Characterization of Endocrine Disrupting Effects of Bisphenol A or 17α-Ethinyl Estradiol in Mouse Uterus

A dissertation submitted to the

Graduate School

of the University of Cincinnati

in partial fulfillment of the

requirements for the degree of

Doctor of Philosophy

in the Department of Pharmacology and Cell Biophysics

of the College of Medicine

by

Jessica A. Kendziorski

BA, DePauw University, 2010

Committee Chair: Scott M. Belcher, PhD

ii  

ABSTRACT

Bisphenol A (BPA) and 17α-ethinyl estradiol (EE) are estrogenic endocrine disrupting

chemicals (EDCs) that adversely affect the structure and function of the uterus. Humans are

ubiquitously exposed to BPA via consumption of contaminated food and beverage from

polycarbonate packaging or food cans. Humans are exposed to EE through the use of oral

contraceptives. While human exposure to these EDCs is widespread, little is known how these

compounds alter physiological responses that lead to increased incidence of uterine pathology.

The purpose of this dissertation was to investigate and characterize uterine pathologies

and associated alterations in immune responsiveness and fibrosis. Two mouse strains were used,

the CD1 and the C57Bl/6N strains. Exposure during adulthood included mating, parturition, and

offspring rearing and mice were exposed for 12-15 weeks. Whole life exposure included

placental transfer, as well as direct oral consumption, exposing mice until postnatal day (PND)

90. In order to closely mimic human exposure, mice were orally exposed through diet to known

concentrations of control (0 ppm), BPA (0.03, 0.3, 3, 30, or 300 ppm), or EE (0.0001, 0.001,

0.01, 0.1, or 1.3 ppm), which resulted in calculated BPA doses of 0.04, 0.4, 4, 40, and 400

mg/kg/day and EE doses of 0.00002, 0.0002, and 0.001 mg/kg/day. Other exogenous estrogenic

compounds from housing, bedding, and water were eliminated.

Two distinct immune-related and fibrotic uterine pathologies were identified. Pyometra

developed in C57Bl/6N mice exposed to BPA or EE during adulthood. An equine endometrosis-

like phenotype, characterized by increased gland nests and stromal and periglandular fibrosis,

naturally occurred in control C57Bl/6N mice. In CD1 mice, this fibrotic phenotype was present

in the 30 ppm BPA group. Exposure to BPA significantly increased Col1a1 and Col3a1

expression and decreased Mmp2 and Timp2 expression. Expression and activity of matrix

metalloproteinases, MMP2 and MMP14, were significantly decreased in both the control

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C57Bl/6N and 30 ppm BPA-exposed CD1 mice compared to control CD1 mice. However, both

strains presented with an increased immune response, quantified as the percentage of F4/80-

positive cells. The C57Bl/6N strain had a significant increase in endometrial macrophages at a

lower dose of BPA (0.03 ppm) than the CD1 strain (30 ppm) exposed during adulthood. In both

exposure models, CD1 mice had increased macrophages in the 30 ppm BPA group.

This dissertation research was the first report of BPA altering the immune response or

collagen accumulation in the uterus, leading to the induction of pyometra or an equine

endometrosis-like phenotype. It was also the first report of strain-specific differences in

sensitivity to the actions of BPA on these endpoints of interest. Finally, this research highlighted

the potential for exposure to estrogenic compounds such as BPA and EE to impact the

development and progression of fibrotic and immune-related uterine diseases. However, based

on the differences observed between the two generations, physiological changes that occur

during mating, pregnancy, and parturition may be necessary for estrogenic compounds to elicit

fibrotic effects. These studies also emphasized the importance of understanding differences in

sensitivity to estrogenic compounds between mouse strains.

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ACKNOWLEDGEMENTS

This dissertation work is the product of the encouragement of several colleagues, family,

and friends. First and foremost, I would like to acknowledge my advisor and mentor, Dr. Scott

Belcher, for his support and assistance during my time as a graduate student. Through his

invaluable mentoring and challenges, I have grown significantly in confidence as a scientist and

a person. I am truly grateful for the guidance and support he has provided over the years.

I would like to extend gratitude for the encouragement and constructive advice given by

my dissertation committee members, Drs, Jo El Schultz, William J. Ball, and Heather Patisaul.

This dissertation would not have been completed without their attention to progress and helping

me to remain focused on the final product.

I am gratefully indebted to the past and present members of the Belcher lab, specifically

Dr. Eric Kendig, Robin Gear, Dana Buesing, Susie Christie, Dr. Vinicius Carreira, and Clifford

Cookman, for their suggestions on experimental procedures, their help in conducting

experiments, and their friendships in general during my time in graduate school. I would like to

especially acknowledge Vini for his help with identifying pathologies, which laid the foundation

for a great part of this dissertation research.

I would like to acknowledge rotating graduate students, undergraduate students, summer

students, and students from England for their contributions to this research and for their

friendship, especially Joni Ford, Rachel Steinher, Ed Durley, and Kev Ungi. Their willingness to

do whatever was asked of them was highly appreciated.

I would like to thank all of the faculty, students, and staff in the Department of

Pharmacology and Biophysics and the associated graduate program. I would like to specifically

acknowledge Nancy Thyberg for everything that she has done, by keeping me updated on

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deadlines, helping to organize committee meetings, and being the person I would go to for any

questions. If she didn’t have the answer, she would find someone that could answer it.

Finally, I would like to thank my family and friends for their encouragement, love, and

support. Even though they had no idea what I was doing or why I was in school for 5+ years, my

parents always asked how things were going. I would like to thank my sister Sarah for always

being able to cheer me up if I was having a rough time. I would like to thank my friends, in

particular Ian, Jamey, Natalie, Josh, Mat, Leslie, Chad, Cassie, Mason, and Kyle, for their ability

to always make sure I had fun and didn’t go too crazy from being in the lab, and my boyfriend

Carson for putting up with my complaining and for his constant reassurance and love. I would

also like to thank Megan, Chenoa, Andrea, and Kelly for their laughs. Even though we were

hundreds of miles apart, they made me smile and uplifted my spirits every time we texted or

called.

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TABLE OF CONTENTS Page

ABSTRACT ii

ACKNOWLEDGEMENTS v

TABLE OF CONTENTS vii

LIST OF FIGURES AND TABLES xiv

LIST OF ABBREVIATIONS xviii

Chapter 1 INTRODUCTION 1

1.1 Statement of Purpose 2

1.1.1 Objective and Rationale 2

1.1.2 Central Hypothesis and Specific Aims 2

1.2 Definition of Endocrine Disrupting Chemicals 3

1.3 Prototypical Estrogenic EDC: Diethylstilbestrol 4

1.4 17α-Ethinyl Estradiol 6

1.5 Bisphenol A 7

1.5.1 Commercial Uses of BPA and Routes of Exposure 7

1.5.2 Pharmacokinetics and Toxicologic Exposure Limits of BPA 8

1.5.3 Receptor Binding of BPA 8

1.5.4 Endocrine Disrupting Effects of BPA in the Uterus 11

1.5.5 Extracellular Matrix Alterations in Response to BPA 12

1.6 Strain Susceptibility to Effects of Estrogenic Compounds 13

1.6.1 Susceptibility to Uterine Weight Increases and Morphological Alterations 15

1.6.2 Differences in Circulating Sex Hormones and ERα Levels 15

1.6.3 Quantitative Trait Loci Controlling Phenotypic Variation 16

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1.6.4 Differential Sensitivity of the Ovary and Vagina to Estrogenic Compounds 17

