bone Microstructure in Rodents Using Micro-

Computed Tomography. J. BONE Miner. Res., 2010, 25, 1468–1486.

[166] Schmitz, N.; Laverty, S.; Kraus, V.B.; Aigner, T. Basic Methods in Histopathology of

Joint Tissues. Osteoarthritis Cartilage, 2010, 18, S113–S116.

[167] Schneider, C.A.; Rasband, W.S.; Eliceiri, K.W. NIH Image to ImageJ: 25 Years of

Image Analysis. Nat. Methods, 2012, 9, 671–675.

[168] Kayath, M.J.; Tavares, E.F.; Dib, S.A.; Vieira, J.G.H. Prospective Bone Mineral

Density Evaluation in Patients With Insulin-Dependent Diabetes Mellitus. J.

Diabetes Complications, 1998, 12, 133–139.

[169] Kemink, S.A.G.; Hermus, A.R.M.M.; Swinkels, L.M.J.W.; Lutterman, J.A.; Smals,

A.G.H. Osteopenia in Insulin-Dependent Diabetes Mellitus; Prevalence and

Aspects of Pathophysiology. J. Endocrinol. Invest., 2000, 23, 295–303.

50

[170] MunozTorres, M.; Jodar, E.; EscobarJimenez, F.; LopezIbarra, P.; Luna, J. Bone

Mineral Density Measured by Dual X-Ray Absorptiometry in Spanish Patients with

Insulin-Dependent Diabetes Mellitus. Calcif. TISSUE Int., 1996, 58, 316–319.

[171] Räkel, A.; Sheehy, O.; Rahme, E.; LeLorier, J. Osteoporosis among Patients with

Type 1 and Type 2 Diabetes. Diabetes Metab., 2008, 34, 193–205.

[172] Martin, L.M.; McCabe, L.R. Type I Diabetic Bone Phenotype Is Location but Not

Gender Dependent. Histochem. Cell Biol., 2007, 128, 125–133.

[173] Hamada, Y.; Kitazawa, S.; Kitazawa, R.; Fujii, H.; Kasuga, M.; Fukagawa, M.

Histomorphometric Analysis of Diabetic Osteopenia in Streptozotocin-Induced

Diabetic Mice: A Possible Role of Oxidative Stress. Bone, 2006, 40, 1408–1414.

[174] The Cost of Diabetes http://www.diabetes.org/advocacy/news-events/cost-of-

diabetes.html (accessed Apr 29, 2018).

[175] Zhang, P.; Lu, J.; Jing, Y.; Tang, S.; Zhu, D.; Bi, Y. Global Epidemiology of Diabetic

Foot Ulceration: A Systematic Review and Meta-Analysis. Ann. Med., 2017, 49,

106–116.

[176] Immonen, J.; Zagon, I.; McLaughlin, P. Selective Blockade of the OGF-OGFr

Pathway by Naltrexone Accelerates Fibroblast Proliferation and Wound Healing.

Exp. Biol. Med., 2014, 239, 1300–1309.

[177] Zagon, I.; Klocek, M.; Sassani, J.; Mauger, D.; McLaughlin, P. Corneal Safety of

Topically Applied Naltrexone. J. Ocul. Pharmacol. Ther., 2006, 22, 377–387.

51

[178] Liang, D.; Sassani, J.W.; McLaughlin, P.J.; Zagon, I.S. Topical Application of

Naltrexone to the Ocular Surface of Healthy Volunteers: A Tolerability Study. J.

Ocul. Pharmacol. Ther., 2016, 32, 127–132.

[179] Ki67 - A Crucial Cellular Proliferation Marker (accessed Jun 13, 2018).

[180] Zagon, I.S.; Wu, Y.; McLaughlin, P.J. Opioid Growth Factor and Organ

Development in Rat and Human Embryos. Brain Res., 1999, 839, 313–322.

[181] Wang, G.; Wang, J.; Fu, Y.; Bai, L.; He, M.; Li, B.; Fu, Q. Systemic Treatment with

Vanadium Absorbed by Coprinus Comatus Promotes Femoral Fracture Healing in

Streptozotocin-Diabetic Rats. Biol. Trace Elem. Res., 2013, 151, 424–433.

[182] Wang, L.X.; Jiang, H.L.; Du, S.L. Observed Impacts of Insulin Therapy on Callus Cell

Transforming Growth Factor-Beta 1 Expression in Diabetic Rats. Genet. Mol. Res.

GMR, 2015, 14, 5076–5084.

[183] Ko, K.I.; Coimbra, L.S.; Tian, C.; Alblowi, J.; Kayal, R.A.; Einhorn, T.A.; Gerstenfeld,

L.C.; Pignolo, R.J.; Graves, D.T. Diabetes Reduces Mesenchymal Stem Cells in

Fracture Healing through a TNFα-Mediated Mechanism. Diabetologia, 2015, 58,

633–642.

[184] Park, A.G.; Paglia, D.N.; Al-Zube, L.; Hreha, J.; Vaidya, S.; Breitbart, E.; Benevenia,

J.; O’Connor, J.P.; Lin, S.S. Local Insulin Therapy Affects Fracture Healing in a Rat

Model. J. Orthop. Res., 2013, 31, 776–782.

[185] Hough, S.; Avioli, L.V.; Bergfeld, M.A.; Fallon, M.D.; Slatopolsky, E.; Teitelbaum,

S.L. Correction of Abnormal Bone and Mineral Metabolism in Chronic

52

Streptozotocin-Induced Diabetes Mellitus in the Rat by Insulin Therapy.

Endocrinology, 1981, 108, 2228–2234.

53

CHAPTER 2: TOPICAL NALTREXONE IS A SASFE AND EFFECTIVE ALTERNATIVE TO

STANDARD TREATMENT OF DIABETIC WOUNDS1

Patricia J. McLaughlin, Jarrett D. Cain, Michelle B. Titunick, Ian S. Zagon __________________________________ 1Chapter 2 consists of a previously published paper approved by the dissertation committee for the use of this dissertation. The paper has been reformatted to fit into this dissertation. The citation for the original paper is: McLaughlin, P., Cain, J., Titunick, M., Sassani, J., & Zagon, I. (2017). Topical naltrexone is a safe and effective alternative to standard treatment of diabetic wounds. Advances in Wound Care, 6(9), 279-288. doi:10.1089/wound.2016.0725

54

2.1. Abstract

2.1.1. Objective:

Diabetes affects more than 29 million individuals in the United States resulting in

$245 billion in healthcare costs. Approximately 15% of these individuals will develop a

chronic, non-healing foot ulcer (DFU), and if untreated, may lead to amputation of part

or the entire lower limb. The current treatments for DFU are expensive, have significant

side-effects, and result in non-compliant patients. A new topical treatment is described

that enhances wound repair and more importantly, focuses on underlying diabetic

pathways.

2.1.2. Approach:

The efficacy of topical naltrexone (NTX), an opioid receptor antagonist, and

Regranex® was compared in preclinical studies using type 1 diabetic (T1D) rats. Dorsal

cutaneous wounds were treated topically with 0.03% NTX, Regranex®, or moisturizing

cream alone. Wound closure, DNA synthesis, and cytokine production were monitored.

2.1.3. Results:

Wound closure rates with topical NTX in diabetic rats were comparable to

Regranex®. Topical NTX accelerated DNA synthesis as measured by BrdU incorporation,

enhanced platelet derived growth factor (PDGF) expression, and increased angiogenesis

as measured by vascular endothelial growth factor (VEGF) expression. Regranex® had

little effect on DNA synthesis and VEGF expression relative to vehicle-treated wounds,

and only temporarily increased PDGF expression. Neither treatment altered fibroblast

growth factor expression over the four day observation period.

55

2.1.4. Innovation and Conclusion:

The data suggest that blockade of the OGF-OGFr axis utilizing 0.03% NTX cream is

a safe, inexpensive and effective alternative for treatment for diabetic wounds. NTX also

increased cell replication and angiogenesis that are two important components of wound

healing. In head-to-head comparisons with Regranex, NTX was equally effective.

2.1. Introduction

More than 29 million individuals in the United States are diagnosed with diabetes,

with type 1 diabetes accounting for approximately 5-10% [1]. It is estimated that an

additional 37% of the population have signs of pre-diabetes

[https://www.sciencedaily.com/releases/2015/09/150908112428.htm]. The estimated

annual healthcare costs exceed $245 billion annually [1-3]. In addition to treatment of the

disease itself, diabetes is accompanied by complications that increase healthcare costs

and reduce the quality of life [3-5]. Several non-life threatening complications involve

delayed epithelialization and/or impaired sensitivity such as dry eye, corneal keratopathy

and delayed cutaneous wound closure, often manifested as diabetic foot ulcers (DFU).

Impaired cutaneous wound healing, even if treated, can result in chronic lesions, ulcers,

epithelial erosion, and amputation of the extremities [1-7]. Approximately 50% of the

170 million people worldwide with diabetes experience complications associated with

delayed cutaneous wound healing [1,2,4,5].

The processes underlying wound healing are dynamic and culminate in the

restoration of proper anatomical function of the tissue [8,9]. Wound closure occurs as a

continuum of overlapping phases that involve re-epithelialization, inflammation,

56

proliferation, and formation of granulation tissue during remodeling. Each phase is

initiated by cytokines and growth factors [7], and the secretion and expression of these

cytokines appears to be compromised in hyperglycemic individuals and animals leading

to delayed cell replication [9-15]. Many of the current therapies used for treatment of

DFU provide topical application of a cytokine or growth factor [10-12], but are not disease

modifying treatments that address underlying defects related to diabetes. Regranex® is

an FDA-approved standard of care for DFUs [16]. Preclinical and clinical studies have

demonstrated that the active ingredient of platelet-derived growth factor is effective in

wound restoration following topical application [17-19]. However, this therapy is

expensive and associated with side-effects and warnings that prevent extended usage.

There remains a need for new alternative treatments for chronic, non-healing wounds.

The novel therapy discussed in this study is safe, inexpensive, effective, and

targets one of the underlying dysregulated pathways in diabetes. The therapy utilizes

topical application of the opioid antagonist naltrexone (NTX) to block the opioid growth

factor (OGF) – OGF receptor (OGFr) regulatory pathway. Preclinical studies have shown

that this small molecular weight compound (377 MW) can be dissolved in a carrier and

topically applied to full-thickness wounds to accelerate full-thickness cutaneous wounds

in type 1 diabetic rats, as well as type 2 genetically diabetic mice (db/db), for accelerated

wound repair [13-15]. As an opioid receptor antagonist, NTX lacks intrinsic biological

activity and works by blocking interactions between an inhibitory peptide and its receptor

[20,21]. The mechanism of action involves extended blockade of the OGF-OGFr

regulatory pathway [22] that has been reported to be dysregulated in diabetes leading to

57

overexpression of the inhibitory peptide, OGF, chemically termed [Met5 enkephalin], is

an endogenous neuropeptide that inhibits cell replication. OGF levels are elevated in

human and animal models of diabetes leading to downregulation of cell proliferation and

renewal processes in wound healing [27-29]. Total opioid receptor blockade by NTX

restores the proliferating homeostasis required for tissue repair [13-15, 22-26].

In this preclinical study, a comparison of effectiveness was made between the new

NTX formulation (0.03%) and Regranex® applied once daily for treatment of cutaneous

wounds in type 1 diabetic rats. In addition to cell replication and cytokine production of

VEGF, PDGF, and FGF were evaluated in order to determine mechanism of action for both

therapies.

2.2. Clinical Problem Addressed

A major complication of diabetes is delayed closure of full thickness cutaneous

wounds. The standard of care therapy, Regranex, works by increasing PDGF, is

expensive and is associated with severe side-effects including cancer, and even death.

An alternative topical therapy that has been shown preclinically to be safe and effective

is NTX. NTX targets the underlying pathophysiology of diabetes and enhances cell

replication at all stages of wound remodeling. NTX is safe, inexpensive, and effectively

targets the disease-modifying pathways in this comparison study of wound closure in

Type 1 diabetic rats. These data support large clinical trials and FDA-approval for this

topical treatment.

58

2.3. Materials and Methods

2.4.1. Animals and Induction of Diabetes

Adult male Sprague-Dawley rats (Charles River Laboratories, Wilmington, MA)

were purchased at 6 weeks of age and housed in AAALAC approved facilities with water

and food (2018 Global Rodent Diet, Teklad®, Indianapolis, IN) provided ad libitum. All

protocols were approved by the IACUC at Penn State University College of Medicine, and

conformed to guidelines of the National Institutes of Health.

At six weeks of age, rats were rendered hyperglycemic by two consecutive

injections (i.p.) of 40 mg/kg streptozotocin (STZ, Sigma, St. Louis, MO) dissolved in citrate

buffer (pH 4.5). Normal rats received only i.p. injections of citrate buffer. This regimen

produced insulin-dependent type 1 diabetes (T1D) within 4-5 days. Although this model

of hyperglycemia and subsequent delayed wound healing does not mimic the long-term

chronic, pressure wounds associated with human diabetes, the model lends itself to study

of mechanism and therapeutic response [30].

Body weights were recorded periodically. Blood glucose measurements were

taken using a True Track Smart System glucometer (Home Diagnostics, Ft. Lauderdale, FL).

Three independent experiments were performed over a period of 9 months; each

experiment had approximately 15 rats rendered hyperglycemic. In each experiment, rats

remained hyperglycemic (blood glucose >350 mg/dL) for 6 weeks prior to study without

insulin supplementation. At the start of each experiment if the rat had blood glucose

measurements greater than 600 mg/dl or appeared lethargic and unwilling to eat, the

animal was not included in the experiment.

59

2.4.2. Cutaneous wound surgery

Full thickness cutaneous wounds were created as described previously [13,14]. In

brief, on the day prior to surgery, the dorsum of each rat was shaved using an electric

razor followed by application of Nair® to remove all hair. Rats were immobilized by i.p.

injection of a mixture of ketamine (60 mg/kg; Ketaject®), xylazine (10 mg/kg,

TranquiVed®, Vedco), and acepromazine (1 mg/kg; acepromazine maleate; Vedco). Four

excisional circular (6 mm diameter) skin wounds were created 1 cm off the midline under

sterile conditions on each rat. Wounds were created to the level of the panniculus muscle

using disposable biopsy punches (Accuderm). After surgery, wounds were swabbed with

an antiseptic surgical scrub and left without dressing. All surgeries were conducted

between 0800-1100 h to alleviate the potential effects of diurnal rhythm.

2.4.3. Wound Treatment and Closure

Topical treatment of wounds with NTX, Regranex®, or vehicle was randomized for

each rat in order to control for placement of wound and treatment response. NTX was

dissolved in sterile saline (v/v) for a final dosage of 0.03% NTX in Neutrogena moisturizing

cream, 0.1 ml sterile saline was dissolved in Neutrogena cream, and Regranex® was used

directly from the pharmaceutical-grade tube. Each formulation was applied once daily (1X)

at 0900 h. The fourth wound received one of the treatments in a random manner.

Residual wound sizes were visualized with 20x magnification on an Olympus BH2-

RFCA microscope, and photographed using Spot Advanced software; the digital camera

was placed on a stable tripod located approximately 15 cm above the animal.

Measurements were calibrated with a ruler photographed adjacent to the wounds.

60

Animals were sedated in a regulated vaporizer with a 3% isoflurane- oxygen mixture and

photographs taken immediately after surgery (day 0) and every other day for two weeks.

Areal analysis of each wound was performed using Image J software, and the percent area

of residual defect was calculated.

2.4.4 BrdU-labeling and DNA synthesis

To assess cell replication, rats were injected with bromodeoxyuridine (100 mg/kg,

0.2ml BrdU, Sigma-Aldrich) at 6 hr and 3 hr prior to euthanasia on day 4. Tissue sections

were stained with anti-BrdU antibody (1:50, Invitrogen) followed by monoclonal

secondary antibody (1:2000, Invitrogen) and counterstained with hematoxylin. BrdU

labeling indexes were determined as the percentage of positive BrdU-labeled basal

epithelial cells per total basal epithelial cells superficial to the wound site.

2.4.5 Histological analysis and immunohistochemistry

On days 1, 2, and 4 following initial surgery, skin was harvested from rats. A 3-cm2

region of skin encompassing the original wound was removed for at least 6

wounds/treatment group, bisected, fixed in 10% neutral buffered formalin and processed

for paraffin embedding. Skin sections (10 μm thick) were stained with hematoxylin-eosin

for general assessment of toxicity and pathology. Angiogenesis monitored by expression

of VEGF was performed by immunohistochemical staining with polyclonal antibodies to

VEGF (1:200, sc-152, Santa Cruz) [31,32]. Other cytokine markers for wound repair were

evaluated by immunohistochemical staining with polyclonal antibodies to FGF-2 (1:500,

Ab106245, Abcam, Cambridge, MA) and PDGF (1:75, Ab21234, Abcam, Cambridge, MA);

all primary reactions were followed by staining with goat anti-rabbit (1:1000;

61

ThermoFisher) secondary antibodies. Antibody retrieval was completed by placing slides

in a pressure cooker for 3 min in sodium citrate buffer or EDTA buffer.

2.4. Statistical Analysis

Body weights and glucose measurements, as well as epithelial thicknesses in

unwounded animals, were evaluated using the Student’s two-tailed t-test. Residual

wound areas, epithelial thicknesses, and BrdU labeling indexes were analyzed using

analysis of variance (ANOVA) with subsequent planned comparisons made using

Newman-Keuls tests. A statistical power analysis for the number of wounds treated, as

well as the number of sections per wound required for morphological reliability was based

on previous experiments. Cytokine expression levels were semi-quantitatively assessed

by either measuring the area of staining or by counting the number of positively stained

cells within a grid.

2.5. Results

2.6.1. Body Weight and Blood Glucose Measurements

The mean weight of all male rats was 170 ± 2 g at the start of experimentation.

Approximately six weeks after STZ injection, the diabetic animals weighed 324 ± 7 g in

comparison to normal controls weighing 399 ± 18 g. Diabetic rats had significantly

elevated blood glucose levels within 2-3 days of STZ injections (~540 mg/dL), and by the

initiation of wound surgery, glucose levels in the DB rats were 563 ±12 mg/dL in

comparison to normal, non-diabetic with glucose levels of 124 ± 6 mg/dL.

62

2.6.2. Full-thickness Wound Closure

Overall analyses of wound closure indicated that wounds in diabetic rats healed

more slowly than those in normal animals. Comparison of Regranex® and NTX treatments

in T1D rats revealed that both treatments accelerated wound closure (Figure 2.1.A, B).

Evaluation of residual wound size over a 14 day period demonstrated that within 48 hr of

treatment, wound sizes in the DB/R and DB/NTX groups were smaller than those in the

DB/Vehicle group, with the mean DB/R group reaching statistical significance. At 4, 8 and

10 days, both treatment groups had wound areas that were significantly smaller than

those in the DB/Vehicle group. By day 12, the DB/NTX group was statistically smaller,

although the wound size was virtually negligible. At no time did the DB/NTX and DB/R

groups demonstrate differences in residual wound size or rates of closure.

63

Figure 2.1. Wound closure is accelerated by treatment of topical NTX or Regranex®. (A)

Photographs of full-thickness wounds following application of topical NTX, Regranex®, or Vehicle

to type 1 diabetic (DB) rats. Wounds (5 mm diameter) were created on the dorsum and treated

once daily (0.05 ml) with 10-5 M NTX (DB/NTX) or saline (DB/Vehicle) dissolved in Neutrogena

moisturizing cream, or Regranex® (DB/R). Photographs were taken immediately after surgery

(day 0) and on days 4, 8, and 12. Bar = 5 mm. (B) Residual wound areas (%) following surgical

wounding on the dorsum of DB mice receiving saline dissolved in Neutrogena moisturizing cream

(Normal + Vehicle), 10-5 M NTX dissolved in Neutrogena moisturizing cream (DB/NTX), or

Regranex® (DB/R). Significantly different between DB/Vehicle and NTX or Regranex® treatment

at *P<0.05, **P<0.01. Rate of healing or residual wound size did not differ between NTX-

treatment and Regranex®.

64

2.6.3. Histological Analysis of Skin

Tissue sections of wounds receiving all 3 treatments and stained with hematoxylin

and eosin revealed that neither Regranex® nor NTX distorted wound closure or appeared

to have abnormal pathology.

2.5.4. DNA Labeling Indexes

Given that most of the processes involved with wound closure require cell

replication, levels of DNA synthesis were evaluated to determine mechanism of action for

NTX and Regranex® (Figure 2.2.). Evaluation of wounds from normal rats treated with

vehicle revealed BrdU labeling indexes of 47.6 ± 3.5% in comparison to those with 29.3

±1.9% DNA incorporation in diabetic rats receiving vehicle. However, BrdU incorporation

in wounds of diabetic rats receiving NTX demonstrated 41.9 ± 2.3% labeling, a 43%

increase over DB/Vehicle rats (p<0.001), and comparable to that of normal animals.

Regranex® treatment had little effect on cell proliferation as the DNA synthesis rates were

21.0 ± 3.6, nearly 50% less than naltrexone treatment.

65

Figure 2.2. Cell proliferation in the basal epithelial layer in renewed skin. (A) Photomicrographs

of BrdU labeling in the basal layer of epithelium positioned over the granulation tissue of closed

wounds. Type 1 diabetic (DB) rats were treated with 10-5 M NTX (DB/NTX) or saline (DB/Vehicle)

dissolved in Neutrogena moisturizing cream, or Regranex® (DB/R) for 4 days. A separate group

of Normal rats received daily application of vehicle (N/Vehicle). Bar = 50 µm. (B) BrdU labeling

indexes (%) were calculated from the number of BrdU positive basal cells relative to the total

number of basal cells in sections of skin treated topically with either NTX or saline dissolved in

cream, or Regranex®; N/Vehicle rats received vehicle only. Values represent mean ± SEM.

Significantly different from N/Vehicle group at ***P<0.001; significantly different from

DB/Vehicle at ++P<0.01 and +++P<0.001.

66

2.5.5. Tissue Pathology and Immunohistochemical Studies

Hematoxylin and eosin stained sections of wounds did not show significant cell

death or structural pathology in response to treatment with either 0.03% NTX or

Regranex®.

2.5.6. PDGF Expression

Positive stained PDGF cells were located at the edge of the wound and deep in the

wound matrix, embedded within the granulation tissue. Cells were counted within a grid

and presented as the number of cells per mm2. Within 1 day of treatment, groups of DB

animals receiving either Regranex® or NTX had more than 1.5-fold (p<0.01) more PDGF+

stained cells than vehicle treated wounds. The elevation in PDGF+ cells continued on day

2 and day 4, whereas Regranex® treatment had no further effect, and levels returned to

those observed in the DB/Vehicle rats (Figure 2.3.).

67

Figure 2.3. PDGF expression in full-thickness skin wounds from type 1 diabetic (DB) rats receiving

daily treatment of a single drop of 0.03% NTX (DB/NTX) or saline (DB/Vehicle) dissolved in

moisturizing cream, or Regranex® (DB/R) for 1, 2, and 4 days. (A) Representative

photomicrographs of granulation tissue positively stained for PDGF on day 4 following wounding

(bar = 25 μm). (B) Values represent mean ± SEM number of PDGF positive stained cells within

one mm2. Significantly different from the DB/Vehicle group at *P<0.05 and **P<0.01;

significantly different between the DB/NTX and DB/R groups at +P<0.05.

2.5.7. VEGF Expression

The percentage of VEGF-positive stained blood vessels on day 4 was 43% more for

the NTX-treated group relative to diabetic controls (Figure 2.4.). Nearly 60% of all vessels

in the NTX-treated group were positive for VEGF, relative to less than 40% of the vessels

being VEGF-positive in either the control of Regranex®-treated rats.

68

Figure 2.4. VEGF expression in full-thickness skin wounds from type 1 diabetic (DB) rats receiving

daily treatment of a single drop of topical NTX or saline dissolved in moisturizing cream, or

Regranex® (DB/R) for 1, 2, and 4 days. (A) Representative photomicrographs of granulation tissue

with microvessels positively stained for VEGF on day 4 following wounding (bar = 25 μm). (B)

Values represent the percentage (± SEM) of VEGF-positive vessels within one mm2. Significantly

different from the DB/Vehicle group at ***P<0.001; significantly different between the DB/NTX

and DB/R groups at +++P<0.001.

2.5.8. FGF-2 Staining

Following wounding and initial mast cell infiltration, cytokines including FGF-2,

PDGF and VEGF are found circulating in the blood. FGF-2 cytokines are at the highest

levels within 24 hr of wounding. Examination of tissue surrounding the wound on days 1,

2 and 4 revealed that FGF staining was comparable among all treatment groups.

69

2.6. Discussion

Delayed wound healing is a major complication associated with diabetes,

particularly uncontrolled and/or chronic diabetes such as that associated with T2D.

Pressure-related wounds and diabetic foot ulcers affect nearly half of all persons with

diabetes at some point in their lifetime. In addition to the disabling side-effects of

unhealed wounds, the diabetic patients become more vulnerable to infections that may

in fact lead to more serious medical conditions requiring amputation [1-3]. Previous

studies in our laboratory have defined mechanisms that relate to reversing delayed

wound repair, particularly in T1D rats [13-15]. These preclinical studies have

demonstrated that topical application of NTX enhances epithelial cell replication,

angiogenesis, and increases cytokine profiles related to formation of skin [13,14].

Treatment with NTX enhanced wound closure by XXXX in T1D rats relative to rats

receiving three applications of vehicle only. DNA synthesis experiments revealed that

epithelialization was accelerated, as was blood vessel formation. These preclinical studies

were also confirmed in a mouse model of T2D [33]. The db/db mouse was surgically

wounded and 10-5M NTX dissolved in Vigamox® was applied three times daily. Wound

size was markedly reduced relative to diabetic mice receiving saline in Vigamox®. This

work corroborates studies on corneal surface epithelialization where topical NTX

enhanced repair of corneal abrasions in type 1 diabetic rats and rabbits, as well as type 2

diabetic mice [34].

In the present study, a novel formulation of NTX was used - 0.03% NTX dissolved

in moisturizing cream and the application was reduced to one time daily. This regimen

70

was consistent with, or even less, than that prescribed for Vigamox® therapy. The results

with one application of 0.03% NTX correlated well with those reported for 3 daily

applications of 10-5 M NTX [13-15]. In comparison to Regranex®, wound healing was

comparable in the T1D rat and both treatments accelerated wound closure relative to rats

receiving vehicle. Moreover, NTX targeted an underlying mechanisms associated with

diabetic complications by blocking the interfacing of OGF and its overexpression in

diabetes and OGFr. This overexpression of OGF inhibits cell replication and thus delays

many of the phases of wound closure. BrdU-labeling of the rats demonstrated that topical

0.03% NTX accelerated DNA synthesis and epithelial replication relative to both

Regranex® treatment and vehicle controls. Evaluation of cytokine expression

demonstrated that NTX also elevated VEGF expression whereas Regranex® had no effect

of this angiogenesis marker. Both Regranex® and NTX increased PDGF expression, and

NTX extended the enhanced expression for 48 hr relative to the effects seen following

Regranex® treatment that lasted only one day. FGF-2 expression did not change, at least

at the time points evaluated in these experiments.

The present investigation extends the work demonstrating that continuous

blockade of the OGF and OGFr regulatory pathway using NTX is an effective novel therapy

for non-healing ulcers in individuals with diabetes. The new formulation of 0.03% NTX

applied topically once daily substantiates an alternative, effective therapy for DFUs, and

lends support for clinical trials. Given that the population of diabetes is approaching 29

million in the United States [1], there is an urgent need to development safe, inexpensive,

71

and effective treatments for one of the many complications associated with the chronic

disorder.

2.7. Innovation

Topical application of an opioid antagonist for treatment of cutaneous wounds is

as effective as the standard of care and provides a safe, inexpensive, and effective

therapeutic for wound care in diabetes.

72

2.8. Key Findings

• Topical application of NTX accelerates closure of full thickness cutaneous wounds in

T1D rats

• Topical application of NTX heals full thickness cutaneous wounds at a rate comparable

to that of the standard care, Regranex®

• Topical NTX increases DNA synthesis in basal epithelial cells of the skin, whereas

Regranex® has little or no effect on cell replication

• Topical NTX increases PDGF+ stained cell number

• Topical NTX increases angiogenesis based on VEGF+ stained vessels

• Blockade of the OGF-OGFr axis with NTX targets an underlying pathway in diabetes

• Preclinical data support proof-of-concept clinical trials using topical NTX as a safe,

effective, and inexpensive method of treatment of diabetic foot ulcers

2.9. Acknowledgments and Funding Sources

Work was supported in part by private funding to the laboratories of Drs Zagon and

McLaughlin.

2.10. Author Disclosure and Ghostwriting: None

2.11. Abbreviations and Acronyms

BrdU, 5-bromo-2’-deoxyuridine

FGF-2, fibroblast growth factor -2

NTX, naltrexone

OGF, opioid growth factor

OGFr, opioid growth factor receptor

73

PDGF, platelet-derived growth factor

VEGF, vascular endothelial growth factor

74

2.12. References

[1] Center of Disease Control and Prevention. “Facts about diabetes: a leading cause of death

in the U.S. National Diabetes Educational Program.” 2013. 24 Nov. 2013.

[2] http://www.diabetes.org/living-with-diabetes/complications/

[3] Alberti KGMM, Zimmet PZ. Definition, diagnosis and classification of diabetes mellitus and

its complications. Part 1: Diagnosis and classification of diabetes mellitus. Provisional

Report of a WHO consultation. Diabetic Med 1998;15:539-553.

[4] http://www.diabetes.org/advocacy/news-events/cost-of-diabetes.html

[5] National Diabetes Information Clearinghouse (NDIC). National Institute of Diabetes and

Digestive and Kidney Disease (NIDDK). Insulin Resistance and Prediabetes. 2013. 25 Nov

2013.

[6] Diegelmann RF and Evans MC. Wound healing: an overview of acute, fibrotic and delayed

healing. Frontiers Bioscience 2004;9:283-289.

[7] Werner S, Grose R. Regulation of wound healing by growth factors and cytokines. Physiol.

Rev. 83: 835-870, 2002.

[8] Stadelmann WK, Digenis AG, Tobin GR. Physiology and healing dynamics of chronic

cutaneous wounds. Am J Surg 1998;176:26S-38S

[9] Gurtner GCX, Werner S, Barrandon Y, Longaker MT. Wound repair and regeneration.

Nature 453:314-321, 2008

[10] Lim Y-C, Bhatt MP, Kwon M-H, Park D, Na SH, Kim YM, Ha KS. Proinsulin C-peptide prevents

impaired wound healing by activating angiogenesis in diabetes. J Invest Dermatol 135:269-

278, 2015.

75

[11] Ikizaki S, Ito Y, Hosono K et ae. Vascular endothelial growth factor rec eptor Type-1

signaling prevents delayed wound healing in diabetes by attenuating the production of IL-

1β by recruited macrophages. Am J Pathol 186:1481-1498, 2016.

[12] Guillemin Y, Le Broc D, Segalen C, Kurkdjian E, Gouze JN. Efficacy of a collagen-based

dressing in an animal model of delayed wound healing. J Wound Care 25:406-413, 2016.

[13] McLaughlin PJ, Pothering CA, Immonen JA, Zagon IS. Topical treatment with the opioid

antagonist naltrexone facilitates closure of full-thickness wounds in diabetic rats. Exp Biol

Med 2011;236:1122–32.

[14] McLaughlin PJ, Immonen JA, Zagon IS. Topical naltrexone accelerates full-thickness wound

closure in type 1 diabetic rats by stimulating angiogenesis. Exp Biol Med 2013;238:733-

743.

[15] Immonen, JA, McLaughlin PJ, Zagon IS. Topical treatment with the opioid antagonist

naltrexone accelerates the remodeling phase of full-thickness wound healing in type 1

diabetic rats. Exp Biol Med 2013;238:1127-1135.

[16] http://www.regranex.com/ Faster Healing of Diabetic Neuropathic Ulcers. Accessed

December 2016.

[17] Chan RK, Liu PH, Pietramaggiori G et al. Effect of recombinant platelet-derived growth

factor (Regranex®) on wound closure in genetically diabetic mice. J Burn Care Res 27:202-

205, 2006.