1.6.5 Differential Sensitivity of Other Tissues to Estrogenic Compounds 18

1.7 The Endocrine System 19

1.8 Nuclear Hormone Receptors: ERα and ERβ 20

1.8.1 Structure 20

1.8.2 Nuclear Transactivation, Membrane-Initiated, 22

and Rapid Signaling Mechanisms

1.8.3 Role of ERs in Uterine Structure and Reproductive Function 26

1.8.3.1 Postnatal Uterine Development 26

1.8.3.2 Impact on Gene Expression 27

1.8.3.3 ERα in the Brain and Impact on Cyclicity 28

1.8.3.4 Role of GPR30 in the Uterus 28

1.9 The Cycling and Pregnant Uterus 29

1.9.1 Circulating Hormones and Tissue Remodeling 29

1.9.2 Role of Extracellular Matrix in Uterine Structure and Function 32

1.9.3 Role of Extracellular Matrix during Pregnancy 36

1.10 Uterine Disorders 39

1.10.1 Pyometra 39

1.10.1.1 Description, Etiology, and Treatment of Pyometra 39

1.10.1.2 Role of Bacteria and the Immune System 40

in the Development of Pyometra

1.10.1.3 Involvement of Sex Hormones and Collagen Accumulation 41

in Pyometra

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1.10.2 Endometriosis 41

1.10.2.1 Description, Symptoms, and Treatment of Endometriosis 41

1.10.2.2 Role of Extracellular Matrix in Pathogenesis of Endometriosis 42

1.10.3 Leiomyoma 43

1.10.3.1 Description, Symptoms, and Treatment of Leiomyoma 43

1.10.3.2 Etiology of Leiomyoma 45

1.10.3.3 Regulation of Collagen Accumulation in Leiomyoma 47

1.10.4 Equine Endometrosis 47

1.10.4.1 Description, Etiology, and Treatment of Equine Endometrosis 47

1.10.4.2 Progression of Equine Endometrosis 50

1.10.4.3 Characterization of Extracellular Matrix Alterations 51

in Equine Endometrosis

1.10.4.4 Fetal Foal Loss 52

1.11 Statement of Purpose 54

1.11.1 Objective and Rationale 54

1.11.2 Central Hypothesis and Specific Aims 54

Chapter 2 Methods 55

2.1 BPA and EE Exposure 56

2.1.1 Animal Housing 56

2.1.2 Exposure 56

2.1.3 Breeding 59

2.1.4 Vaginal Cytology 59

2.2 Assessment of Pathology 60

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2.2.1 Necropsy and Tissue Preparation 60

2.2.2 Hematoxylin and Eosin Staining 61

2.2.3 Myometrial Wall Thickness 62

2.2.4 Histologic Assessment of Uterine Pathology 62

2.3 Assessment of Collagen Synthesis and Degradation and Immune Response 63

2.3.1 Picrosirius Red Staining and Collagen Accumulation 63

2.3.2 Immunohistochemistry 63

2.3.3 Quantitative RT-PCR 66

2.3.4 SDS-PAGE and Immunoblotting 70

2.3.5 Gelatin Zymography 72

2.4 Statistical Methods 74

Chapter 3 Strain Specific Induction of Pyometra and Differences in Immune 75

Responsiveness in Mice Exposed to 17α-Ethinyl Estradiol or the

Endocrine Disrupting Chemical Bisphenol A

3.1 Abstract 77

3.2 Introduction 78

3.3 Materials and Methods 80

3.3.1 Animal Husbandry and Dietary Exposures 80

3.3.2 Monitoring of Estrous 81

3.3.3 Necropsy and Tissue Preparation 82

3.3.4 Immunohistochemistry 82

3.3.5 Statistical Analysis 83

3.4 Results 84

xi  

3.4.1 Diet Consumption 84

3.4.2 Effects on Body Weight 86

3.4.3 Fertility and Fecundity 88

3.4.4 Necropsy and Organ Weights 90

3.4.5 Uterine Pathology: C57Bl/6 Pyometra 92

3.4.6 Uterine Pathology: Effects on Uterine Morphology and 94

Macrophage Infiltration

3.5 Discussion 99

3.6 Acknowledgements 103

3.7 Supplemental Results 104

3.7.1 Uterine Morphology: Myometrial Wall Thickness 104

3.7.2 Immune Response in Uteri of CD1 and C57Bl/6N Mice 105

Chapter 4 Strain-Specific Induction of Endometrial Periglandular Fibrosis in Mice 108

Exposed During Adulthood to the Endocrine Disrupting Chemical Bisphenol A

4.1 Abstract 110

4.2 Introduction 111

4.3 Materials and Methods 116

4.3.1 Animal Husbandry and Necropsy 116

4.3.2 Histology and Immunohistochemistry 117

4.3.3 Picrosirius Red Stain and Collagen Assessment 118

4.3.4 Quantitative RT-PCR 119

4.3.5 SDS-PAGE and Immunoblotting 119

4.3.6 Gelatin Zymography 120

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4.3.7 Statistical Analysis 121

4.4 Results 122

4.4.1 Uterine Pathology: Gland Nests and Periglandular Fibrosis in C57Bl/6N 122

and CD-1 Mice

4.4.2 Analysis of Collagen and MMP mRNA Expression 126

4.4.3 Analysis of Protein Expression and Activity of MMPs 127

4.4.4 Immunolocalization of MMP2 and MMP14 in the Endometrium 128

4.4.5 Immunohistochemical Characterization of an Endometrosis-Like Phenotype 131

4.5 Discussion 133

4.6 Acknowledgements 139

Chapter 5 Effects of Bisphenol A and 17α-Ethinyl Estradiol in Nulligravida Uterus 140

of CD1 and C57Bl/6N Mice: Whole Life Exposure

5.1 Introduction 141

5.2 Methods 143

5.2.1 Exposure 143

5.2.2 Myometrial Wall Thickness 143

5.2.3 Uterine Pathology 144

5.2.4 Assessment of Collagen Accumulation and Immune Response 144

5.2.5 Statistical Analysis 145

5.3 Results 146

5.3.1 Uterine Morphology: Myometrial Wall Thickness 146

5.3.2 Uterine Pathology 147

5.3.3 Collagen Accumulation 148

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5.3.4 Quantification of Gland Nest Density 150

5.3.5 Immune Response 151

5.4 Discussion 153

Chapter 6 SUMMARY AND CONCLUSIONS 156

6.1 Significance 157

6.2 Limitations 160

6.3 Future Directions 161

LIST OF PUBLICATIONS AND ABSTRACTS 167

BIBLIOGRAPHY 169

xiv  

LIST OF FIGURES AND TABLES Page

FIGURES

Chapter 1 INTRODUCTION

Figure 1 Chemical structure, name, and CAS ID of estrogenic compounds 3

Figure 2 HPG axis and negative feedback loop 20

Figure 3 Expression of ERα and ERβ in human tissues 21

Figure 4 Schematic of domain structure of ERα and ERβ 22

Figure 5 Classical model of nuclear hormone receptor translocation 23

Figure 6 Normal uterine structure of adult CD1 mouse 30

Figure 7 Simplified representation of the human menstrual cycle 31

Figure 8 Regulation of collagen synthesis and degradation 33

Figure 9 Schematic representation of MMP structures 35

Figure 10 Schematic of decidualization and implantation in the human uterus 37

Chapter 2 METHODS

Figure 1 Exposure paradigm for sires, dams, and offspring in control and BPA or 57

EE exposure groups

Chapter 3 Pyometra and Immune Responsiveness

Figure 1 Histological and immunohistochemical assessments of C57Bl/6 uterus 93

affected by severe pyometra

Figure 2 Effects of oral exposure to BPA on uterine morphology and macrophage 95

infiltration in C57Bl/6 mice without overt pyometra

Figure 3 Effects of oral exposure to BPA on uterine morphology and macrophage 97

xv  

infiltration in CD1 mice

Figure 4 Quantitative comparison of F4/80 immunoreactive cell numbers in the uterus 98

of control and BPA-exposed C57Bl/6 and CD1 mice

Figure 5 Myometrial wall thickness in mice across exposure groups and strain 105

Figure 6 Quantification of F4/80-positive cells in BPA- or EE-exposed CD1 and 107

C57Bl/6N adult mice

Chapter 4 Equine Endometrosis-Like Phenotype and Fibrosis

Figure 1 Effects of oral exposure to BPA or EE on uterine morphology and 123

collagen accumulation in C57Bl/6N mice

Figure 2 Effects of oral exposure to BPA or EE on uterine morphology and 124

collagen accumulation in CD-1 mice

Figure 3 Quantification of gland nest density in the uteri of control and 125

BPA-exposed CD-1 and C57Bl/6N mice

Figure 4 Relative expression levels and activity of mRNA and proteins involved in 126

collagen synthesis and degradation

Figure 5 Strain differences in the immunohistochemical localization of MMP2 and 130

MMP14 in the uteri of control CD-1 and C57Bl/6N mice

Figure 6 Immunohistochemical characterization of endometrosis-like phenotype 131

across BPA exposure groups or between strains

Chapter 5 Effects of BPA and EE in Uterus: Whole Life Exposure

Figure 1 Exposure paradigm for sires, dams, and offspring in control and BPA or 143

EE exposure groups

xvi  

Figure 2 Myometrial wall thickness in CD1 mice across exposure groups 146

Figure 3 Effects of oral BPA or EE exposure on uterine morphology 150

and collagen accumulation in CD1 offspring

Figure 4 Quantification of gland nest density in the endometrium 151

of BPA- and EE-exposed CD1 and C57Bl/6N mice at PND90

Figure 5 Quantification of F4/80-positive cells in BPA- or EE-exposed CD1 offspring 152

TABLES

Chapter 1 INTRODUCTION

Table 1 Summary of studies examining sensitivity of rodent strains to 14

estrogenic compounds

Table 2 Characteristics of equine endometrosis phenotypes 51

Chapter 2 METHODS

Table 1 Composition of defined control and test diets 58

Table 2 Amounts of EE, BPA, and dyes incorporated into diets 58

Table 3 Primary and secondary antibodies 66

Table 4 RNA concentrations 68

Table 5 TaqMan probe sets used for qRT-PCR analysis 69

Chapter 3 Pyometra and Immune Responsiveness

Table 1 Estimated food consumption, caloric intake, and dose 85

Table 2 Female body weight 87

Table 3 Measures of fertility and fecundity 89

Table 4 Organ weights 91

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Chapter 4 Equine Endometrosis-Like Phenotype and Fibrosis

Table 1 Incidence of uterine pathology in CD-1 and C57Bl/6N mice 122

at 19-23 weeks of age

Chapter 5 Effects of BPA and EE in Uterus: Whole Life Exposure

Table 1 Uterine pathology in CD1 and C57Bl/6N offspring 149

Chapter 6 SUMMARY AND CONCLUSIONS

Table 1 Comparison of relative expression and activity of collagens and MMPs 159

in mouse strains

xviii  

LIST OF ABBREVIATIONS

11β-HSD2 11β-hydroxysteroid dehydrogenase

α-MG α-macroglobulin

α-SMA α-smooth muscle actin

ABC avidin-biotin complex

AF1 ligand-independent activation factor 1

AF2 ligand-dependent activation factor 2

ANOVA analysis of variance

APS ammonium persulfate

AR androgen receptor

BM basement membrane

BPA bisphenol A

BSA bovine serum albumin

CAL calbindinD9k

cAMP cyclic adenosine monophosphate

CEH cystic endometrial hyperplasia

CO2 carbon dioxide

DAB 3’3-diaminobenzidine

DBD DNA binding domain

DES diethylstilbestrol

dH2O distilled water

DMSO dimethyl sulfoxide

E2 17β-estradiol

eAD-MSC equine adipose derived-mesenchymal stem cell

ECM extracellular matrix

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EDC endocrine disrupting chemical

EE 17α-ethinyl estradiol

EGFR epidermal growth factor receptor

Endo endometrium

eNOS endothelial nitric oxide synthase

EPF endometrial periglandular fibrosis

ER estrogen receptor

ERα estrogen receptor α

ERβ estrogen receptor β

ERE estrogen response element

ERK extracellular signal-regulated kinase

ERR estrogen-related receptor

ERRα estrogen-related receptor α

ERRβ estrogen-related receptor β

ERRγ estrogen-related receptor γ

ERRE estrogen-related receptor response element

F0 parental generation

F4/80 glycoprotein marker on mouse macrophages

FDA US Food and Drug Administration

FSH follicle stimulating hormone

g gram

GH growth hormone

GnRH gonadotrophin releasing hormone

GPER1 G protein-coupled estrogen receptor 1

GPR30 G protein-coupled receptor 30

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H&E hematoxylin and eosin

H2O2 hydrogen peroxide

hCG human chorionic gonadotropin

HPG hypothalamus-pituitary-gonadal

HSP heat shock protein

IC50 half maximal inhibitory concentration

IL interleukin

IM inner myometrium

kcal kilocalorie

KD dissociation constant

kg kilogram

Kiss1 kisspeptin

LBD ligand binding domain

LE luminal epithelium

LH luteinizing hormone

LPS lipopolysaccharide

LRP low density lipoprotein receptor-related protein

µg microgram

µL microliter

µm micrometer

µM micromolar

M molar

MAPK mitogen-activated protein kinase

MED12 mediator complex subunit 12

mg milligram

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mL milliliter

mm millimeter

mM millimolar

MMP matrix metalloproteinase

MR magnetic resonance

MSC mesenchymal stem cell

Myo myometrium

n sample size

NBF neutral buffered formalin

NET neutrophil extracellular trap

NHR nuclear hormone receptor

NIEHS National Institute of Environmental Health and Safety

ng nanogram

nM nanomolar

NO nitric oxide

NOAEL no observed adverse effect level

NTD N-terminal domain

OECD Organisation for Economic Co-operation and Development

OM outer myometrium

P4 progesterone

PBS phosphate buffered saline

PGC-1α peroxisome proliferator-activated receptor coactivator-1α

PGES prostaglandin E2 synthase

PGFS prostaglandin F2α synthase

PI3K phosphoinositide-3-kinase

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PKA protein kinase A

PKB protein kinase B

pM picomolar

PND postnatal day

ppm parts per million

PR progesterone receptor

Prl prolactin

PTGS-2 prostaglandin-endoperoxide synthase

QTL quantitative trait loci

RBA relative binding affinity

RT room temperature

SD standard deviation

SDS sodium dodecyl sulfate

SDS-PAGE sodium dodecyl sulfate-polyacrylamide gel electrophoresis

SEM standard error of the mean

SNP single nucleotide polymorphism

T testosterone

TBS Tris-buffered saline

TBST Tris-buffered saline with Tween-20

TEMED tetramethylethylenediamine

TG2 tissue transglutaminase 2

TGF-β1 transforming growth factor-β1

THR thyroid hormone receptor

TIMP tissue inhibitor of metalloproteinase

TLR toll-like receptor

xxiii  

UC uterocalin

UF uteroferrin

UG uteroglobin

1  

CHAPTER 1

INTRODUCTION

2  

1.1 Statement of Purpose

1.1.1 Objective and Rationale

The overall objective of this dissertation research was to characterize the endocrine