[18] Niezgoda JA, Van Gils CC, Frykberg RG et al. Randomized clinical trial comparing OASIS

wound matrix to Regranex® gel for diabetic ulcers. Adv Skin Wound Care 18:258-266, 2005.

76

[19] Howard JD, Sarojini H, Wan R, Chien S. Rapid granulation tissue regeneration by

intracellular ATP delivery-A comparison with Regranex. PLoS One 9:e91787 2014.

[20] Blumberg H, Dayton HB. Naloxone, naltrexone, and related noroxymorphones. In: Braude

MC, Harris LC, May EL, Smith JP, Villarreal JE, eds. Advances in Biochemical

Psychopharmacology. 2nd ed. Narcotic Antagonists. New York: Raven Press, 1974:33–43.

[21] Gutstein HB, Akil H. Opioid analgesics. In: Hardman JG, Limbard LE, eds. The

Pharmacological Basis of Therapeutics, 10th edition. New York: McGraw Hill, 2001:569–

619.

[22] McLaughlin, P.J. and I.S. Zagon. 2015. Duration of opioid receptor blockade determines

clinical response. Biochem. Pharmacol. 97:236-246.

[23] Zagon IS, Donahue RN, McLaughlin PJ. Opioid growth factor – opioid growth factor

receptor axis is a physiological determinant on cell proliferation in diverse human cancers.

Am J Physiol Regul Integr Comp Physiol 2009;297:R1154–61.

[24] Zagon IS, McLaughlin PJ. Naltrexone modulates tumor response in mice with

neuroblastoma. Science 1983;221:671–3.

[25] Zagon IS, McLaughlin PJ. Duration of opiate receptor blockade determines tumorigenic

response in mice with neuroblastoma: a role for endogenous opioid systems in cancer. Life

Sci1984;35:409–16.

[26] Donahue RD, McLaughlin PJ, Zagon IS. Cell proliferation of human ovarian cancer is

regulated by the opioid growth factor – opioid growth factor receptor axis. Am J Physiol

Regul Integr Comp Physiol 296:R1716–25, 2009.

77

[27] Fallucca F, G Tonnarini, N Di Biase, M D'Alessandro, M Negri. Plasma met-enkephalin levels

in diabetic patients: Influence of autonomic neuropathy. Metabolism 45:1065-1068, 1996.

PMID:8781292

[28] Negri M, G Tonnarini, M D'Alessandro, F Fallucca. Plasma met-enkephalin in type 1

diabetes. Metabolism 41:460-461, 1992. PMID:1588823

[29] Timmers K, NR Voyles, C Zalenski, S Wilkins, L Recant. Altered β-endorphin, met- and leu-

enkephalins, and enkephalin-containing peptides in pancreas and pituitary of genetically

obese diabetic (db/db) mice during development of diabetic syndrome. Diabetes 35:1143-

1151, 1986. PMID:2944783

[30] King A, Bowe J. Animal models for diabetes: understanding the pathogenesis and finding

new treatments. Biochem Pharmacol 99:1-10, 2016.

[31] Van der Loos CM, Meijer-Jorna LB, Broekmans MEC, Pleogmakers HPHM, Teeling P, de

Boer OJ, van der Wal AC. Anti-human vascular endothelial growth factor (VEGF) antibody

selection for immunohistochemical staining of proliferating blood vessels. J Histochem

Cytochem 58:109-118, 2010.

[32] Lim Y-C, Bhatt MP, Kwon M-H, et al. Preinsulin C-peptide prevents impaired wound healing

by activating angiogenesis in diabetes. J Invest Dermatol 135:269-278, 2015.

[33] Klocek MS, Sassani JW, McLaughlin PJ, Zagon IS. Topically applied naltrexone restores

corneal reepithelialization in diabetic rats. J Ocul Pharmacol Ther 2007;23:89-102.

[34] Sassani J.W., McLaughlin PJ, Zagon IS. The Yin and Yang of the opioid growth regulatory

system: Focus on diabetes: The Lorenz E. Zimmerman Tribute Lecture. J. Diabetes Res.,

Volume 2016, article ID 9703729. http://dx.doi.irg/10.1155/2016/9703729.

78

[35] Zagon I.S., J.W. Sassani, J.A. Immonen and P.J. McLaughlin. 2014. Ocular surface

abnormalities related to Type 2 diabetes are reversed by the opioid antagonist naltrexone.

Clin. Exp. Ophthalmol. 42:159-168. PMID: 23777539.

[36] Immonen, J.A., I.S. Zagon, and P.J. McLaughlin. 2014. Topical naltrexone as treatment for

type 2 diabetic cutaneous wounds. Advances Wound Care 3: 419-427. PMID:24940556

[37] Immonen, J.A., I.S. Zagon, and P.J. McLaughlin. 2014. Selective blockade of the OGF-OGFr

pathway by naltrexone accelerates fibroblast proliferation and wound healing. Exp. Biol.

Med. 239:1300-1309. PMID:25050485

79

2.13. About the authors

Patricia J. McLaughlin, M.S., D.Ed., is Professor of Neural and Behavioral Science at Penn

State University College of Medicine.

Jarrett Cain, DPM is a podiatric surgeon at Penn State Hershey and co-Director of the Foot

Clinic. Dr. Cain has an active service treating diabetic foot ulcers and other complications

from diabetes.

Michelle Titunick, BA completed these experiments as a portion of her doctoral research

in the Anatomy Graduate Program at Penn State University College of Medicine.

Ian S. Zagon, M.S., Ph.D, is Distinguished University Professor at Penn State University

College of Medicine.

80

CHAPTER 3: THE OPIOID GROWTH FACTOR-OPIOID GROWTH FACTOR RECEPTOR

AND DIABETIC BONE COMPOSITION

M.B. Titunick, J.D. Cain, I.S. Zagon, P.J. McLaughlin

81

3.1. Abstract

3.1.1. Background:

Complications from diabetes, a disease affecting more than 29 million individuals

in the United States, include increased fracture rates and delays in bone fracture repair

that contribute to the healthcare costs of $322 billion. Treatments to accelerate bone

healing are needed.

3.1.2. Methods:

Type 1 diabetes was induced in rats by injection of streptozotocin. Open fractures

were created in the right femur of 5 week hyperglycemic animals and fixed internally.

Naltrexone (5x10-5 M) was dissolved in calcium sulfate solution used for fixation and

applied generously to the fracture site. Animals were allowed to recover for 7 days prior

to euthanasia. Callus tissue and bone were prepared for morphological studies. Tissues

were immunohistochemically stained for expression of opioid growth factor (OGF), OGF

receptor (OGFr), as well as for osteoclasts, cartilage and bone by staining with tartrate

resistant alkaline phosphatase and safranin O, respectively. Antibodies to osteocalcin and

Ki-67 were used to assess osteoblasts and proliferation rates.

3.1.3. Results:

Semiquantitative analyses of OGF and OGFr indicated the presence of both

peptide and receptor throughout development in rat bone tissue, and increased

expression levels in diabetic rat bone. Importantly, serum levels of OGF, chemically

termed [Met5]-enkephalin, by ELISA indicated elevated serum OGF levels in diabetic rats

relative to values in non-diabetic rats. Diabetic bone was diminished with respect to area

82

of cartilage, proliferative cells, and VEGF+ cells. Analysis of callus tissue revealed an

increase in cartilage and bone, and compensatory decrease in granulation tissue in NTX-

treated diabetic fractures compared to vehicle-treated diabetic fractures.

3.1.4. Conclusions:

Diabetes is associated with defective levels of the inhibitory peptide OGF leading

to delayed proliferation. Bone healing processes were supported by addition of NTX to

block the interaction of OGF and OGFr. These data suggest that additional studies are

warranted to assess the value of NTX as a therapeutic in diabetic bone repair.

3.1.5. Keywords:

Naltrexone, type 1 diabetes, bone composition, enkephalin, femur fracture

3.2. Abbreviations:

DAPI, 4’,6- diamidino-2-phenylindole

DB, type 1 diabetes

ELISA, enzyme-linked immunosorbent assay

NTX, naltrexone hydrochlorid

OGF, opioid growth factor

OGFr, opioid growth factor receptor

PBS, phosphate buffered saline

SEM, standard error of the mean

T1D, type 1 diabetes

TRAP, tartrate resistant alkaline phosphatase

83

3.3. Introduction

An estimated 30.3 million people in the U.S. have diagnosed or undiagnosed

diabetes, of which 5-10% is type 1 diabetes (T1D) [1]. According to the 2017 report by

the Center for Disease Control, healthcare costs for diabetes costs in the U.S. are

estimated to be $322 billion [2]. Complications arising from diabetes include poor wound

healing, increased risk of bone fracture, and delayed fracture healing [1-4]. In human

studies, diabetes has been associated with decreased linear bone growth, lower bone

mineral density, increased fracture risk, and poor bone healing [5-9]. Preclinical changes

in bone resorption, trabecular connectivity, bone porosity, and spicule/marrow space

ratio, as well as decreased osteoblastic recruitment have been reported in animal models

of diabetes [10, 11]. Collagen reduction was noted within two weeks post diabetes

induction in a rat model of diabetes followed by prolonged fracture callus maturation and

delayed bone bridging [10]. Complications associated with diabetes in mice have also

been shown to disrupt normal bone formation by reducing cellular processes required for

osteoblast formation and differentiation [11]. Specific causes of these diabetic

complications are unclear, but the high cellular glucose levels that increase reactive

oxygen species and pro-inflammatory cytokine release may be contributing factors to

defects in bone repair [12, 13].

Current therapies to increase bone quality are not completely effective [14-16].

Despite the lack of a direct correlation between bone density and risk for fracture in type

2 diabetes, calcium supplementation, strict regulation of insulin and glucose, diet, and

exercise are often recommended to limit the risk of osteopenia and osteoporosis [14,15].

84

These health initiatives are secondary to the primary complication of delayed bone repair;

thus, there remains an unmet medical need to identify specific treatments for bone

healing in diabetic individuals.

Another modifier of the healing process is the neuropeptide methionine

enkephalin or opioid growth factor (OGF) [17]. OGF is an endogenous opioid peptide and

negative growth regulator that binds selectively to OGFr to maintain cellular homeostasis.

The mechanism of action involves alteration of the p16 or p21 cyclin-dependent inhibitory

kinases that alter the G0/G1 phase of the cell cycle and inhibit DNA synthesis [18]. In

preclinical and clinical studies, increased plasma levels of [Met5]-enkephalin have been

reported in diabetic animals and humans [19-21], suggesting that the tonic interaction

impacting cell homeostasis is disrupted favoring inhibited cell replication. Blockade of the

OGF-OGFr pathway with opioid antagonists such as naltrexone (NTX) that provide a

continuous receptor blockade results in enhanced cell proliferation [22]. Studies in our

laboratory using sufficient dosages of NTX to produce a continuous blockade of OGFr have

shown efficacy in the treatment of diabetic corneal wounds [23,24] and full-thickness

cutaneous wounds in diabetic (type 1 diabetes) [25-27] and db/db mice (type 2 diabetes)

[28]. The mechanism of action in these studies indicated increased proliferation of

epithelial cells [27, 29], endothelial cells contributing to angiogenesis [25], and

granulation tissue formation [26].

In this study, we investigated the hypothesis that excess OGF ([Met5]-enkephalin)

in diabetes may contribute to changes in bone quality, and that blockade of the OGF-OGFr

axis will restore diabetic bone quality, and fracture repair processes.

85

3.4. Materials and Methods

3.4.1. Animals and induction of diabetes

Male 6-week-old Sprague Dawley rats (Charles River Laboratories, Wilmington,

MA) were housed under controlled environmental conditions with food and water ad

libitum. All protocols were approved by Penn State College of Medicine Institutional

Animal and Use Committee. Type 1 diabetes was induced in a subset of rats by

intraperitoneal injection of streptozotocin (40 mg/kg; Sigma) on two consecutive days;

control rats received sodium citrate buffer only [23,27]. Serum glucose levels were

measured with a TRUETrack® glucometer (Nipro Diagnostics). Rats with a blood glucose

reading of at least 250 mg/dL were considered to be hyperglycemic.

3.4.2. Bone fracture surgical model

Under sterile conditions 37 male rats were anesthetized using a cocktail of

ketamine (50mg/kg), xylazine (5mg/kg), and acepromazine (1mg/kg). Animals were

shaved between the thigh and knee, sprayed with 70% ethanol and swabbed with iodine.

An open mid-diaphyseal fracture was created by opening the skin superficial to the femur

and cutting through the quadricep muscles to expose the distal portion of the femur. A

bore hole was created by a 22 G needle and a 25 G spinal needle was used as internal

fixation. The spinal needle was retracted from the marrow cavity to create the fracture

using a Dremel saw. The two ends were attached using the spinal needle, fixing the

needle’s end into the proximal portion of the femur. Surgical Simplex® P calcium sulfate

with either 5x10-5M NTX or sterile water was applied to the fracture bone. The calcium

sulfate degraded over time and released naltrexone in situ. The fracture site was closed

86

and skin sutured; animals were given carprofen (0.2ml/100g) during surgery and as

needed following the procedure. The rats were euthanized 1 week after surgery with

Euthasol (Virbac Animal Health).

3.4.3. Immunohistochemistry

Femurs were fixed in 10% neutral buffered formalin for 3 days and placed in 14%

ethylenediaminetetra-acetic acid for 2-3 weeks to decalcify. Tissue was dehydrated in

increasing dilutions of ethanol and xylene, embedded in paraffin, and sectioned at 5 µm.

The sections were rehydrated and incubated with pepsin for 10 minutes for antigen

retrieval. Following washes, tissue was blocked with bovine serum albumin for 1 hour.

Primary antibodies (1:200 OGF, 1:200 OGFr, 1:500 osteocalcin) were incubated overnight

at 4°C; Ki67 (1:300) and VEGF (1:100) were incubated for 30 min at room temperature.

Anti-OGF was generated in our laboratory [30], and all other antibodies were purchased

from the following sources: OGFr (Bethyl Laboratories, Montgomery, TX), Ki67 (Merck

Millipore, Burlington, MA), osteocalcin (Santa Cruz Biotechnology, Inc, Dallas TX), and

VEGF from Neo Markers (ThermoFisher Scientific; Waltham, MA). Following primary

antibody incubation, tissues were washed and incubated with goat anti-rabbit secondary

antibody (Alexa Fluor® 568, A11011, ThermoFisher) for 2 hours. Coverslips were

mounted using 50:50 glycerol:PBS with DAPI (1:2000). Intact bone was assessed at the

mid-shaft, whereas fractured bone was evaluated within the callus. Fluorescent images

were taken with an Olympus IX81 and Spot RT3 camera, and cell number evaluated in 3-

5 sections per bone, 3-6 bones per treatment group. Semiquantitative measurements of

TRITC intensity were assessed using Slidebook 5.0 (Silicon Graphics).

87

3.4.4. Histomorphometry

To assess composition of the callus, bones were processed as described above for

paraffin embedding, and sections stained with hematoxylin, fast green, and safranin O

[31] to determine the percent area of granulation tissue (where present), cartilage, and

bone. Adjacent sections were stained with tartrate resistant alkaline phosphatase (TRAP)

by staining with naphthol AS-MX phosphate, Fast Red (>1 hour) and counterstained with

methyl green to evaluate the number of osteoclasts [32]. The percentage of cartilage or

bone was assessed in intact femur, as well as in the callus region of fractured bone by

placing a grid adjacent to the fracture edge. Area measurements were collected to

determine the percentage of cartilage per grid, as well as calculation of cartilage for the

entire area of the callus. Stained sections were photographed on an Olympus BX51 with

a Spot RTKE camera. Areal analysis of safranin O and TRAP were assessed using ImageJ

[33].

3.4.5. Measurement of serum enkephalin

Blood samples from T1D and normal, non-diabetic rats were collected

immediately post-mortem and allowed to coagulate on ice; serum was separated and

frozen at -80oC until assayed. Levels of [Met5]-enkephalin were measured in duplicate

using an ELISA kit (MyBioSource.com; MBS9342519).

3.4.6. Statistical analyses

Data were analyzed using GraphPad Prism 7.0 software (Graph Pad). In most

cases, data were analyzed with one-way analysis of variance (ANOVA) followed by

88

Newman-Keuls post-hoc test. Student’s two-tailed unpaired t-test was used where

indicated. Significance was determined at α-value of 0.05 for all tests.