disrupting effects of BPA or EE exposure in the uterus of two mouse strains. Two exposure

paradigms were used: 1) exposure in adulthood, during which time mice mated and produced a

litter, and 2) whole life exposure without breeding. Previous studies examined incidence of

pathologies after a much longer exposure time period than is proposed in this dissertation

research. Uterine immune responsiveness and fibrosis are components of several human and

animal uterine pathologies, and the effect of BPA or EE exposure on these endpoints has not

previously been studied.

1.1.2 Central Hypothesis and Specific Aims

The central hypothesis for this dissertation was that exposure to estrogenic EDCs, such as

BPA or EE, will influence E2-regulated uterine physiology, leading to an increased incidence of

uterine pathology. A second hypothesis was that a greater sensitivity in the C57Bl/6N strain to

the estrogenic actions of EDCs also increases the incidence of uterine pathology compared to the

CD1 strain.

The specific aims of this dissertation research included 1) investigating the endocrine

disrupting actions of EDCs in both exposure models on the development of uterine pathologies,

2) examining the effects of EDCs in both exposure models on the immune responsiveness in the

uterus, 3) investigating the ECM changes associated with EDC exposure in both models, and 4)

comparing the pathological, immune, and ECM alterations between the C57Bl/6N and CD1

strains.

3  

1.2 Definition of Endocrine Disrupting Chemicals

The first use of the term “endocrine disrupting chemical” (EDC) in a scientific paper was

by Colborn et al. (1993), leading to a surge of research on the deleterious effects of EDCs on

reproduction, development, and other processes and organ systems. According to the National

Institute of Environmental Health Sciences (NIEHS), EDCs were defined as “naturally occurring

compounds or man-made substances that may mimic or interfere with the function of hormones

in the body” (NIEHS 2010). The Endocrine Society defined EDCs as “an exogenous chemical,

or mixture of chemicals, that interferes with any aspect of hormone action” (Gore et al. 2014).

Given such a broad definition and the numerous hormones and hormone targets present in the

body, thousands of natural and synthetic compounds may have endocrine-disrupting properties.

Of particular interest to this dissertation research were compounds that lead to estrogenic

endocrine disrupting activities, such as diethylstilbestrol (DES), 17α-ethinyl estradiol (EE), or

bisphenol A (BPA) (Figure 1).

Figure 1. Chemical structure, name, and CAS ID of estrogenic compounds.

4  

1.3 Prototypical Estrogenic EDC: Diethylstilbestrol

Diethylstilbestrol (DES) was one of the first compounds characterized as an estrogenic

EDC. Due to its known estrogenic effects, DES is often used as a positive control in EDC

studies. It was developed in 1938 as a synthetic estrogenic compound and prescribed to pregnant

women in the 1940s to early 1970s to prevent miscarriage and other pregnancy complications

(CDC 2012, NCI 1976). In the 1950s, DES treatment was found to be ineffective in reducing the

incidence of miscarriage and potentially led to premature labor (Dieckmann et al. 1953). In 1971,

it was found that using DES during pregnancy led to an increased incidence of a rare type of

cancer called vaginal clear cell adenocarcinoma in exposed female offspring as young as 15

years of age (Herbst et al. 1971). These findings led the US Food and Drug Administration

(FDA) to issue a warning to physicians to stop prescribing DES (FDA 1972).

Adverse pathophysiological effects are still being diagnosed in women exposed to DES

during pregnancy (“DES mothers”), their male and female offspring (“DES sons and

daughters”), and even in the third generation (“DES grandsons and granddaughters”). DES

mothers had an increased risk of developing breast cancer, but not other hormonal cancers such

as ovarian or endometrial cancer (Titus-Ernstoff et al. 2001, Colton et al. 1993). Animal studies

reported increased mammary tumor incidences (Greenman et al. 1987, Greenman et al. 1986), as

well as fibrosis and degeneration in ovaries (Hong et al. 2010) and thyroid neoplasias (Greenman

et al. 1990). However, these effects appeared to be strain dependent (Eastin et al. 2001).

It was already noted that DES daughters developed vaginal clear cell carcinoma, an

extremely rare cancer especially in young women. Other abnormalities were found in these

women as well, including cervical dysplasia (Robboy et al. 1984) and numerous forms of

gynecological cancers (Verloop et al. 2000). In animal studies, prenatal or neonatal DES

5  

exposure caused numerous morphological and pathological alterations either in development or

adulthood, including hyperplasia, adenocarcinoma, pyometra, distended or cystic glands, and

decreased epithelial cell proliferation (Bosquiazzo et al. 2013, Yoshida et al. 2011, Yamamoto et

al. 2003, Yoshida et al. 1999, McLachlan et al. 1980), partially due to DES binding to ERα

(Couse and Korach 2004). Alterations in gene expression related to carcinogenesis, leiomyoma,

and energy metabolism in epithelial cells were reported in DES-exposed animals (Yin et al.

2012, Greathouse et al. 2008, Newbold et al. 2007a), as well as alterations in methylation

patterns (Bromer et al. 2009, Sato et al. 2009), suggesting that DES exposure also has epigenetic

effects on the offspring of DES-exposed animals and humans. In DES sons, there is much debate

as to whether in utero DES exposure increased the risk for testicular or prostate cancer due to the

inconclusive results from epidemiological studies conducted (IARC 2012). In utero DES

exposure resulted in testicular atrophy, decreased spermatogenesis, and alterations in gene

expression in animal studies (Warita et al. 2012, LaRocca et al. 2011, Li et al. 2011, Hong et al.

2010).

Children of mothers who were exposed in utero to DES are now being studied for

potential detrimental effects in the third generation. Increases in reported birth defects in the

third generation were mainly associated with skeletal/bone, heart, or male reproductive system

(Titus-Ernstoff et al. 2010). In third generation daughters, there were irregular menstrual cycles

and the potential for fertility issues (Titus-Ernstoff et al. 2006). In rodent studies, third

generation rodents had increased incidence of uterine adenocarcinomas and ovarian

cystadenocarcincomas (Newbold et al. 1998, Turusov et al. 1992, Walker 1984). Even though

direct DES exposure has not occurred in over 40 years, endocrine disrupting effects are still

6  

being found in the offspring of prenatally exposed humans, supported by multigenerational

effects observed in rodent studies.

1.4 17α-Ethinyl Estradiol

17α-ethinyl estradiol (EE) is an orally bioavailable synthetic estrogen pharmaceutical

commonly used in oral contraceptive formulations. Due to its structural similarities to E2, EE

also binds to and activates the ERs (Blair et al. 2000). The major form of metabolism of EE is

oxidation by cytochrome P450 3A4 (Guengerich 1990, 1988). However, EE can also interact

with and inactivate P450 2B1 and 2B6 (Kent et al. 2006, Kent et al. 2002), preventing the

enzymes from metabolizing their target compounds and potentially leading to adverse drug-drug

interactions. Excretion occurs via the urine or feces and leads to contamination of waterways and

subsequent alterations in aquatic wildlife species (Bhandari et al. 2015).

Due to its estrogenic activity, EE is used as a positive control for estrogen-like effects

when studying oral exposures to other estrogenic EDCs. Exposure to high concentrations of EE

had detrimental effects on several parameters associated with reproduction and uterine structure

and function. In a multigenerational study, EE exposure resulted in an earlier age for vaginal

opening and increased the percentage of rats with abnormal cyclicity and cycle length (Delclos et

al. 2009). Pathology such as hyperplasia and alterations in uterine gland numbers was also

observed (Takahashi et al. 2014, Delclos et al. 2009). Uterine weight was increased by EE under

all tested conditions across species, strain, laboratory, and housing (Kanno et al. 2001). Induction

of E2-responsive gene expression in the uterus was similar between EE exposure and E2

treatment (Hyder et al. 1999); however, EE exposure decreased ERα and ERβ expression in both

the uterus and the female hypothalamus (Rebuli et al. 2014, Takahashi et al. 2014, Kang et al.

2003). Gene expression alterations in response to EE exposure included genes involved in cell

7  

proliferation and apoptosis, protein metabolism and transport, regulation of transcription, water

imbibition, Wnt signaling, ECM remodeling, alternative complement activation, and estrogen-

mediated Ca2+ signaling (Fong et al. 2007, Heneweer et al. 2007, Katayama et al. 2006, Naciff et

al. 2003).

1.5 Bisphenol A

1.5.1 Commercial Uses of BPA and Routes of Exposure

Bisphenol A (BPA) is a synthetic compound that leaches from polycarbonate plastics and

epoxy resins of food cans to contaminate the contents of the food or beverage container, in

particular under heated or acidic conditions (Cooper et al. 2011, Carwile et al. 2009, Grumetto et

al. 2008, Le et al. 2008, Krishnan et al. 1993). The main route of human exposure occurs orally

through consumption of contaminated food and beverages (Hoekstra and Simoneau 2013,

Teeguarden et al. 2011, Carwile et al. 2009, Grumetto et al. 2008, Vandenberg et al. 2007a).

Humans have transdermal exposure by handling of thermal receipt paper and intravenous

exposure through tubing in medical equipment (Ehrlich et al. 2014, Calafat et al. 2009). Prenatal

exposure occurs via placental transfer to the amniotic fluid (Vom Saal et al. 2014, Edlow et al.

2012, Nishikawa et al. 2010, Ikezuki et al. 2002, Yamada et al. 2002). Greater than 93% of

humans have detectable urinary levels of BPA and metabolites (Calafat et al. 2008), highlighting

ubiquitous BPA exposure in humans. Of importance for consistency between rodent

experimental studies were controlling the potential exogenous sources of BPA and other

estrogenic compounds. Used polycarbonate caging leached BPA into drinking water

(Howdeshell et al. 2003). Diet, bedding, and drinking water also played a major factor in excess

and uncontrolled BPA exposure in rodent studies (Thigpen et al. 2013). Finally, route of

exposure had a significant impact. Because the main route of exposure in humans is orally,

8  

experimental animal studies should also use the oral route of exposure in order to extrapolate

results to humans.

1.5.2 Pharmacokinetics and Toxicologic Exposure Limits of BPA

Depending on route of exposure, species, and dose, BPA concentrations reached a

maximal serum concentration between 1-6 hours and declined by 24 hours (Sieli et al. 2011,

Taylor et al. 2011). The main form of metabolism of BPA was via glucuronidation in the liver in

both humans and rodents (Vandenberg et al. 2007a, Yokota et al. 1999). However, humans

excreted BPA and its metabolites in the urine, while the main form of excretion in rodents was in

the feces (Vandenberg et al. 2007a). An acceptable oral limit of BPA exposure in humans was

set at 0.05 mg/kg body weight/day (IRIS 1988). Based on several multigenerational studies in

rodents, the no observed adverse effect level (NOAEL) for BPA was set at 5 mg/kg/day (Tyl et

al. 2008, Tyl et al. 2002). The NOAEL indicates the highest concentration of a compound that

causes no detectable alterations in the endpoint being studied and is dependent on the

experimental design.

1.5.3 Receptor Binding of BPA

Based on in vitro studies, BPA bound to several nuclear hormone receptor targets, but

had agonist or antagonist roles depending on the receptor. The main receptor targets for BPA

were estrogen receptor (ER) α and ERβ, which BPA activated at nM concentrations (Li et al.

2013, Wetherill et al. 2007, Matthews et al. 2001, Kuiper et al. 1998). The relative binding

affinity (RBA) of BPA to ERα was very low and was calculated to be between 0.008 and

0.065%, compared to 100% for E2 (Yiu et al. 2014, Blair et al. 2000). The RBA indicates the

affinity of a ligand to bind to a specific target when compared to a ligand with a known affinity

9  

set to 100%. A low RBA, in the case of BPA, means that there are fewer intermolecular forces

between it and ERα compared to between E2 and ERα and that E2 will bind to ERα before BPA

binds. The IC50 is an indicator of the concentration of ligand needed to displace a known ligand

from the binding pocket of a receptor. Using an ERα competitive binding assay with [3H]-E2, E2

displaced 50% of the radioligand at 8.99 x 10-10 M and BPA had an IC50 of 1.17 x 10-5 M (Blair

et al. 2000), meaning that it took a higher concentration of BPA to displace E2 from ERα.

However, the on-rate of BPA binding to ERα was very fast compared to its RBA. The kon for

BPA was 11 x 103 M-1 s-1, compared to the kon for E2 of 1000 x 103 M-1 s-1 (Yiu et al. 2014),

suggesting that the fast on-rate of binding could produce a transient surge in downstream effects.

This transient effect could be prolonged with the recruitment of a co-activator to keep BPA

bound longer. Through mathematical modeling, it was proposed that a 10 minute surge of BPA

from pM to nM concentration in the blood could increase the fraction of BPA-activated ERα to

high enough levels to induce a physiological response (Yiu et al. 2014, Welshons et al. 2003).

The binding and activation of ERβ by BPA had significant physiological effects in the

brain, heart, and pancreas. Rapid activation of ERβ by BPA in estrogen-sensitive developing

cerebellar granule cells led to E2-like neurotoxicity and cell death (Le and Belcher 2010). In the

heart, BPA promoted sex-specific arrhythmogenesis in female rodents via rapid ERβ-mediated

alterations in Ca2+ release and reuptake (Liang et al. 2014, Gao et al. 2013, Belcher et al. 2012,

Yan et al. 2011). Similarly, BPA exposure significantly altered sex-specific Esr2 expression in

multiple areas of the brain, with potential adverse downstream effects related to brain

development. In the developing amygdala, hypothalamus, and other areas related to sociosexual

behavior, altered expression of Esr2 led to BPA-induced anxiogenic behavior (Cao et al. 2014,

10  

Patisaul et al. 2012) and the potential for later-in-life defects in neuroendocrine function related

to reproduction (Rebuli et al. 2014, Cao et al. 2013, Cao et al. 2012).