3.5. Results

3.5.1. Expression levels of OGF and OGFr during bone development

Throughout development (1 day to 4 months), there were no significant

differences in bone OGF levels (Fig.1A) with mean intensity ranging from 384.1 on day 21

to 487.5 on day 5. In contrast, OGFr levels were significantly higher immediately following

birth (1 day) in comparison to all other time points through 4 months of age. OGFr

expression was 1201 ± 127.6 on day 1 in comparison to levels of 266 ± 36.5 on day 5 and

643 ± 118.5 on day 21 (Fig. 1B). Mean values on days 5, 10, 30, and 120 were significantly

less than those recorded on days 1 and 21.

89

3.5.2. Levels of OGF and OGFr in bone and serum of T1D animals

Figure 3.1.: Expression of opioid growth factor (OGF) (A) and opioid growth factor

receptor (OGFr) (B) in long bone of developing rats on days 1, 5, 10, 21, 30, and 4 months

of age. Histograms represent mean ± SEM intensity of TRITC. Significantly different from

day 1 at **** p<0.0001; significantly different from day 21 at + p<0.05 and ++ p<0.01.

90

Figure 3.2.: Expression of OGF and OGFr in long bone of type 1 diabetic (DB) and normal

(N) rats. Immunofluorescence images of OGF (A) and OGFr (C) stained bone with merged

DAPI staining. Scale bar=10um. Inset shows secondary only stained tissue.

Semiquantitative immunohistochemical assessment of OGF (B) and OGFr (D) in sections

of femur from DB and N rats. Histogram represents mean ± SEM intensity TRITC.

Significantly different from N values at *p<0.05. (E) Serum levels of [Met5]-enkephalin

(ng/ml) in DB and N rats assayed by ELISA. Values represent mean ± SEM. Diabetic serum

levels of OGF differed from normal serum levels at * p=0.0327

91

Immunohistochemical staining for OGF and OGFr in intact femur tissue of 14-

week-old T1D (hyperglycemic for 6 weeks) and normoglycemic male Sprague-Dawley rats

demonstrated specific staining (Figs 2A, C). Semiquantitative analysis of TRITC

fluorescence in intact cortical bone revealed that T1D femurs exhibited a mean intensity

of OGF-TRITC staining of approximately 495 units in comparison to normal femurs with a

mean intensity of OGF-TRITC staining of 253.6 units, with a significant difference p<0.05

(Fig 2B). OGFr staining in T1D cortical bone was assessed at approximately 645 units, an

84% increase (p<0.05) from the TRITC intensity measured in cortical bone of

normoglycemic rats (Fig. 2D).

Serum levels of OGF were 3.5 ± 0.2 ng/ml in normal non-fractured rats and 4.3 ±

0.2 ng/ml in the serum of T1D rats; this represented a 23% increase over baseline or

normal values (Fig. 2E). Blood collected 7 days after fracture surgery was assayed, and

there were no differences in OGF levels in the serum.

3.5.3. Cellular composition of T1D bone

The cellular composition of intact bone tissue from T1D and normal male rats was

assessed following TRAP and revealed no significant differences in the number of

osteoclasts. Evaluation of the bone tissues with safranin O indicated a 73% decrease in

the area of calcified cartilage in diabetic rats (Fig. 3A). Tissue stained for proliferation

revealed a 66% decrease in Ki67 positive cells in diabetic bones compared to normal

bones (20%) (Fig. 3B). VEGF staining to assess the number of endothelial cells associated

with blood vessels decreased to approximately 30% in diabetic bones compared to

approximately 41% in normal bones (Fig. 3C).

92

Figure 3.3.: Analyses of bone tissue and markers in DB and N adult male rats. (A) Safranin

O stained sections were analyzed for the area (percent) of cartilage. Cortical bone sections

were evaluated for the percentage of Ki67+ cells (B) and VEGF+ vessels (C). Values

represent means ± SEM for 1-5 sections/rat and 6 rats per group. Significantly different

from N values at *p<0.05 and **p<0.01.

3.5.4. Cellular composition of T1D calluses following fracture

Experiments were conducted twice with a total of 7 normal and 13 diabetic rats

monitored. Femurs collected 7 days following fracture were analyzed to assess cellular

composition in the entire callus area, as well as within a smaller grid adjacent to the

fracture edge which may represent a more actively dividing region. Tissue from intact

femurs exhibited a 73% increase in cartilage of normal femurs compared to diabetic

93

femurs. There is a significant 2% increase in bone of diabetic femurs (99%) compared to

normal femurs (97%) (Fig. 4).

Figure 3.4.: Callus in N and DB rats following femur fracture. Fractures were treated with

sterile water (DB/V) or 5x10-5M NTX (DB/NTX) dissolved in calcium sulfate bone cement

and topically applied to the fracture site. (A) Photographs of safranin O stained fracture

calluses at 7 days post-fracture. Arrowheads indicate the location of a 1mm2 counting

grid. (B) Histogram represents composition (percentage) of total callus tissue 7 days post-

fracture for normal vehicle-treated (N/Vehicle), type 1 diabetic vehicle-treated

(DB/Vehicle), and type 1 diabetic naltrexone-treated (DB/NTX) fractures. No differences

were noted. Values were collected from 9-13 sections/rat and 4-6 rats per treatment

group.

94

With regard to granulation tissue in fractured bone, there is 84% more granulation

tissue remaining in the callus of DB vehicle-treated rats at 7 days post-fracture than in

N/Vehicle calluses, and 52% more than in DB NTX-treated calluses (Fig. 5A). NTX-treated

DB rat calluses had 88% more cartilage than DB vehicle-treated calluses (Fig. 5B). There

was 24% less bone in DB vehicle-treated calluses than in N/V calluses and 35% more bone

in DB NTX-treated calluses than in DB vehicle-treated calluses (Fig. 5C). DB rats treated

with vehicle showed significantly less Ki67 staining than normal rat bones (Fig. 6). NTX

treatment of DB rats increased Ki67 staining of the callus by 35% compared to DB rats

treated with vehicle only (32%) (Fig. 6).

Blood was collected 7days after fracture. OGF serum levels increased 28% in DB

rats with fractures treated with vehicle compared to the serum of DB rats without a

fracture (4.3 ng/ml); OGF serum levels were 13% less in DB rats treated with NTX relative

to that recorded for diabetic vehicle-treated rats.

95

Figure 3.5.: Composition of callus tissue measured at the fracture site. Areas (percent)

of granulation tissue (A), cartilage (B), and (C) bone within 1mm2 grids of callus tissue

adjacent to fracture site as indicated by the arrowheads in Figure 3.3. at 7 days post-

fracture for normal vehicle-treated (N/Vehicle), type 1 diabetic vehicle-treated

(DB/Vehicle), and type 1 diabetic naltrexone-treated (DB/NTX) fractures. Fractures were

treated with H2O or 5x10-5M NTX within calcium sulfate bone cement. Histograms

represent means ± SEM; values were collected from multiple sections/rat and 4-6 rats per

treatment group. Significantly different from N/Vehicle at **p<0.01 or *** p<0.001;

significantly different between DB treatment groups at +p<0.05 (Student’s t-test) or +++

p<0.001.

96

Figure 3.6.: Proliferative cells in callus tissue at 7 days following femur fracture in DB and

N rats. (A) Photomicrographs of Ki67+ cells in normal vehicle-treated (N/Vehicle), type 1

diabetic vehicle-treated (DB/Vehicle), and type 1 diabetic naltrexone-treated (DB/NTX)

fractured rat femurs; Bar = 10 µm. (B) Histogram represents the percent Ki67+ cells within

1mm2 grids of callus tissue adjacent to fracture site as indicated by the arrowheads in

Figure 3.3. Values represent means ± SEM. Significantly different from N/Vehicle at

*p<0.05 and significantly different between DB treatment groups at +p<0.05.

97

3.6. Discussion

While T1D is more often associated with reduced bone mass and increased risk of

fracture, type 2 diabetes is associated with increased risks of hip and foot fractures [34].

The impact of diabetes on the normal stages of bone healing has been extensively

documented to show increased required healing times that can lead to delayed union [4].

This study investigated the presence and functioning of the OGF-OGFr axis and

diabetic bone quality. OGF and OGFr are present in bone throughout development and

into adulthood. There was an increased expression of OGF in diabetic bone as well as in

the serum of diabetic rats. The OGF-selective receptor, OGFr, was also reported to be

elevated in T1D rats suggesting that this pathway, involved with homeostatic cellular

replication, is aberrant in diabetes. These findings in bone are not unique, as OGF and

OGFr levels are elevated in epithelial tissues.

Utilizing NTX to block OGFr limits the activity of OGF providing a potential

therapeutic to increase fracture healing in diabetes. In this study we began to examine

callus composition in the presence of NTX following fracture. With a follow-up of only 7

days, it was evident that topical application of NTX may have an effect on the proportion

of cartilage and bone within the callus. Histomorphometric analyses indicated that NTX-

treated diabetic fracture calluses contained more cartilage and bone and less granulation

tissue than vehicle-treated diabetic fractures. This would support our hypotheses that

NTX blocks the inhibitory action of OGF and therefore enhance the timing of bone repair.

Reports on well controlled diabetes demonstrate there is delayed fracture healing

suggesting that hyperglycemia alone is not responsible for the delay in bone healing.

98

Elevated serum levels of the inhibitory growth factor OGF may contribute to the delayed

cellular renewal required for bone fracture repair.

One of the limitations of the study is the evaluation of the treatment after only 7

days. This study was limited to 7 days post-fracture. While the initial phases of bone

healing involve stimulation by inflammatory responses of recruitment of mesenchymal

stem cells that lead to differentiation, a further study will be necessary to investigate the

time frame of bone bridging, and the mechanical properties of the bone once fully healed.

Future studies also need to measure the release rate of NTX from the calcium sulfate to

determine effective dosage level. In addition, damage to the muscle and periosteum, two

of three major blood supplies to the bone, are damaged in the open fracture model, and

potentially limit vascularity to the wound.

Despite the experimental limitations, the current data support and extend our

knowledge about the modulation of the OGF-OGFr axis and complications of diabetes. In

preclinical studies utilizing both type 1 and type 2 diabetes animal models, delayed wound

closure was reversed following topical application of NTX [26, 28]. The mechanism of

action appeared to be enhanced cell replication, with the mechanism of action utilizing

the specific OGF receptor action site [29]. The growing evidence of an increase in OGF in

diabetes and the potential for dysregulation of the OGF-OGFr regulatory axis that plays a

role in cellular homeostasis supports the need for further study on NTX as a therapeutic

agent for diabetic fracture repair. Moreover, research is encouraged to examine the use

of systemic NTX to thwart other bone diseases including osteopenia or osteoporosis.

99

3.7. Conclusions

The OGF-OGFr regulatory axis is present in long bone throughout rat

development. However, in type 1 diabetic models, OGFr is overexpressed in tissue and

OGF is overexpressed in tissue and serum possibly contributing to the decreased quality

of long bone, and the changes observed in a small window of time in callus tissue

following femur fracture. Blockade of the OGF-OGFr pathway with NTX appears to

increase cell replication and capillary formation, leading to accelerated bone repair in

diabetes.

3.8. Acknowledgements

Special thanks to Michael D. Ludwig, PhD who helped throughout the surgeries.

3.9. Author contributions

Study design: MBT, PJM; Study conduct: MBT, JDC; Data collection and analyses: MBT,

PJM

Data interpretation: MBT, PJM, JDC; Drafting, editing and approving final manuscript:

MBT, JDC, ISZ, PJM

3.10. Funding

Research was supported in part by discretionary gift funds to ISZ and PJM, as well

as a grant with the Pennsylvania Department of Health using Tobacco CURE Funds (PJM).

The Department specifically disclaims responsibility for any analyses, interpretations or

conclusions.

3.11. Competing interests - None.

100

3.12. References

[1] National Center for Chronic Disease Prevention and Health Promotion (CDC), National

Diabetes Statistics Report, 2017 http://www.diabetes.org/assets/pdfs/basics/cdc-

statistics-report-2017.pdf

[2] American Diabetes Association, The Staggering Costs of Diabetes.

http://www.diabetes.org/diabetes-basics/statistics/infographics/adv-staggering-cost-of-

diabetes.html

[3] Marin C, Lyuyten FP, Van der Schueren B, Kerckhofs G, Vandamme K. The impact of type

2 diabetes on bone fracture healing. Front. Endocrinol. 9:6

doi:10.3389/fendo.2018.00006, 2018.

[4] Jiao H, Xiao E, Graves DT. Diabetes and its effect on bone and fracture healing. Curr.

Osteoporos. Rep. 13 (2015) 327-335.

[5] Giannini C, Mohn A, Chiarelli F. Growth abnormalities in children with type 1 diabetes,

juvenile chronic arthritis, and asthma. Int. J. Endocrinol. vol 2014 article ID 265954.

[6] Vestergaard, P. Discrepancies in bone mineral density and fracture risk in patients with

type 1 and type 2 diabetes—a meta-analysis. Osteoporos. Int. 18 (2007) 427–444.

[7] Janghorbani M, Van Dam R M, Willett WC, Hu F B. Systematic review of Type 1 and Type 2

diabetes mellitus and risk of fracture. Am. J. Epidemiol. 166 (2007) 495–505.

[8] Hofbauer LC, Brueck CC, Singh SK, Dobnig H. Osteoporosis in patients with diabetes

mellitus. J. Bone Miner. Res. 22 (2007) 1317–1328.

101

[9] Retzepi M, Calciolari E, Wall I, Lewis MP, Donos N. The effect of experimental diabetes

and glycaeemic control on guided bone regeneration: histology and gene expression

analyses. Clin.Oral Impl. Res. 29 (2018) 139-154.

[10] Kayal RA, Tsatsas D, Bauer MA, Allen B, Al-Sebaei MO, Kakar S, Leone CW, Morgan EF,

Gerstenfeld LC, Einhorn TA, Graves DT. Diminished bone formation during diabetic

fracture healing is related to the premature resorption of cartilage associated with

increased osteoclast activity. J. Bone Miner. Res. 22 (2007) 560-568.

[11] Lu H, Kraust D, Gerstenfeld LC, Graves DT. Diabetes interferes with the bone formation by

affecting the expression of transcription factors that regulate osteoblast differentiation.

Endocrinology 144 (2003) 346-352.

[12] Yamagishi S. Role of advanced glycation end products (AGEs) in osteoporosis in diabetes.

Curr. Drug Targets 12 (2011) 2096-2102

[13] Cruz NB, Sousa LP, Sousa MO, Pietrani NT, Fernandes AP, Gomes KB. The linkage between

inflammation and type 2 diabetes. Diab. Res. Clin. Pract. 99 (2013) 85-92.

[14] Almubarak S, Nethercott H, FreebergM, Beaudon C, Jha A, Jackson W, Marcucio R, Miclau

T, Health K, Bahney C. Tissue engineering strategies for promoting vascularized bone

regeneration. Bone 83 (2016) 197-209

[15] Eimar H, Alebrahim S, Manickam G, AL-Subaie A, Abu-Nada L, Murshed M, Tamimi F.

Donepezil regulates energy metabolism and favors bone mass accrual. Bone 84 (2016)

131-138.

[16] Wey A, Cunningham C, Hreha J, Breitbart E, Cottrell J, et al. Local ZnCl2 accelerates fracture

healing. J Orthopaedic Res 32 (2014) 834-841.

102

[17] Zagon IS, Wu Y, McLaughlin PJ. Opioid growth factor inhibits DNA synthesis in mouse

tongue epithelium in a circadian rhythm-dependent manner. Amer. J. Physiol. 267 (1994)

R645-R652.

[18] Cheng F, McLaughlin PJ, Verderame MF, Zagon IS. The OGF-OGFr axis utilizes the p16INK4a

and p21WAF1/CIP1 pathways to restrict normal cell proliferation. Mol. Biol. Cell 20 (2009)

319-327. PMCID:PMC2613082 PMID: 18923142

[19] Negri M, Tonnarini G, D’Alessandro M, Fallucca F. Plasma met-enkephalin in type I

diabetes. Metabolism. 41 (1992) 460–461.

[20] Greenberg J, Ellyin F, Pullen G, Ehrenpreis S, Singh SP, Cheng J. Methionine-enkephalin

and β-endorphin levels in brain, pancreas, and adrenals of db/db mice. Endocrinology

(1985) 116: 328-331.