Other receptor targets for BPA include the androgen receptor (AR) and thyroid hormone

receptor (THR), to which BPA bound and inhibited at µM concentrations in vitro (Teng et al.

2013, Moriyama et al. 2002, Paris et al. 2002, Sohoni and Sumpter 1998). This antagonistic

activity led to significant impairment of stromal and glandular cell function in the prostate and

could be involved in enhancing the invasiveness of prostate cancer (Ramos et al. 2001) or on

vertebrate development (Heimeier et al. 2009). The binding of BPA to AR also reduced sperm

quality and disrupted the hypothalamus-pituitary-gonadal (HPG) axis (Wisniewski et al. 2015,

Ramos et al. 2003).

Another receptor that BPA was shown to bind to in vitro is estrogen-related receptor

(ERR) γ. When examining crystal structures, BPA bound to the ligand binding domain of ERRγ

without changing internal structures of the binding pocket; in fact, BPA appeared to stabilize the

constitutively active conformation of ERRγ (Liu et al. 2014, Matsushima et al. 2007). Through

the use of a saturation binding assay, BPA had a KD of 5.5 nM for ERRγ and only one of the

phenol hydroxyl groups was essential for full binding (Okada et al. 2008). Accumulation of BPA

in the placenta was in part mediated by its binding to ERRγ, leading to increased placental

transfer to the developing fetus and significant prenatal exposure (Takeda et al. 2009). Otolith

development in zebrafish was also impaired by the interaction of BPA and ERRγ, suggesting a

potential role in deafness or hearing loss (Tohme et al. 2014)

Rapid signaling actions of BPA occurred through binding to G protein-couple receptor

(GPR) 30. In an ER-negative cell line, BPA had an IC50 of 650 nM for displacing E2 from the

transfected GPR30 (Thomas and Dong 2006). Treatment with BPA also increased

11  

phosphorylation of extracellular signal-regulated kinase (ERK) in several cell types via GPR30

activation (Ge et al. 2014, Watson et al. 2014, Dong et al. 2011). Rapid signaling of BPA

binding to GPR30 affected spermatogonial cell proliferation, as well as testicular seminoma

tumor cell and breast cancer cell proliferation (Sheng et al. 2013, Chevalier et al. 2012, Pupo et

al. 2012, Sheng and Zhu 2011).

1.5.4 Endocrine Disrupting Effects of BPA in the Uterus

Exposure to BPA led to endocrine disrupting effects in the uterus, including both gross

and microscopic alterations in structure and significant impairment of uterine function. The

uterotrophic assay assessing uterine weight increases was often used to determine the estrogen-

like activity of EDCs, based on a protocol outlined by the Organisation for Economic Co-

operation and Development (OECD) (Kanno et al. 2003, Kanno et al. 2001). In rodents, BPA

exposure significantly increased uterine weights (Kendig et al. 2012, Markey et al. 2001,

Papaconstantinou et al. 2000, Ashby and Tinwell 1998), supporting that BPA has estrogen-like

activity in the uterus.

Later in life morphological alterations in the uterus from a prenatal or developmental

BPA exposure included changes in luminal and glandular epithelial and stromal thickness and

changes in the number of apoptotic or proliferative cells (Vigezzi et al. 2015, Mendoza-

Rodriguez et al. 2011, Schonfelder et al. 2004, Steinmetz et al. 1998). Increased incidence of

cystic endometrial hyperplasia, squamous metaplasia, adenomyosis, leiomyoma, and stromal

polyps due to BPA exposure were also observed (Vigezzi et al. 2015, Newbold et al. 2009,

2007b, Markey et al. 2005). While BPA altered ERα and PR expression, it is debated if

expression was increased or decreased (Vigezzi et al. 2015, Mendoza-Rodriguez et al. 2011,

Markey et al. 2005, Schonfelder et al. 2004).

12  

Exposure to BPA not only impacted uterine structure, but it also significantly impaired

the function of the uterus. Epidemiological studies suggested an association between increasing

urinary or serum BPA concentrations and decreased rates of implantation success or successful

reproductive outcomes (Shen et al. 2015, Peretz et al. 2014, Ehrlich et al. 2012a, Ehrlich et al.

2012b, Mok-Lin et al. 2010). The number of implantation sites was greatly reduced in BPA-

exposed rodents due to decreased Hoxa10 expression, which led to significant impairment of

fertility (Varayoud et al. 2011). Contractility of the uterus was also decreased, which would

result in a reduced ability to deliver a full-term fetus (An et al. 2013). Expression of Hox and Wnt

genes critical for organ development was significantly decreased in the uteri of fetal rhesus

macaques (Calhoun et al. 2014), leading to impaired embryonic development. Exposure to BPA

also increased expression of c-fos, which stimulates cell proliferation, Pgr or the progesterone

receptor, and proteins involved in Ca2+ binding and the immune response (Hewitt and Korach

2011, Hong et al. 2006, Naciff et al. 2002, Steinmetz et al. 1998). In two- and three-generational

reproductive toxicity studies in rodents, only very high BPA concentrations altered fertility

parameters (Tyl et al. 2008, Tyl et al. 2002), but otherwise very few studies have examined the

effects of BPA exposure during adulthood in sexually mature rodents.

1.5.5 Extracellular Matrix Alterations in Response to BPA

Few studies have examined the effects of BPA exposure on the extracellular matrix

(ECM) in any tissue. In the fetal mammary gland, collagen localization but not density was

altered by BPA exposure (Vandenberg et al. 2007b). Whole life exposure to BPA led to sex-

specific alterations in ECM genes and proteins in the mouse heart (Belcher et al. 2015). In the

human ovarian cancer cell line OVCAR-3, BPA significantly increased MMP2 and MMP9

mRNA expression and activity of the enzymes (Ptak et al. 2014). Granulosa-lutein cells treated

13  

with BPA had increased MMP9 secretion (Dominguez et al. 2008). While the impact of BPA

exposure on ECM alterations has been studied in various tissues, it still remains unknown if BPA

exposure alters ECM genes and proteins in the uterus.

1.6 Strain Susceptibility to Effects of Estrogenic Compounds

Genetic variation between rodent strains can impact physiological or pathological

responses to estrogenic compounds (Wall et al. 2014). Differences in sensitivity between strains

may lead to false conclusions and interpretations when studying the potential adverse effects of

estrogenic compounds. Therefore, the selection of strain needs to be carefully considered for all

studies that use potential or known estrogenic compounds. Table 1 summarizes several strain-

dependent endpoints with differences in response to estrogenic compound exposure.

14  

Table 1. Summary of Studies Examining Sensitivity of Rodent Strains to Estrogenic Compounds

Endpoint Estrogenic Compound

Summary of Results Citation

Uterine Weight

DES ↑ in Alpk and C57Bl/6J at low and high dose, ↑ in CD1 and B6CBF1 at high dose only Ashby et al. (2003)

EE ↑ in DA/Han, Wistar, and Sprague-Dawley

Diel et al. (2004) Genistein ↑ in DA/Han, Wistar, and Sprague-Dawley at 50 and 100 mg/kg/day

BPA ↑ in Da/Han and Sprague-Dawley at 200 mg/kg/day

Uterine Morphology

Estradiol Benzoate cystic, hyperplastic glands in C57Bl/6 one week after treatment; Silberberg and Silberberg

(1951) develop two months after treatment in strain A/J

Circulating Sex Hormones

larger prolactin and P peaks in strains with small litter size; DeLeon et al. (1990)

smaller P peak in strains with large litter size

Epithelial ERα Expression

expressed in CD1 at PND3; expressed in BALB/c and C57Bl/6J at PND5

Bigsby et al. (1990), Taguchi et al. (1988)

DES advanced ERα ontogeny in C57Bl/6J Taguchi et al. (1988)

QTL

E2 Est2 and Est3control phenotypic variation in uterine weight Wall et al. (2013), Roper et

al. (1999)

E2 Est1 control phenotypic variation in eosinophil infiltration Griffith et al. (1997)

E2/DES Eutr1 (E2) and Eutr2 (DES) control phenotypic variation in pyometra development Gould et al. (2005), Pandey

et al. (2005)

Ovary

number of oocytes in neonatal development higher in CD1 than C57Bl/6;

Pepling et al. (2010) CD1 faster rate of loss of oocytes

E2 ↑ in oocytes in CD1 and C57Bl/6; greater increase in C57Bl/6

BPA ↓ follicular growth and steroidogenesis in CD1 and C57Bl/6 Peretz and Flaws (2013),

Peretz et al. (2013)

Vagina

BPA ↑ cell proliferation in F344 compared to Sprague-Dawley Long et al. (2000)

Estradiol Valerate C57Bl/6 and BALB/c susceptible and CD1 resistant to Candida albicans infection Mosci et al. (2013),

Clemons et al. (2004), Calderon et al. (2003)

Anterior Pituitary BPA hyperprolactinema in F344, no effect in Sprague-Dawley Steinmetz et al. (1997)

Mammary BPA/E2 changes in morphology in CD1 and C57Bl/6N Wadia et al. (2007)

Male Reproductive

E2 ↓ testicular weight and spermatogenesis in C57Bl/6J, CD1 resistant Spearow et al. (2001),

Spearow (1999)

15  

1.6.1 Susceptibility to Uterine Weight Increases and Morphological Alterations

The uterotrophic assay is often used to determine the estrogen-like activity of a

compound. However, differences in sensitivity to estrogens between rodent strains may lead to

false results. For example, when comparing uterine weights in four DES-exposed mouse strains,

the Alpk and C57Bl/6J strains demonstrated significant increases at 1 µg/kg/day DES, while the

CD1 and B6CBF1 strains did not. All strains had significant uterine weight increases at 10

µg/kg/day DES though (Ashby et al. 2003). Several other studies using mice or rats observed

varied susceptibility to uterine weight increases in response to EE, BPA, DES, genistein, and

other E2 derivatives (Diel et al. 2004, Greenman et al. 1977, Drasher 1955).

Morphological features of the glands in the endometrium of the uterus differed between

strains in response to estradiol benzoate treatment. The C57Bl/6 strain had cystic and

hyperplastic glands one week after treatment, while it took two months before the same altered

morphology was observed in the A/J strain (Silberberg and Silberberg 1951). These findings

suggest that differences in sensitivity to estrogenic compounds impact alterations in uterine

morphology, eventually developing into pathology.

1.6.2 Differences in Circulating Sex Hormones and ERα Levels

Strain-dependent differences in circulating sex hormone levels may influence variances

in sensitivity to exogenous estrogenic compounds. Mouse strains selected for small or large litter

size had differing circulating sex hormone levels. Smaller litters coincided with larger prolactin

and P peaks, with the prolactin peak occurring in estrus. Larger litter size correlated with a

smaller P peak during proestrus (DeLeon et al. 1990). It was also possible that E2 levels differed

between strains due to alterations in circulating P levels and negative feedback mechanisms.

16  

Uterine epithelial ERα expression during development also differed greatly between

strains. The CD1 strain expressed ERα as early as PND3, compared to BALB/c or C57Bl/6J

mice that expressed ERα at PND5 (Bigsby et al. 1990, Taguchi et al. 1988). Estradiol is

necessary for epithelial cell proliferation, and this finding suggests that CD1 mice have a more

advanced glandular and luminal ontogeny compared to inbred strains. However, DES exposure

advanced ERα ontogeny in epithelial cells of C57Bl/6J mice so that the uterus looked similar to

the uterus of the CD1 strain (Taguchi et al. 1988).

1.6.3 Quantitative Trait Loci Controlling Phenotypic Variation

Genetic variation between strains or even substrains influences phenotypic differences at

numerous endpoints. The two most commonly used inbred mouse strains, C57Bl/6J and

C57Bl/6N, have distinct genetic differences that led to phenotypic alterations at endpoints related

to behavior, ophthalmology, immune function, cardiovascular parameters, and metabolism

(Simon et al. 2013, Mekada et al. 2009). Specific quantitative trait loci (QTL) regulated

susceptibility to alterations in response to estrogenic compound exposure for uterine wet weight,

cellular infiltration, and the development of pyometra. Phenotypic variation in uterine wet weight

was controlled by two interacting QTL in the mouse, Est2 on chromosome 5 and Est3 on

chromosome 11. A third QTL, Est4 on chromosome 10, encoded an interacting factor that

influenced the variation of Est2 and Est3. These QTL contained several E2-responsive genes,

leading to the possibility that estrogenic EDCs may also alter uterine weight via interactions with

these genes (Wall et al. 2013, Roper et al. 1999). Eosinophil infiltration in response to E2

treatment also appeared to be genetically controlled by Est1 on chromosome 4, which interacted

with loci on chromosomes 10 and 16 (Griffith et al. 1997). Two separate but overlapping QTL

on chromosome 5 were responsible for controlling the development of pyometra in rats, Eutr1

17  

for E2-induced pyometra and Eutr2 for DES-induced pyometra (Gould et al. 2005, Pandey et al.

2005). These studies highlighted that sensitivity to estrogenic compounds was in part regulated

by underlying genetic differences between strains due to specific chromosomal locations being

identified and involved in genetic control of estrogen-responsive endpoints.

1.6.4 Differential Sensitivity of the Ovary and Vagina to Estrogenic Compounds

Other organs in the female reproductive system, including the ovaries and vagina, also

showed physiological differences or alterations in susceptibility to estrogenic compounds

between rodent strains. During normal neonatal ovarian development, the number of oocytes in

the ovaries of CD1 mice was significantly higher compared to the C57Bl/6 strain. However, the

rate of loss of oocytes in the ovaries of CD1 mice was faster compared to the C57Bl/6 strain. The

same number of oocytes were present at PND4 in the CD1 and C57Bl/6 strains (Pepling et al.

2010). When comparing sensitivity to E2, both the CD1 and C57Bl/6 strain had a significantly

increased number of oocytes compared to non-treated conditions. However, the increase was

much greater in C57Bl/6 mice, suggesting that this strain is more sensitive to estrogenic

compounds (Pepling et al. 2010). Inhibited follicle growth and steroidogenesis in response to

BPA exposure in the adult ovary occurred in both CD1 and C57Bl/6 strains, suggesting that the

response of the ovary to BPA at this endpoint was not dependent on genetic variation between

mouse strains (Peretz et al. 2013, Peretz and Flaws 2013).

Another estrogen-responsive tissue, the vagina, showed differences in cell proliferation

but not BPA clearance, ERα binding, or c-fos mRNA expression levels between Fischer 344 and

Sprague-Dawley rats (Long et al. 2000). Infection with Candida albicans also differed between

C57Bl/6, BALB/c, and CD1 and derived CD10 and CD3 strains, following similar patterns of

18  

susceptibility as to estrogen sensitivity (Mosci et al. 2013, Clemons et al. 2004, Calderon et al.