[21] Fallucca F, Tonnarini G, Di Blasé N, D’Allessandro M, Negri M. Plasma met-enkephalin

levels in diabetic patients: influence of autonomic neuropathy. Metabolism (1996) 45:

1065-1068.

[22] McLaughlin PJ, Zagon I.S. Duration of opioid receptor blockade determines clinical

response. Biochem. Pharmacol. 97 (2015) 236-246.

[23] Klocek MS, Sassani JW, McLaughlin P J, Zagon IS. Topically applied naltrexone restores

corneal reepithelialization in diabetic rats. J. Ocul. Pharmacol. Ther. 23 (2007) 89–102.

[24] Zagon IS, Sassani JW, Carroll M A, McLaughlin PJ. Topical application of naltrexone

facilitates reepithelialization of the cornea in diabetic rabbits. Brain Res. Bull. 81 (2010)

248–255.

103

[25] McLaughlin PJ, Immonen JS, Zagon IS. Topical naltrexone accelerates full-thickness wound

closure in Type 1 diabetic rats by stimulating angiogenesis. Exp. Biol. Med. 238 (2013) 733-

743. PMID: 23788174. Doi: 10.1177/1535370213492688

[26] Immonen JA, Zagon I S, Lewis G S, McLaughlin PJ. Topical treatment with the opioid

antagonist naltrexone accelerates the remodeling phase of full-thickness wound healing in

type 1 diabetic rats. Exp. Biol. Med. 238 (2013) 1127–1135.

[27] McLaughlin PJ, Cain J D, Titunick MB, Sassani JW, Zagon IS. Topical naltrexone Is a safe and

effective alternative to standard treatment of diabetic wounds. Adv. Wound Care 6 (2017)

279–288.

[28] Immonen JA, Zagon IS, McLaughlin, PJ. Topical naltrexone as treatment for type 2 diabetic

cutaneous wounds. Adv. Wound Care 3 (2014) 419-427. PMID:24940556

[29] Immonen JA, Zagon IS, McLaughlin PJ. Featured Article. Selective blockade of the OGF-

OGFr pathway by naltrexone accelerates fibroblast proliferation and wound healing. Exp.

Biol. Med. 239 (2014)1300-1309.

[30] Zagon IS, McLaughlin PJ. Production and characterization of polyclonal and monoclonal

antibodies to the zeta (ξ) opioid receptor. Brain Res. 630 (1993), 295–302.

[31] Schmitz N, Laverty S, Kraus VB, Aigner T. Basic methods in histopathology of joint tissues.

Osteoarthritis Cartilage 18 (2010) S113–S116.

[32] Erlebacher A, Derynck R. Increased expression of TGF-beta 2 in osteoblasts results in an

osteoporosis-like phenotype. J. Cell Biol. 132 (1996) 195–210.

[33] Schneider CA, Rasband WS, Eliceiri KW. NIH Image to ImageJ: 25 years of image analysis.

Nat. Methods 9 (2012) 671–675.

104

[34] Adami S. Bone health in diabetes: considerations for clinical management. Curr Med Res

Opin 25 (2009) 1057-1072.

105

CHAPTER 4: THE OPIOID ANTAGONIST NALTREXONE PROTECTS

BONE IN A DIABETIC RAT MODEL

M.B. Titunick, G.S. Lewis, I.S. Zagon, P.J. McLaughlin

106

4.1 ABSTRACT

4.1.1. Aims:

Type 1 diabetes (DB) may result in a dysregulated Opioid Growth Factor (OGF)-

Opioid Growth Factor Receptor (OGFr) regulatory pathway. This research investigated

whether modulation of this axis by naltrexone (NTX) alters bone characteristics in DB rats.

4.1.2. Methods:

DB was induced by streptozotocin (n=16 male rats); 4 control rats received buffer.

Hyperglycemic animals were injected i.p. daily for 3 weeks with either 30 mg/kg NTX or

phosphate buffered saline; normoglycemic rats were untreated. At 21 days, bones were

processed for immunohistochemistry, μCT scanning, or 3-point bending to failure.

4.1.3. Results:

Relative to normal bone, DB decreased the strength (26%), osteocalcin expression

(17%), and Ki67 staining (33%) of femurs, and increased OGFr levels (126%). Systemic NTX

treatment increased the strength (21%) and energy absorbed (105%) in bone tissue relative

to tissues from saline-treated rats. Cortical and cross-sectional areas of femurs decreased

with diabetes.

4.1.4. Conclusions:

Bones from rats with diabetes were weaker and absorbed less energy than normal

bones, had increased OGFr expression, and decreased expression of osteocalcin and Ki67.

Three weeks of systemic NTX decreased OGFr levels and restored strength and energy

absorption, suggesting that NTX may protect bone quality in diabetes.

107

4.1.5. Keywords: Naltrexone, type 1 diabetes, bone composition, enkephalin

108

4.2. INTRODUCTION

More than 9% of the U.S. population has diabetes with 5-10% of the individuals

diagnosed with type 1 diabetes[1]. The estimated healthcare cost of diabetes in the U.S.

exceeds $325 billion[2]. Complications associated with diabetes include neuropathy,

retinopathy, nephropathy, cardiovascular disease[3], poor wound healing[4], and delayed

fracture healing[5].

Over time, and particularly if uncontrolled, diabetes can decrease adolescent

linear bone growth[6], reduce bone mineral density (BMD)[7], result in poor osseous

healing[8], and increase fracture risk[9]. Type 1 diabetes is associated with decreased

osteoblast recruitment and activity. Studies have shown bone resorption to be unaltered

or decreased in experimental diabetes. Loss of collagen can be measured within 2 weeks

of diabetes induction in animals with type 1 diabetes[10], and serum alkaline phosphatase

and osteocalcin levels are lower in T1D animals than in control animals. Studies have

shown decreased trabecular bone in the tibia in both streptozotocin (STZ)-induced

diabetic mice and nonobese spontaneously diabetic mice[11].

Osteopenia accompanies the onset of type 1 diabetes[12]. Levin et al. revealed

decreased skeletal mass in 50% of type 1 and type 2 diabetes patients[13]. Guo et al.[14]

examined STZ-induced diabetic Sprague Dawley rats after 6 months of hyperglycemia and

reported that saline-treated diabetic rats had a lower relative bone volume, trabecular

number, and trabecular thickness in comparison to normoglycemic controls. Load-to-

failure, energy absorption, and stiffness were also significantly decreased in diabetic

vehicle-treated rats. In some reports insulin therapy for 8 weeks was found to be

109

ineffective at reversing defects in diabetic bone[15]. STZ-induced diabetic Wistar-Albino

rats treated with insulin for 8 weeks had significant decreases in bone mineral density,

load, and energy relative to normal rats suggesting that insulin alone cannot restore the

biomechanical properties of bone in STZ-induced T1D rats.

Naltrexone (NTX), an opioid receptor antagonist, blocks the interaction of OGF

with OGFr, and continuous blockade of the receptor leads to increased cell

proliferation[16]. Preclinical studies in our labs investigating NTX modulation of the OGF-

OGFr regulatory pathway have documented that both systemic and topical NTX enhanced

corneal epithelial wound healing in rats[17,18] without adverse effects [19]. NTX

treatment also enhanced full-thickness cutaneous wound healing in diabetic rats by

accelerating wound closure, increasing angiogenesis, and collagen production [20,21].

Based on these observations, it was hypothesized that type 1 diabetes is

associated with a dysregulated OGF-OGFr axis and that modulation of this axis by high

dose NTX may alter bone composition. This study examines the effects of a continuous

blockade of the OGF-OGFr pathway by NTX on bone composition.

4.3. MATERIALS AND METHODS

4.3.1. Animals, induction of diabetes, and treatment

Twenty 6-week-old male Sprague Dawley rats (Charles River Laboratories,

Wilmington, MA) were housed under controlled environmental conditions with food and

water available ad libitum. All protocols were approved by Penn State College of

Medicine Institutional Animal and Use Committee. Type 1 diabetes was induced in 16

rats by intraperitoneal (i.p.) injection of streptozotocin (40 mg/kg; Sigma) on two

110

consecutive days; control rats (n=4) received sodium citrate buffer only [19,21]. Serum

glucose measurements were obtained using a TRUETrack® glucometer (Nipro

Diagnostics). Animals with serum glucose levels of at least 250 mg/dL were considered

hyperglycemic. Hyperglycemic rats were injected i.p. daily for 3 weeks with either 30

mg/kg naltrexone (NTX; 8 animals) or phosphate buffered saline (PBS; 8 animals); normal

animals received no treatment.

4.3.2. Three-point bending

A 225N load cell was used in conjunction with the ElectroForce LM1 Testbench®

system (Bose/TA Instruments, New Castle, DE) with standard three-point bend fixture.

The two supports were spread 1.5 cm apart, and femurs were oriented for an anterior to

posterior bend to failure. Maximum force (failure point), stiffness, and energy-to-failure

were recorded.

4.3.3. MicroCT

High-resolution images of the femur were acquired with a μCT imaging system

(vivaCT 40, Scanco, Brüttisellen, Switzerland) and analyzed based on our previously

described protocol[22]. Briefly, soft tissue was removed and the femurs were stored in

PBS until time of scan. Femurs were scanned with a voxel size of 10.5 μm. Analysis

regions were selected proximal to the distal femur growth plate (trabecular) and at the

midshaft (cortical). Standard parameters[23] calculated include bone volume (BV), total

volume (TV), relative bone volume (BV/TV), trabecular number, trabecular thickness,

trabecular separation, bone mineral density (BMD), tissue mineral density (TMD), cross-

sectional area, cortical area, cortical area fraction, cortical thickenss, and cortical tissue

111

mineral density (BMD) were calculated using the Scanco software at the distal and mid-

shaft scanning regions. Seventy-five slices were used for trabecular calculations proximal

to the distal growth plate and 20 slices were used for cortical calculations at the

midshaft[22].

4.3.4. Immunohistochemistry

Bones were fixed in 10% neutral buffered formalin for 3 days and allowed to

decalcify in 14% ethylenediaminetetra-acetic acid for 2-3 weeks. The bones were

dehydrated in ethanol and xylene, embedded in paraffin, and sectioned at 5 µm. Sections

were rehydrated and underwent antigen retrieval with pepsin for 10 minutes. Following

washes, tissue was blocked using bovine serum albumin for 1 hour. Primary antibodies

requiring overnight incubation at 4°C included 1:200 OGF, 1:200 OGFr (Bethyl

Laboratories, Montgomery, TX), and 1:500 osteocalcin (Santa Cruz Biotechnology, Inc,

Dallas, TX). Ki67 (1:300; Merck Millipore, Burlington, MA) and VEGF (1:100; Neo Markers

ThermoFisher Scientific, Waltham, MA) were incubated for half an hour at room

temperature. Anti-OGF was generated in our laboratory[24]. Following washes, sections

were incubated with goat anti-rabbit secondary antibody (Alexa Fluor® 568, A11011,

ThermoFisher) for 2 hours, washed and mounted with 50:50 glycerol:PBS with DAPI

(1:2000). Double labeling of bone tissues for OGFr and osteocalcin were performed by

incubation with both primary antibodies overnight at 4°C followed by washes and

incubation with Alexa Fluor® 488 goat anti-mouse (1:1000) and Alexa Fluor® 568 goat

anti-rabbit (1:1000) for 2 hours. An Olympus IX81 and Spot RT3 camera were used to

112

obtain fluorescent images. Intensity of TRITC staining was obtained semiquantitatively

using Slidebook software.

4.3.5. Histomorphometry

Bones were processed as described above for paraffin embedding, sectioned, and

stained with fast green and safranin O[26] to differentiate between bone (green) and

cartilage (red). Adjacent sections were stained using hematoxylin and eosin (H & E).

Cartilage and bone were assessed in femora using photographs taken on an Olympus

BX51 with a Spot RTKE camera. Analysis was performed using ImageJ[26].

4.3.6. Data Analyses

Data were analyzed using GraphPad Prism 7.0 software (Graph Pad) with one-way

analysis of variance (ANOVA) followed by Newman-Keuls post-hoc test. Student’s two-

tailed unpaired t-test was used where indicated; significance was determined at α-

value of 0.05.

4.4. RESULTS

4.4.1. Three-point bending

The force necessary to break normal bones averaged 95.4N (figure 1). There was

a 26% decrease in strength of diabetic vehicle-treated bones compared to normal bones,

whereas NTX increased the strength of diabetic bones by 21%. The energy-to-failure of

normal bones averaged 52.4±5.7Nmm. Diabetic vehicle-treated bones required 37% less

energy to break. In our samples, naltrexone-treated diabetic bones had a trend of being

113

37% stiffer than those in diabetic rats receiving vehicle, but this did not reach significance

(p=0.064).

Figure 4.1: Diabetes altered the responses to 3-point bending to failure. Bars (means ±

SEM) represent the force (A) necessary to break bones by 3-point bending, the energy (B)

absorbed by the bones prior to breaking, and the stiffness (C) of the bones. Significantly

different from N at * p<0.05, ** p<0.01, and between diabetes treatment groups at +

p<0.05. N = normal rats; DB/V = diabetic rat receiving saline; DB/NTX = diabetic rats

receiving 30mg/kg NTX for 21 days. At least 3 specimens were tested per group

4.4.2. MicroCT

There were no significant differences between the 3 groups in trabecular μCT

measurements (table 1). Relative to normal rats, cortical μCT measurements (table 2)

114

revealed significant decreases of 21% and 18% in cross-sectional area (inside periosteal

envelope) of vehicle-treated rats with diabetes and NTX-treated diabetic rats,

respectively, as well as significant decreases of 22% and 16% in cortical area of diabetic

vehicle-treated rats and NTX-treated diabetic rats, respectively.

TV (mm3) BV (mm3) BV/TV (%)

trabecular number (1/mm)

trabecular thickness (mm)

trabecular separation (mm)

BMD (mgHA/ cm3)

TMD (mgHA/ cm3)

N 11.01±

0.58 3.06±

0.46 31.07±

0.04 7.15±

0.64 0.071±

0.002 0.16±

0.01 287±

22 753±

6

DB/V 9.95±

0.40 2.06±

0.84 29.50±

0.07 6.26±

1.12 0.067±

0.004 0.20±

0.03 273±

47 774±

12

DB/NTX 10.38±

0.53 4.63±

0.78 44.45±

0.07 8.38±

1.45 0.070±

0.002 0.15±

0.04 368±

45 764±

8 Table 4.1: Mean ± SEM trabecular measurements from μCT evaluation. N = normal rats;

DB/V = diabetic rats receiving saline; DB/NTX = diabetic rats receiving 30mg/kg NTX for

21 days. TV = total volume; BV = bone volume; BV/TV = relative bone volume; BMD =

bone mineral density; TMD = tissue mineral density. n ≥ 5 per group.

total cross-sectional area (mm)

cortical area (mm2)

cortical area fraction (%)

cortical thickness (mm)

TMD (mgHA/cm3)

N 2.12±0.05 1.34±0.04 63.45±0.02 0.50±0.02 1099±7.3DB/V 1.68±0.06a 1.05±0.05b 62.61±0.02 0.49±0.02 1112±4.3DB/NTX 1.74±0.06a 1.13±0.04b 64.68±0.01 0.51±0.03 1111±8.7

Table 4.2: Mean ± SEM cortical measurements from μCT evaluation. Significantly

different from N/V at ap<.001 and bp<.01. N = normal rats; DB/V = diabetic rats receiving

saline; DB/NTX = diabetic rats receiving 30mg/kg NTX for 21 days. TMD = tissue mineral

density. n ≥ 4 per group.

115

4.4.3. Immunohistochemistry

There was a significant increase (126%) in OGFr expression (figures 2A and 2B) in

diabetic femurs compared to that recorded in normal bone that had a mean TRITC

intensity of 84.8 units. Naltrexone-treated diabetic rats had a reduced amount (57%) of

OGFr immunostaining in comparison to diabetic vehicle-treated rats. In normal femur

tissue 31.5% of cells stained positive for osteocalcin (figures 3A and 3B), an osteoblast

marker, whereas only 26% of the cells in the femurs of diabetic rats receiving vehicle.

Although not reaching significance, NTX-treated diabetic rat bones had increased (32%)

osteocalcin staining compared to vehicle-treated bone.

Tissue sections of femora from normal rats had 48% of cells stained positive for

Ki67 (figure 4), a proliferation marker. Relative to normal values, there was a 33%

decrease in Ki67 staining in diabetic vehicle-treated rat bones and a 31% decrease in

diabetic NTX-treated bones.

116

Figure 4.2: OGF and OGFr immunohistochemistry. OGF immunohistochemistry (A) and

histogram representing staining intensity of OGF in an individual cell (B); OGFr

immunohistochemistry (C) and histogram representing staining intensity of OGFr in an

individual cell (D); insets show secondary antibody only staining. Scale bar = 10μm.

Values represent means ± SEM for at least 3 specimens per group; significantly different

from N at *** p<0.001 and between different treatment groups of diabetic rats at ++

p<0.01. N = normal rats; DB/V = diabetic rats receiving saline; DB/NTX = diabetic rats

receiving 30mg/kg NTX for 21 days.

117

Figure 4.3: The presence of OGFr and osteocalcin in bone tissue from normal and diabetic

rats. Immunohistochemistry double-labeling of osteocalcin and OGFr (A) and histogram

representing the percentage of osteocalcin-positive cells within a field of view (B). Scale

bar = 10μm. Values represent means ± SEM; significantly different from N at * p<0.05

(Student’s t-test). N = normal rats; DB/V = diabetic rats receiving saline; DB/NTX = diabetic

rats receiving 30mg/kg NTX for 21 days. n ≥ 7 measurements per group.

118

Figure 4.4: Ki67 staining. Histogram representing the percentage of Ki67-positive cells in

a field of view. Values represent means ± SEM for at least 3 fields from at least 3 bones

per group. Significantly different from N at * p<0.05, ** p<0.01. N = normal rats; DB/V =

diabetic rats receiving saline; DB/NTX = diabetic rats receiving 30mg/kg NTX for 21 days.

4.4.4 Histomorphometry

Analyses of safranin O stained sections revealed no significant differences in the

percentage of cartilage and bone between the 3 groups, and hematoxylin-eosin stained

tissues showed no difference in the morphology, suggesting that systemic NTX treatment

did not result in any gross pathology of the bone tissue.

4.5. DISCUSSION

Diabetes affects more than 30 million Americans[1] and of those with type 1

diabetes, 50-60% have osteopenia[27,28] and 14-20% have osteoporosis[28,29]. The

mechanism connecting poor bone quality to diabetes is still unclear[30]. The results of

this study support current literature that there is a decrease in osteoblast

119

activity/maturity. There was a decrease in osteocalcin staining in vehicle-treated diabetic

rat bones compared to normal bones, suggesting a decrease in the number of mature

osteoblasts; however, systemic NTX treatment appeared to be reestablishing the

osteoblast population. Histomorphological studies using the proliferative marker Ki67

showed decreased staining in diabetic bones from animals treated with either saline or

NTX compared to normal bones. The lack of change in Ki67 may be due to the relative

lack of dynamics occurring in an intact bone. Important to the work on modulation of the

OGF-OGFr axis in bone fracture repair, OGFr staining was significantly increased in

vehicle-treated diabetic bone, whereas systemic treatment with NTX reduced the number

of receptors in diabetic femurs to that of normal bone.

Studies involving STZ-induced diabetic mice showed significant decreases in tibia,

femur, and vertebrae trabecular bone volume fraction[11,31]. Although we did not see

any significant change in trabecular bone composition it may be that the rats did not have

diabetes long enough for these changes to become significant, and a longer period of

hyperglycemia may be required to see changes in bone composition. Hamada et al. did

not see significant differences in bone mineral density between STZ-induced diabetes and

normal mice at 4 weeks post STZ injection[32]. It is also predicted that longer periods of

NTX treatment, as well as beginning NTX treatment 4-5 weeks after rats become

hyperglycemic may be important for ascertaining the role of systemic NTX on bone

composition. Guo et al.[14] and Erdal et al.[15] observed decreases in bone quality at 6

months and 8 weeks of hyperglycemia, respectively.

120

4.6. CONCLUSION

In conclusion systemic NTX treatment for 21 days in hyperglycemic rats reversed

certain important deficiencies in the femur. Bone tissue in diabetes rats receiving saline

had more OGFr present suggesting the greater availability of receptors to interact with

the inhibitory peptide OGF and thus reduce cell proliferation. Ki67 staining was decreased

in diabetic vehicle-treated bones compared to normal bones indicating a decrease in

proliferative cells. Importantly, diabetic bone required less force to break under 3-point

bending, a parameter which was rescued by NTX, suggesting that blockade of the OGF-

OGFr axis may increase bone strength in diabetes.

4.7. Acknowledgements

Special thanks to Hwa Bok Wee for assistance with μCT scanning.

4.8. Author contributions

Study design: MBT, PJM; Study conduct: MBT, GSL; Data collection and analyses: MBT,

PJM

Data interpretation: MBT, PJM, GSL; Drafting, editing and approving final manuscript:

MBT, GSL, ISZ, PJM

4.9. Funding

Research was supported in part by discretionary gift funds to ISZ and PJM.

4.10. Competing interests - None.

121

4.11. References

[1] National Diabetes Statistics Report, 2017; Centers for Disease Control and

Prevention: Atlanta, GA, 2017.

[2] Yang, W.; Dall, T.M.; Beronjia, K.; Lin, J.; Semilla, A.P.; Chakrabarti, R.; Hogan, P.F.

Economic Costs of Diabetes in the U.S. in 2017; Diabetes Care; American Diabetes

Association, 2018.

[3] Nathan, D.M. Long-Term Complications of Diabetes Mellitus. N. Engl. J. Med.,

1993, 328, 1676–1685.

[4] Falanga, V. Wound Healing and Its Impairment in the Diabetic Foot. The Lancet,

2005, 366, 1736–1743.

[5] Hofbauer, L.C.; Brueck, C.C.; Singh, S.K.; Dobnig, H. Osteoporosis in Patients With

Diabetes Mellitus. J. Bone Miner. Res., 2007, 22, 1317–1328.

[6] Salerno, M.; Argenziano, A.; DiMaio, S.; Gasparini, N.; Formicola, S.; DeFilippo, G.;

Tenore, A. Pubertal Growth, Sexual Maturation, and Final Height in Children With

IDDM: Effects of Age at Onset and Metabolic Control. Diabetes Care, 1997, 20,

721–724.

[7] Vestergaard, P. Discrepancies in Bone Mineral Density and Fracture Risk in

Patients with Type 1 and Type 2 Diabetes—a Meta-Analysis. Osteoporos. Int.,

2007, 18, 427–444.

[8] Cozen, L. Does Diabetes Delay Fracture Healing? Clin. Orthop., 1972, 82, 134–140.

122

[9] Janghorbani, M.; Van Dam, R.M.; Willett, W.C.; Hu, F.B. Systematic Review of

Type 1 and Type 2 Diabetes Mellitus and Risk of Fracture. Am. J. Epidemiol., 2007,

166, 495–505.

[10] Motyl, K.; McCabe, L.R. Streptozotocin, Type I Diabetes Severity and Bone. Biol.

Proced. Online, 2009, 11, 296–315.

[11] Botolin, S.; McCabe, L.R. Bone Loss and Increased Bone Adiposity in Spontaneous

and Pharmacologically Induced Diabetic Mice. Endocrinology, 2007, 148, 198–

205.

[12] McNair, P.; Madsbad, S.; Christiansen, C.; Faber, O.K.; Transbøl, I.; Binder, C.

Osteopenia in Insulin Treated Diabetes Mellitus: Its Relation to Age at Onset, Sex

and Duration of Disease. Diabetologia, 1978, 15, 87–90.

[13] Levin, M.E.; Boisseau, V.C.; Avioli, L.V. Effects of Diabetes Mellitus on Bone Mass

in Juvenile and Adult-Onset Diabetes. N. Engl. J. Med., 1976, 294, 241–245.

[14] Guo, J.; Dong, W.; Jin, L.; Wang, P.; Hou, Z.; Zhang, Y. Hydrogen-Rich Saline

Prevents Bone Loss in Diabetic Rats Induced by Streptozotocin. Int. Orthop., 2017,

41, 2119–2128.

[15] Erdal, N.; Gürgül, S.; Demirel, C.; Yildiz, A. The Effect of Insulin Therapy on

Biomechanical Deterioration of Bone in Streptozotocin (STZ)-Induced Type 1

Diabetes Mellitus in Rats. Diabetes Res. Clin. Pract., 2012, 97, 461–467.

[16] McLaughlin, P.J.; Zagon, I.S. Duration of Opioid Receptor Blockade Determines

Biotherapeutic Response. Biochem. Pharmacol., 2015, 97, 236–246.

123

[17] Zagon, I.S.; Sassani, J.W.; McLaughlin, P.J. Re-Epithelialization of the Rat Cornea Is

Accelerated by Blockade of Opioid Receptors. Brain Res., 1998, 798, 254–260.

[18] Zagon, I.; Sassani, J.; McLaughlin, P. Cellular Dynamics of Corneal Wound Re-

Epithelialization in the Rat - II. DNA Synthesis of the Ocular Surface Epithelium

Following Wounding. BRAIN Res., 1999, 839, 243–252.

[19] Klocek, M.S.; Sassani, J.W.; McLaughlin, P.J.; Zagon, I.S. Topically Applied

Naltrexone Restores Corneal Reepithelialization in Diabetic Rats. J. Ocul.

Pharmacol. Ther., 2007, 23, 89–102.

[20] McLaughlin, P.J.; Pothering, C.A.; Immonen, J.A.; Zagon, I.S. Topical Treatment

with the Opioid Antagonist Naltrexone Facilitates Closure of Full-Thickness

Wounds in Diabetic Rats. Exp. Biol. Med., 2011, 236, 1122–1132.

[21] McLaughlin, P.J.; Cain, J.D.; Titunick, M.B.; Sassani, J.W.; Zagon, I.S. Topical

Naltrexone Is a Safe and Effective Alternative to Standard Treatment of Diabetic

Wounds. Adv. Wound Care, 2017, 6, 279–288.

[22] Lloyd, S.A.; Lang, C.H.; Zhang, Y.; Paul, E.M.; Laufenberg, L.J.; Lewis, G.S.;

Donahue, H.J. Interdependence of Muscle Atrophy and Bone Loss Induced by

Mechanical Unloading. J. Bone Miner. Res., 2014, 29, 1118–1130.

[23] Bouxsein, M.; Boyd, S.; Christiansen, B.; Guldberg, R.; Jepsen, K.; Muller, R.

Guidelines for Assessment of Bone Microstructure in Rodents Using Micro-

Computed Tomography. J. BONE Miner. Res., 2010, 25, 1468–1486.

124

[24] Zagon, I.S.; McLaughlin, P.J. Production and Characterization of Polyclonal and

Monoclonal Antibodies to the Zeta (ξ) Opioid Receptor. Brain Res., 1993, 630,

295–302.

[25] Schmitz, N.; Laverty, S.; Kraus, V.B.; Aigner, T. Basic Methods in Histopathology of

Joint Tissues. Osteoarthritis Cartilage, 2010, 18, S113–S116.

[26] Schneider, C.A.; Rasband, W.S.; Eliceiri, K.W. NIH Image to ImageJ: 25 Years of

Image Analysis. Nat. Methods, 2012, 9, 671–675.

[27] Kayath, M.J.; Tavares, E.F.; Dib, S.A.; Vieira, J.G.H. Prospective Bone Mineral

Density Evaluation in Patients With Insulin-Dependent Diabetes Mellitus. J.

Diabetes Complications, 1998, 12, 133–139.

[28] Kemink, S.A.G.; Hermus, A.R.M.M.; Swinkels, L.M.J.W.; Lutterman, J.A.; Smals,

A.G.H. Osteopenia in Insulin-Dependent Diabetes Mellitus; Prevalence and

Aspects of Pathophysiology. J. Endocrinol. Invest., 2000, 23, 295–303.

[29] MunozTorres, M.; Jodar, E.; EscobarJimenez, F.; LopezIbarra, P.; Luna, J. Bone

Mineral Density Measured by Dual X-Ray Absorptiometry in Spanish Patients with

Insulin-Dependent Diabetes Mellitus. Calcif. TISSUE Int., 1996, 58, 316–319.

[30] Räkel, A.; Sheehy, O.; Rahme, E.; LeLorier, J. Osteoporosis among Patients with

Type 1 and Type 2 Diabetes. Diabetes Metab., 2008, 34, 193–205.

[31] Martin, L.M.; McCabe, L.R. Type I Diabetic Bone Phenotype Is Location but Not

Gender Dependent. Histochem. Cell Biol., 2007, 128, 125–133.

125

[32] Hamada, Y.; Kitazawa, S.; Kitazawa, R.; Fujii, H.; Kasuga, M.; Fukagawa, M.

Histomorphometric Analysis of Diabetic Osteopenia in Streptozotocin-Induced

Diabetic Mice: A Possible Role of Oxidative Stress. Bone, 2006, 40, 1408–1414.

126

CHAPTER 5: DISCUSSION

127

5.1. Discussion

Over 30 million Americans have diabetes, 5-10% of that population have type 1

diabetes[1,2]. Diabetes costs the U.S. over $300 billion each year[1] including the cost of

complications associated with diabetes such as non-healing foot ulcers. Diabetic foot

ulcers (DFU) are also associated with peripheral neuropathy. DFUs are often undetected

by individuals due to sensory neuropathy in addition to the fact that the initial wound is

small. Eighty-five percent of amputations in diabetics were preceded by a non-healing

ulcer[3]. High incidences of diabetic foot ulceration is not unique to North America,

although it does have the highest prevalence at 13%[3]. Non-healing foot ulcers affect

6.3% of the world’s population[3]. There needs to be research on treating not just the

complications on a superficial level but at a mechanistic level as well.

Type 1 diabetic (T1D) bone has a lower bone mineral density (BMD) at the neck of

the femur than normal bone [4]. However, the same difference is not observed at in the

lumbar spine. This slight decrease in BMD is not great enough to account for the highly

increased fracture risk among T1D patients. Microvascular complications such as

retinopathy and neuropathy, coincide with low BMD. Trabecular bone score (TBS) is an

index used to evaluate pixel variation in lumbar spine DXA images and correlates with the

trabecular bone volume to tissue ratio, trabecular number, and connectivity. Osteocalcin,

an osteoblast marker, and CTX, an osteoclast marker, were both lower in diabetic bone

than normal bone. No differences were discerned among calcium, phosphorus, and

parathyroid hormone levels[4]. A study examining trabecular bone indicated no

differences in bone formation or resorption parameters between the two groups[5].

128

Computed tomography suggested lower cortical thickness and cross-sectional area of the

femur in T1D patients. There was also a larger endosteal circumference amongst the T1D

patients. Studies suggest that type 1 diabetes affects cortical bone to a greater extent

than trabecular bone[4].

Although this dissertation is focused on type 1 diabetes, it is important to briefly

note the relationship of type 2 diabetes and bone. Type 1 diabetes has lower insulin-like

growth factor-1 and higher insulin-like growth factor binding protein-1 serum levels than

type 2 diabetes and normal controls[6]. Levels of 25OHD are significantly lower in both

forms of diabetes[7]. Circulating levels of sclerostin, a protein opposing bone building,

are greater in type 2 diabetes than in either type 1 diabetes or control. Osteocalcin and

amino-terminal propeptide of pro-collagen type 1 are lower in type 2 diabetes compared

to normal controls[8]. Hyperglycemia also increases the levels of anti-osteoblastic

cytokines. Studies indicate that osteoblast precursor cells are decreased in type 2

diabetes while osteoclast precursor cells are increased. Both osteoblast and osteoclast

expression is reduced in type 2 diabetes. Type 2 diabetes medications known as

thiazolidinediones which include rosiglitazone may put patients at a higher risk of

fracture. The drug promotes peroxisome proliferator-activated receptor gamma (PPARγ)

which favors mesenchymal differentiation to adipose rather than osteoblasts[9].

In vivo and in vitro studies of rat and rabbit corneas demonstrated that modulation

of the OGF-OGFr regulatory pathway with naltrexone (NTX) effectively enhanced wound

healing and reepithelialization[10]. Both systemic and topical naltrexone can enhance

129

reepithelialization of the normal rat cornea[11–14]. Topical NTX can also enhance

reepithelialization of full-thickness cutaneous wound healing[15–19] in diabetic rats.

Topical NTX is safe to use on the cornea[20]. Sprague Dawley rats received eye

drops of NTX at concentrations of 10-3, 10-4, 10-5, 10-6, or 10-7 M 4 times daily for 7 days.

No gross or histological pathology was detected. Topical NTX is also well tolerated by

humans [21]. Volunteers administered NTX eye drops of concentrations 1x10-6 M (1 drop

or 4 drops daily), 5x10-6 M (4 drops daily), 1x10-5 M (4 drops daily), 5x10-5 M (4 drops daily)

for 7 days and reported no adverse effects.

Moreover, NTX has been shown to be a safe and effective treatment for full-

thickness wound healing[19]. Specific Aim 1 showed that wounds treated with NTX

healed faster than vehicle-treated wounds surgically created on the dorsum of type 1

diabetic rats. The rate of wound closure was non-inferior to Regranex®-treated wounds,

demonstrating comparable efficacy with the standard of care. Mechanistically, NTX

increased BrdU labeling, indicative of cell proliferation, in the wound matrix, in

comparison to vehicle-treated wounds. NTX also increased expression of VEGF, required

for angiogenesis – necessary for wound healing – relative to both vehicle-treated and

Regranex®-treated wounds. The increased vascularization in NTX-treated wounds is an

important component of skin repair, as diabetes is often associated with poor vascularity

which can inhibit wound healing. A 1.5-fold increase in platelet derived growth factor

(PDGF) staining was seen at day 1 in both NTX-treated and Regranex®-treated wounds

compared to vehicle-treated wounds. The active ingredient in Regranex® is PDGF so it

may be expected that tissue treated with Regranex® would show an increase in PDGF

130

expression. However, by day 2 the tissue levels of PDGF were comparable between

Regranex®-treated and vehicle-treated wounds. NTX-treated wounds at day 2 still

showed an increase in PDGF expression compared to vehicle-treated wounds.

The Regranex® study supports previous studies on corneal defect repair and full-

thickness cutaneous healing, indicating that naltrexone is an effective treatment for

diabetic healing. Naltrexone was also shown to be as effective as the standard of care,

Regranex®[19]. Benefits of using naltrexone include the safety of naltrexone as evidenced

by previous studies in the cornea and skin of rats and rabbits, as well as the decreased

cost of naltrexone. Regranex® comes with a black box warning stating that there is a

higher risk of mortality secondary to malignancy when using more than three tubes of the

gel[22]. The price differential would bring down the cost of diabetic foot and perhaps

make patients more compliant with the treatment.

The Specific Aim 1 study on full-thickness cutaneous wounds [19] lends

justification for a clinical trial to begin. There have been multiple studies performed on

both rats[15,16,19] and mice[23] indicating naltrexone is an effective treatment for full-

thickness wound healing. Safety studies on topical naltrexone have been performed on

the cornea[11] of both rats and rabbits (unpublished) indicating the safety of naltrexone.

A clinical study would be warranted for testing NTX as a topical therapy for non-healing

foot ulcers.

Specific Aim 2 established the presence of the OGF-OGFr axis in bone and the

potential for dysregulation of the pathway in diabetes. While OGF and OGFr have been

identified in bone throughout rat development and into adulthood, there is an increase

131

in OGFr at days 1 and 21. The increase at day 21 may be biologically significant as this is

the point at which pups ween. There may be other demands that take priority at this

stage over bone growth causing an increase in OGFr and a decrease in bone proliferation.

There is a 95% increase in the inhibitory peptide OGF in diabetic rat bone relative to OGF

levels in normal bone. Diabetic bone showed an 84% increase in OGFr staining compared

to normal bone. Serum levels of OGF indicated a 23% increase in diabetic rats over normal

controls. Thus it appears that the OGF-OGFr regulatory pathway is dysregulated in

diabetes leading to elevated peptide and receptor interactions that may contribute to

bone loss and delayed bone healing.

The rat studies documented a 66% decrease in Ki67 staining in type 1 diabetic rat

bone compared to normal bone. Ki67 is a nuclear protein expressed during the active

phases of the cell cycle[24] thus indicating levels of cell proliferation. Vascularization was

also decreased in diabetic bone compared to normal controls, tissue with a difference of

30% in VEGF staining. During fracture repair the callus of vehicle-treated diabetic rats

retained more granulation tissue at day 7 than either diabetic naltrexone-treated and

normal controls. Diabetic vehicle-treated fractures had less cartilage within the callus

compared to normal fractures. There was also a decrease in bone seen in the callus of

the vehicle-treated diabetic fractures compared to those treated with naltrexone. When

examining the region closest to the periosteum there was 84% more granulation tissue in

diabetic vehicle-treated fracture calluses compared to normal control and 52% more than

diabetic naltrexone-treated fractures. There was 88% more cartilage in diabetic

naltrexone-treated fractures than in diabetic vehicle-treated fractures. Diabetic calluses

132

had 24% less bone than normal calluses. Diabetic naltrexone-treated fractures had 35%

more bone than diabetic vehicle-treated calluses. Proliferation was also altered by

naltrexone with 35% more Ki67 staining compared to vehicle-treated diabetic fractures.

The present study corroborates that of Zagon, Wu, and McLaughlin who examined

fetal rat tissue from pregnant dams injected with NTX or sterile water [25]. Indexes of

radiolabeled thymidine were higher in fetuses from NTX-treated dams in comparison to

saline-treated controls, suggesting that prenatal NTX increased proliferation in bone; OGF

treatment was documented to decrease proliferation. Immunocytochemistry indicated

the presence of OGF and OGFr in both vertebrae and ribs of normal rat offspring. Further

evidence of the OGF-OGFr pathway reported in tissue culture studies where NTX and OGF

modulated cellular proliferation in the HT-1080 and SK-ES-1 sarcoma cell lines[26].

Data from the second aim supports the work of Liskov et al.[27], Thakur,

DeBoyace, and Margulies[28], and Petrizzi et al.[29], indicating a role for an opioid

antagonist in osteogenesis. Liskov et al. found a 2-fold increase in the thickness of the

perichondral bone cuff as well as over 6.5 times the number of dividing cells in the growth

plate of femurs. This paper neglected to look at OGFr and utilized a non-mammalian

model[27]. Thakur, DeBoyace, and Margulies used a mouse model to examine cortical

defects in tibiae. Again, naloxone was used. As in our fracture project, animals were

euthanized 7 days post-fracture. There was a 1.5-fold increase in relative bone volume

of naloxone-treated defects compared to control as well as an increase in trabecular

number. Naloxone was able to decrease the defect diameter by 20%. OGF and OGFr

were only studied in the cell culture experiments not in the in vivo experiments[28].

133

Petrizzi et al. augmented naloxone treatment with calcium gluconate allowing for 4 weeks

of treatment of a metacarpal defect in sheep. There were more remodeling spaces and

fewer osteoblasts in the area of the defect, indicating more advanced stages of healing.

The authors did not look into the role of OGF and OGFr in the study[29].

In the fracture study there was significantly more bone and cartilage in diabetic

fracture calluses treated with NTX compared to those treated with vehicle only. At this

time point, 7 days post-fracture, there should still be a significant amount of cartilage. If

we were to have examined the bones at a much later timepoint and there was still an

increase in cartilage, then there would be a delay in the healing process. In addition, the

increase in bone may be from a combination of endochondral ossification and

intramembranous ossification, thus having increased cartilage and bone is reasonable.

The fracture study was limited by the short survival period of the rats. Healing

normally required 4-6 weeks, and the studies should be repeated to evaluate rate of bone

repair and strength (mechanical testing) of the bone following topical NTX application. In

addition, different methods of NTX application should be evaluated. Including micelles

or beads coated with NTX. Coating the wire in a slow-releasing material would be

beneficial in scenarios where internal stabilization or pinning is necessary. Future

research should examine angiogenesis in fracture repair and utilize db/db type 2 diabetic

mice. Type 2 diabetic mice may show differences in preference for differentiation of

newly proliferated progenitor cells toward adipose instead of osteoblasts. This could

further complicate the experiment however determining if NTX affects differentiation

would be an interesting observation. Differentiation may be increased by NTX as the

134

entire process of proliferation is enhanced. Since cells are either dividing or

differentiating having a faster proliferation rate would allow cells to enter differentiation

sooner. The argument against this notion is that cells are stimulated by NTX to divide and

therefore must wait until division is finished to differentiate. This would decrease cell

differentiation overall.

Osteoclast numbers in diabetes has been reported to be increased, decreased, or

unchanged[30]. There was no significant difference in TRAP staining between diabetic

and normal bone (unpublished). It is unknown if hematopoietic cells react to NTX.

Macrophages replicate infrequently[31], and early osteoblast precursors may not

necessarily differentiate into osteoclasts. Therefore, NTX may not have as substantial an

effect on osteoclasts as it does osteoblasts. This would allow for the use of NTX to

increase osteoblastic cells without much of an increase in osteoclasts, resulting in greater

bone deposition.

Although bone loss is not prevented by maintenance of a healthy insulin level[32],

the Lin lab has used local delivery of insulin[33], vanadium[34], and manganese

chloride[35] to improve fracture healing in a rat model. Local insulin inhibits FOXO1

preventing Wnt inhibition. Vanadium[34] and manganese chloride[35] mimic insulin and

may also inhibit FOXO1. This helps to increase the number of osteoblasts. Naltrexone

also attempts to increase osteoblast numbers. Basic fibroblast growth factor normalized

diabetic fracture repair and enhanced healing in non-diabetic animals[36]. FGF was also

able to increase callus formation 3 weeks post-fracture[36]. Platelet-rich plasma high in

135

levels of mitogenic factors increased cell proliferation in diabetic rats and normalized

healing[37]. Naltrexone also has the ability to increase cell proliferation.

Kawaguchi et al.[36], also using Sprague Dawley rats, induced diabetes using STZ

and waited 2 weeks before commencing with fibula fractures. bFGF was introduced to

the fracture site within a fibrin gel. Rats were euthanized 1, 3, and 5 weeks after surgery.

Similar to our findings, Kawaguchi et al. saw intramembranous ossification under the

periosteum within the first week. The lab did not statistically compare the diabetic bone

values to those of non-diabetic rats although there appears to be a difference. bFGF

increased breaking strength and energy absorbed compared to diabetic vehicle-treated

rat bones. As seen in our study there was a decrease in breaking strength and energy

absorbed in diabetic vehicle-treated rats compared to non-diabetic rats.

Wang et al.[38] used STZ-induced diabetic Wistar rats and a closed femoral

fracture model. Fracture healing was evaluated 4 weeks post-fracture. At 4 weeks there

was no significant difference between total volume, bone volume, relative bone volume,

or trabecular thickness between diabetic and normal bones. There were significant

differences between trabecular number and trabecular separation. Load, stiffness, and

absorbed energy were decreased in diabetic bones compared to non-diabetic bones. This

is supported by results from Specific Aim 3.

Wang, Jiang, and Du[39] used rats to examine insulin therapy on Transforming

Growth Factor-Beta 1 (TGF-β1) expression in fracture calluses. Calluses were examined

1, 2, 4, 6, and 8 weeks post-fracture. As in our experiments the lab utilized an open

fracture model. Changes in TGFβ1 were present in the first week following fracture.

136

Biomechanical properties were tested at 6 and 8 weeks post-fracture and showed

significant decreases in diabetic bone.

Cignachi et al.[40] used C57/BL6 mice and induced diabetes using injections of STZ.

Fractures were formed 7 days after the end of diabetes induction, creating a uni-cortical

defect. The RANKL/RANK/OPG system was evaluated using immunohistochemistry 21

days after the surgery. Similar to our results, the lab saw a significant decrease in bone

regeneration in STZ-induced diabetic animals. There was no significant difference

between the two groups in TRAP staining, RANK, OPG, or RANKL expression.

Ko et al.[41] used CD-1 mice to evaluate the mesenchymal stem cell population in

diabetic fracture healing. Diabetes was induced using STZ then mice underwent surgery

3 weeks later. Diabetic mice had 53% less new bone in fracture calluses compared with

non-diabetic mice 16 days after surgery.

Park et al.[42] used BB Wistar rats to examine femoral fracture healing treated

with insulin. Rats were euthanized 7 and 14 days post-fracture showing little change in

fracture repair. Mechanical testing at 4 weeks post-fracture showed an increase in

maximum torque to failure.

Yee et al.[43] utilized C57BL6/J mice injected with STZ as a diabetic model. Two

weeks after STZ injection closed fractures were generated and rats were euthanized 21

and 42 days later. At 21 days there was a decrease in bone volume and total volume, and

at 42 days there was a decrease in cortical bone mineral density.

Hreha et al.[35] used BB Wistar rats with a closed femoral fracture model.

Manganese chloride or saline were injected into the intramedullary canals. Femora were

137

collected at 7, 10, and 14 days post-fracture. Changes in maximum torque were seen at

4 weeks post-fracture. No differences in callus composition were seen at 7 days post-

fracture which resembles our findings. There was an increase in percent new mineralized

tissue in manganese chloride-treated bones at 10 days compared to saline controls. VEGF

and PECAM changes were not seen until day 10.

Specific Aim 3 addressed the ability to modulate the OGF-OGFr axis to prevent bone

loss associated with diabetes. The force necessary to break diabetic bones was significantly

less compared to normal bones. Naltrexone increased the strength of diabetic bones by

21% relative to vehicle-treated diabetic bones. Diabetic vehicle-treated bones absorbed

79% less energy prior to breaking than non-diabetic bones. Naltrexone increased the

bone’s integrity, requiring 105% more energy to break compared to vehicle-treated bones.

Although not statistically significant, normal and diabetic NTX-treated bones were stiffer

than diabetic vehicle-treated bones. No significant differences were seen between the 3

groups in trabecular μCT measurements. There was a decrease in cross-section and cortical

area in diabetic bone compared to non-diabetic bone. There was a 126% increase in OGFr

staining of diabetic bone compared to normal bone. Naltrexone reduced OGFr levels in

diabetic bone by 57%. There was a 26% decrease in osteocalcin staining of diabetic vehicle-

treated bones compared to normal controls. Naltrexone increased staining by 32% over

vehicle-treated bone. Diabetic vehicle-treated bone had 33% less Ki67 staining compared

to normal bone. Naltrexone did not adversely affect the morphology of the tissue, as

indicated by H & E staining.

138

The preventative study indicates there is a potential benefit to systemic

naltrexone in regards to glucose regulation. Additional studies should be conducted to

examine this phenomenon. If the study is allowed to continue past 3 weeks there may

be a more significant difference between diabetic and normal femurs, making the effect

of naltrexone more substantial. Also using systemic NTX as a treatment for bone

complications instead of a preventative by waiting a month to begin the treatment would

be another route to consider. This may result in fewer changes in the cortical bone but

may show a greater discrepancy between trabecular measurements in diabetic bone

relative to normoglycemic controls since there would be a longer duration of diabetes

and more chance for bone turnover to be slowed down. Without a trauma to spur

proliferation and repair there may be little change in NTX-treated groups. In addition, it

would be interesting to use systemic NTX as a treatment in an osteoporosis animal model.

Insulin therapy has been evaluated as a treatment for osteopenia in diabetes.

Hough et al.[44] induced diabetes in Wistar-Lewis rats with STZ and began insulin therapy

5 days later. Treatment was continued until day 48 when the rats were euthanized. There

was an increase in percent relative osteoid volume in insulin-treated tibiae compared to

that of diabetic saline-treated rats.

Guo et al.[45] injected Sprague Dawley rats with STZ to induce diabetes. Animals

were treated with a hydrogen-rich saline lavage for 3 months and rats were collected 12

weeks after the last injection. Diabetic saline-treated rats had a lower relative bone

volume, trabecular number, and trabecular thickness compared to non-diabetic controls.

There was an increase in trabecular separation as well. Ultimate load, energy absorption,

139

and stiffness were significantly decreased in diabetic vehicle-treated animals compared

to non-diabetic rats.

Erdal et al.[46] used Wistar-Albino rats injected with STZ as a model for diabetes.

Insulin treatments extended for 8 weeks. There was a significant decrease in length, cross

sectional area, bone mineral density, load, and energy in diabetic saline-treated bones

compared to non-diabetic bones. The study concluded that insulin is unable to restore

biomechanical deterioration of bone in STZ-induced T1D.

Treatments may be developed to work at various phases of the healing process.

Increasing cell proliferation and increasing vascularity are two such targets the lab looks

to utilize. Another phase prime for exploitation is the inflammatory phase. Much of the

delay in both full-thickness cutaneous wound healing and fracture repair in diabetes

relates to a prolonged inflammatory phase. There is over activation of the immune

system and this inflammatory process cannot be easily reduced to allow the subsequent

phases to begin.

In summary, NTX is an effective and safe treatment for full-thickness cutaneous

wounds and bone fractures. There appears to be merit to further exploration of

naltrexone’s protective effect in bone. The experiments supported work by various labs.

Further literature searches suggest an increase in treatment time for Specific Aims 2 and

3 may yield better results. Overall, NTX is a likely candidate as treatment for multiple

diabetic complications associated with skin and bone.

140

5.1. References

[1] The Cost of Diabetes http://www.diabetes.org/advocacy/news-events/cost-of-

diabetes.html (accessed Apr 29, 2018).

[2] National Diabetes Statistics Report, 2017; Centers for Disease Control and

Prevention: Atlanta, GA, 2017.

[3] Zhang, P.; Lu, J.; Jing, Y.; Tang, S.; Zhu, D.; Bi, Y. Global Epidemiology of Diabetic

Foot Ulceration: A Systematic Review and Meta-Analysis. Ann. Med., 2017, 49,

106–116.

[4] Shah, V.N.; Carpenter, R.D.; Ferguson, V.L.; Schwartz, A.V. Bone Health in Type 1

Diabetes. Curr. Opin. Endocrinol. Diabetes Obes., 2018, 25, 231–236.

[5] Armas, L.A.G.; Akhter, M.P.; Drincic, A.; Recker, R.R. Trabecular Bone

Histomorphometry in Humans with Type 1 Diabetes Mellitus. Bone, 2011, 50, 91–

96.

[6] Jehle, P.; Jehle, D.; Mohan, S.; Bohm, B. Serum Levels of Insulin-like Growth Factor

System Components and Relationship to Bone Metabolism in Type 1 and Type 2

Diabetes Mellitus Patients. J. Endocrinol., 1998, 159, 297–306.

[7] Gennari, L.; Merlotti, D.; Valenti, R.; Ceccarelli, E.; Ruvio, M.; Pietrini, M.G.;

Capodarca, C.; Franci, M.B.; Campagna, M.S.; Calabrò, A.; Cataldo, D.; Stolakis, K.;

Dotta, F.; Nuti, R. Circulating Sclerostin Levels and Bone Turnover in Type 1 and

Type 2 Diabetes. J. Clin. Endocrinol. Metab., 2012, 97, 1737–1744.

[8] Sassi, F.; Buondonno, I.; Luppi, C.; Spertino, E.; Stratta, E.; Di Stefano, M.;

Ravazzoli, M.; Isaia, G.; Trento, M.; Passera, P.; Porta, M.; Isaia, G.C.; D’Amelio, P.

141

Type 2 Diabetes Affects Bone Cells Precursors and Bone Turnover. BMC Endocr.

Disord., 2018, 18, 1–8.

[9] Stage, T.B.; Christensen, M.-M.H.; Jørgensen, N.R.; Beck-Nielsen, H.; Brøsen, K.;

Gram, J.; Frost, M. Effects of Metformin, Rosiglitazone and Insulin on Bone

Metabolism in Patients with Type 2 Diabetes. Bone, 2018, 112, 35–41.

[10] Zagon, I.S.; Sassani, J.W.; McLaughlin, P.J. Re-Epithelialization of the Rabbit

Cornea Is Regulated by Opioid Growth Factor. Brain Res., 1998, 803, 61–68.

[11] Zagon, I.S.; Sassani, J.W.; McLaughlin, P.J. Re-Epithelialization of the Rat Cornea Is

Accelerated by Blockade of Opioid Receptors. Brain Res., 1998, 798, 254–260.

[12] Zagon, I.S.; Jenkins, J.B.; Sassani, J.W.; Wylie, J.D.; Ruth, T.B.; Fry, J.L.; Lang, C.M.;

McLaughlin, P.J. Naltrexone, an Opioid Antagonist, Facilitates Reepithelialization

of the Cornea in Diabetic Rat. Diabetes, 2002, 51, 3055–3062.

[13] Klocek, M.S.; Sassani, J.W.; McLaughlin, P.J.; Zagon, I.S. Naltrexone and Insulin Are

Independently Effective but Not Additive in Accelerating Corneal Epithelial

Healing in Type I Diabetic Rats. Exp. Eye Res., 2009, 89, 686–692.

[14] Klocek, M.S.; Sassani, J.W.; McLaughlin, P.J.; Zagon, I.S. Topically Applied

Naltrexone Restores Corneal Reepithelialization in Diabetic Rats. J. Ocul.

Pharmacol. Ther., 2007, 23, 89–102.

[15] McLaughlin, P.J.; Pothering, C.A.; Immonen, J.A.; Zagon, I.S. Topical Treatment

with the Opioid Antagonist Naltrexone Facilitates Closure of Full-Thickness

Wounds in Diabetic Rats. Exp. Biol. Med., 2011, 236, 1122–1132.

142

[16] Immonen, J.A.; Zagon, I.S.; Lewis, G.S.; McLaughlin, P.J. Topical Treatment with

the Opioid Antagonist Naltrexone Accelerates the Remodeling Phase of Full-

Thickness Wound Healing in Type 1 Diabetic Rats. Exp. Biol. Med., 2013, 238,

1127–1135.

[17] McLaughlin, P.J.; Immonen, J.A.; Zagon, I.S. Topical Naltrexone Accelerates Full-

Thickness Wound Closure in Type 1 Diabetic Rats by Stimulating Angiogenesis.

Exp. Biol. Med., 2013, 238, 733–743.

[18] Immonen, J.; Zagon, I.; McLaughlin, P. Selective Blockade of the OGF-OGFr

Pathway by Naltrexone Accelerates Fibroblast Proliferation and Wound Healing.

Exp. Biol. Med., 2014, 239, 1300–1309.

[19] McLaughlin, P.J.; Cain, J.D.; Titunick, M.B.; Sassani, J.W.; Zagon, I.S. Topical

Naltrexone Is a Safe and Effective Alternative to Standard Treatment of Diabetic

Wounds. Adv. Wound Care, 2017, 6, 279–288.

[20] Zagon, I.; Klocek, M.; Sassani, J.; Mauger, D.; McLaughlin, P. Corneal Safety of

Topically Applied Naltrexone. J. Ocul. Pharmacol. Ther., 2006, 22, 377–387.

[21] Liang, D.; Sassani, J.W.; McLaughlin, P.J.; Zagon, I.S. Topical Application of

Naltrexone to the Ocular Surface of Healthy Volunteers: A Tolerability Study. J.

Ocul. Pharmacol. Ther., 2016, 32, 127–132.

[22] Warning for Regranex—Cream for Leg and Foot Ulcers; Consumer Updates;

Federal Drug Administration, 2008.

[23] Immonen, J.; Zagon, I.; McLaughlin, P. Topical Naltrexone as Treatment for Type 2

Diabetic Cutaneous Wounds. Adv. Wound Care, 2014, 3, 419–427.

143

[24] Ki67 - A Crucial Cellular Proliferation Marker (accessed Jun 13, 2018).

[25] Zagon, I.S.; Wu, Y.; McLaughlin, P.J. Opioid Growth Factor and Organ

Development in Rat and Human Embryos. Brain Res., 1999, 839, 313–322.

[26] Zagon, I.S.; Donahue, R.N.; McLaughlin, P.J. Opioid Growth Factor-Opioid Growth

Factor Receptor Axis Is a Physiological Determinant of Cell Proliferation in Diverse

Human Cancers. Am. J. Physiol. - Regul. Integr. Comp. Physiol., 2009, 297, 1154–

1161.

[27] Liskov, A.V.; Solnyshkova, T.G.; Frolov, B.A.; Pavlovichev, S.A. Effect of Naloxone

Hydrochloride on Osteogenesis in Chick Embryos. Bull. Exp. Biol. Med., 2005, 139,

331–333.

[28] Thakur, N.; DeBoyace, S.; Margulies, B. Antagonism of the Met5-Enkephalin-

Opioid Growth Factor Receptor-Signaling Axis Promotes MSC to Differentiate into

Osteoblasts: Antagonism of OGFR by Naloxone Increases Bone Formation. J.

Orthop. Res., 2015, n/a-n/a.

[29] Petrizzi, L.; Mariscoli, M.; Valbonetti, L.; Varasano, V.; Langhoff, J.; Von

Rechenberg, B. Preliminary Study on the Effect of Parenteral Naloxone, Alone and

in Association with Calcium Gluconate, on Bone Healing in an Ovine “Drill Hole”

Model System. BMC Musculoskelet. Disord., 2007, 8, 43–43.

[30] McCabe, L.R. Understanding the Pathology and Mechanisms of Type 1 Diabetic

Bone Loss. J. Cell. Biochem., 2007, 102, 1343–1357.

[31] Jenkins, S.J.; Ruckerl, D.; Cook, P.C.; Jones, L.H.; Finkelmsan, F.D.; van Rooijen, N.;

MacDonald, A.S.; Allen, J.E. Local Macrophage Proliferation, Rather than

144

Recruitment from the Blood, Is a Signature of [T.Sub.H]2 Inflammation. Science,

2011, 332, 1284.

[32] Campos Pastor, M.M.; López-Ibarra, P.J.; Escobar-Jiménez, F.; Serrano Pardo,

M.D.; García-Cervigón, A. Intensive Insulin Therapy and Bone Mineral Density in

Type 1 Diabetes Mellitus: A Prospective Study. Osteoporos. Int., 2000, 11, 455–

459.

[33] Gandhi, A.; Beam, H.A.; O’Connor, J.P.; Parsons, J.R.; Lin, S.S. The Effects of Local

Insulin Delivery on Diabetic Fracture Healing. Bone, 2005, 37, 482–490.

[34] Ippolito, J.A.; Krell, E.S.; Cottrell, J.; Meyer, R.; Clark, D.; Nguyen, D.; Sudah, S.;

Muñoz, M.; Lim, E.; Lin, A.; Lee, T.J.H.; O’Connor, J.P.; Benevenia, J.; Lin, S.S.

Effects of Local Vanadium Delivery on Diabetic Fracture Healing. J. Orthop. Res.,

2017, 35, 2174–2180.

[35] Hreha, J.; Wey, A.; Cunningham, C.; Krell, E.S.; Brietbart, E.A.; Paglia, D.N.;

Montemurro, N.J.; Nguyen, D.A.; Lee, Y.; Komlos, D.; Lim, E.; Benevenia, J.;

O’Connor, J.P.; Lin, S.S. Local Manganese Chloride Treatment Accelerates Fracture

Healing in a Rat Model. J. Orthop. Res., 2015, 33, 122–130.

[36] KAWAGUCHI, H.; KUROKAWA, T.; HANADA, K.; HIYAMA, Y.; TAMURA, M.; OGATA,

E.; MATSUMOTO, T. STIMULATION OF FRACTURE REPAIR BY RECOMBINANT

HUMAN BASIC FIBROBLAST GROWTH-FACTOR IN NORMAL AND

STREPTOZOTOCIN-DIABETIC RATS. ENDOCRINOLOGY, 1994, 135, 774–781.

145

[37] Gandhi, A.; Doumas, C.; O’Connor, J.P.; Parsons, J.R.; Lin, S.S. The Effects of Local

Platelet Rich Plasma Delivery on Diabetic Fracture Healing. Bone, 2006, 38, 540–

546.

[38] Wang, G.; Wang, J.; Fu, Y.; Bai, L.; He, M.; Li, B.; Fu, Q. Systemic Treatment with

Vanadium Absorbed by Coprinus Comatus Promotes Femoral Fracture Healing in

Streptozotocin-Diabetic Rats. Biol. Trace Elem. Res., 2013, 151, 424–433.

[39] Wang, L.X.; Jiang, H.L.; Du, S.L. Observed Impacts of Insulin Therapy on Callus Cell

Transforming Growth Factor-Beta 1 Expression in Diabetic Rats. Genet. Mol. Res.

GMR, 2015, 14, 5076–5084.

[40] Cignachi, N.; Pesquero, J.; Oliveira, R.; Etges, A.; Campos, M. Kinin B1 Receptor

Deletion Affects Bone Healing in Type 1 Diabetic Mice. J. Cell Physiol., 2015, 230,

3019–3028.

[41] Ko, K.I.; Coimbra, L.S.; Tian, C.; Alblowi, J.; Kayal, R.A.; Einhorn, T.A.; Gerstenfeld,

L.C.; Pignolo, R.J.; Graves, D.T. Diabetes Reduces Mesenchymal Stem Cells in

Fracture Healing through a TNFα-Mediated Mechanism. Diabetologia, 2015, 58,

633–642.

[42] Park, A.G.; Paglia, D.N.; Al-Zube, L.; Hreha, J.; Vaidya, S.; Breitbart, E.; Benevenia,

J.; O’Connor, J.P.; Lin, S.S. Local Insulin Therapy Affects Fracture Healing in a Rat

Model. J. Orthop. Res., 2013, 31, 776–782.

[43] Yee, C.S.; Xie, L.; Hatsell, S.; Hum, N.; Murugesh, D.; Economides, A.N.; Loots,

G.G.; Collette, N.M. Sclerostin Antibody Treatment Improves Fracture Outcomes

in a Type I Diabetic Mouse Model. Bone, 2015, 82, 122–134.

146

[44] Hough, S.; Avioli, L.V.; Bergfeld, M.A.; Fallon, M.D.; Slatopolsky, E.; Teitelbaum,

S.L. Correction of Abnormal Bone and Mineral Metabolism in Chronic

Streptozotocin-Induced Diabetes Mellitus in the Rat by Insulin Therapy.

Endocrinology, 1981, 108, 2228–2234.

[45] Guo, J.; Dong, W.; Jin, L.; Wang, P.; Hou, Z.; Zhang, Y. Hydrogen-Rich Saline

Prevents Bone Loss in Diabetic Rats Induced by Streptozotocin. Int. Orthop., 2017,

41, 2119–2128.

[46] Erdal, N.; Gürgül, S.; Demirel, C.; Yildiz, A. The Effect of Insulin Therapy on

Biomechanical Deterioration of Bone in Streptozotocin (STZ)-Induced Type 1

Diabetes Mellitus in Rats. Diabetes Res. Clin. Pract., 2012, 97, 461–467.

Vita Michelle B. Titunick

EDUCATION:

• Ph.D. Candidate, Human Anatomy December 2018 Pennsylvania State University, College of Medicine, Hershey, PA

• Graduate School Teaching Certificate March 2018 Pennsylvania State University, College of Medicine, Hershey, PA

• Charter Foundation Program December 2010 St. George’s University, Grenada, West Indies

• Bachelor of Arts, Cell Biology & Neuroscience and Evolutionary Anthropology May 2010 Rutgers University, New Brunswick, NJ

PUBLICATIONS:

• McLaughlin, P.J.; Cain, J.D.; Titunick, M.B.; Sassani, J.W.; Zagon, I.S. Topical Naltrexone Is a Safe and Effective Alternative to Standard Treatment of Diabetic Wounds. Adv. Wound Care, 2017, 6, 279–288.

• Stockdale, D.P.; Titunick, M.B.; Biegler, J.M.; Reed, J.L.; Hartung, A.M.; Wiemer, D.F.; McLaughlin, P.J.; Neighbors, J.D. Selective Opioid Growth Factor Receptor Antagonists Based on a Stilbene Isostere. Bioorg. Med. Chem., 2017, 25, 4464–4474.

AWARDS:

• Graduate Education Travel Award Fall 2016 TEACHING EXPERIENCE:

• Dissection- and lecture-based gross anatomy o Penn State Hershey Medical School 2016-2018 o Penn State Hershey Physician’s Assistant Program 2015-2018 o Penn State Hershey Graduate Program in Anatomy 2016-2018

• Dissection-based neuroanatomy o Penn State Hershey Medical School 2016-2017

• Lecture-based neuroanatomy o Penn State Hershey Neurology Residents 2015-2018