2003).

1.6.5 Differential Sensitivity of Other Tissues to Estrogenic Compounds

Tissues other than those in the female reproductive tract also exhibited differential

sensitivity to estrogens based on rodent strain. In the anterior pituitary, BPA exposure resulted in

hyperprolactinemia in Fischer 344 rats but did not affect the Sprague-Dawley strain (Steinmetz

et al. 1997). When examining the effects of BPA exposure on mammary gland morphology, the

CD1 strain had slightly more pronounced alterations compared to C57Bl/6N mice; however,

both strains were sensitive to the actions of E2 on mammary gland morphology (Wadia et al.

2007). The male reproductive system also displayed phenotypic variations to E2 treatment in

different mouse strains. The CD1 strain was resistant to E2 effects on inhibition of testicular

weight and spermatogenesis at doses 16 fold higher than those that led to the abolishment of

spermatogenesis in the C57Bl/6J strain (Spearow et al. 2001, Spearow 1999), demonstrating that

estrogen sensitivity is not specific to females. Genetic background also played an important role

in estrogen sensitivity in the heart (Kararigas et al. 2014), the adrenal (Westberg et al. 1957),

macrophage function, (Passwell et al. 1974), and anxiety-related behavior and associated

inflammatory responses (Benatti et al. 2011).

Important knowledge gaps still remained concerning differences in sensitive populations

and BPA exposure. In regards to uterine structure, function, and potential for the development of

pathologies, it was unknown if genetic variation at least in part controlled phenotypic

differences. It was also unclear if baseline physiological differences within the uterus between

strains had any effect on sensitivity to the endocrine disrupting actions of BPA.

19  

1.7 The Endocrine System

The endocrine system is comprised of a series of interacting organs and secreted

hormones to regulate numerous biological processes and maintain an overall homeostasis in the

organism. Endocrine glands throughout the body produce and secrete hormones directly into the

bloodstream. These hormones then act on target cells, usually through binding to specific

receptors (Molina 2013). Hormones released by organs in the endocrine system affect almost any

tissue in the body; of particular interest to these studies is the uterus and response to estrogenic

compounds.

Often, hormone secretion is regulated by a feedback loop system to either increase

(positive feedback) or decrease (negative feedback) further hormone production and secretion.

An example of one such feedback loop is the hypothalamic-pituitary-gonadal (HPG) axis (Figure

2). In its simplest form, the hypothalamus produces and secretes gonadotropin-releasing

hormone (GnRH), stimulating the anterior pituitary to secrete luteinizing hormone (LH) and

follicle stimulating hormone (FSH). These hormones then stimulate the ovaries to produce

progesterone (P4) and 17β-estradiol (E2) in females or the testes to produce testosterone (T) in

males. These hormones then have regulatory roles in these reproductive organs, but they are also

secreted into circulation to play stimulatory roles in other target tissues, such as the uterus,

mammary, or prostate. Sex hormones also feed back to the hypothalamus and anterior pituitary

to work in an inhibitory fashion. High levels of E2 leads to decreased FSH secretion and

increased LH secretion, leading to the LH surge associated with ovulation. The presence of high

levels of P4 then decreases LH secretion (Molina 2013).

20  

 

Figure 2. Simplified representation of HPG axis and negative feedback loop. As described, GnRH released by the hypothalamus stimulates the anterior pituitary to release LH and FSH. These hormones stimulate the production of E2 and P4 in the ovaries and T is the testis, which then act on target tissues. However, the sex hormones can also inhibit further release of LH and FSH from the anterior pituitary.

1.8 Nuclear Hormone Receptors: ERα and ERβ

1.8.1 Structure

Estrogens and estrogenic compounds bind to specific nuclear hormone receptors, the

estrogen receptors ERα and ERβ, and act as a ligand-activated transcription factor to modulate

responsive gene expression through recruitment of stimulatory or inhibitory complexes.

Estrogens binding to ERs can also have rapid signaling effects. The most potent endogenous

estrogen is E2. Experimental evidence in support of the idea of a receptor specific for E2 was

first reported by Jensen and Jacobson (1962), but the cDNA and amino acid sequences of the

human ER were not reported until over 20 years later (Green et al. 1986, Greene et al. 1986). In

21  

1996, a second novel ER was cloned and named ERβ or ESR2 to distinguish it from the previous

ER, now named ERα or ESR1 (Kuiper et al. 1996, Mosselman et al. 1996). Using BioGPS

software, the highest mRNA expression of human ESR1 occurs in the uterus, pituitary, and

prostate. Highest mRNA expression of human ESR2 occurs in cardiomyocytes (Figure 3) (Wu et

al. 2013, Wu et al. 2009). The chromosomal location of ESR1 in humans is 6q25.1,

corresponding to a chromosomal location for Esr1 of 10 2.03 cM in mice. ESR2 in humans is

located on 14q22-24, corresponding to the location of Esr2 in mice on 12 33.52 cM.

Figure 3. Expression of ERα and β in human tissues. Shown are graphical representations of mRNA expression levels in various human tissues for ERα and ERβ. The graph for ERα is also shown zoomed in to show lower expression levels in numerous tissues. Graphs were generated at biogps.org using Dataset GeneAtlas U133A, gcrma for both receptors and Probeset 205225_at for ERα and Probeset 211120_x_at for ERβ.

The ERα and ERβ proteins are members of the nuclear hormone receptor (NHR)

superfamily. All NHRs are made up of three interacting domains, the A/B or N-terminal domain

(NTD) containing a ligand-independent activation factor (AF1), the C or DNA-binding domain

(DBD), and the D/E/F or ligand-binding domain (LBD) containing a ligand-dependent activation

22  

factor (AF2) important for coregulator recruitment and receptor dimerization (Figure 4). The

DBD contains two zinc finger domains that facilitate dimerization and DNA binding of the

receptor to specific palindromic estrogen response elements (EREs) in a DNA sequence. The D

region in the LBD is a hinge region which contains a nuclear translocation signal. The LBD

contains a hydrophobic ligand binding pocket, important for ligand specificity (Planey et al.

2014, Ellmann et al. 2009).

Figure 4. Schematic of domain structure of ERα and β. Each receptor contains an A/B or N-terminal domain (NTD), a C or DNA-binding domain (DBD), and a D/E/F or ligand-binding domain (LBD). Sequence homologies between the human receptors for each domain are noted. 1.8.2 Nuclear Translocation, Membrane-Initiated, and Rapid Signaling Mechanisms

The mechanism of ER-mediated nuclear translocation begins with free E2 diffusing

through the plasma membrane and binding to an ER/heat shock protein (HSP) complex, made up

of HSP56 and 90, in the cytosol. Once E2 binds to the ER, the HSP complex dissociates and two

E2/ER molecules dimerize. The dimerized product then enters the nucleus, binds to EREs in the

DNA sequence, and recruits primary transcription factor coregulators which can then recruit

secondary coregulators to either promote or repress transcription of estrogen-responsive genes

(Figure 5) (Belcher 2008).

23  

Figure 5. Classical model of nuclear hormone receptor translocation. Shown is a cartoon describing the “classical” nuclear receptor mechanism of ligand-bound estrogen receptor (ER)-mediated effects on gene transcription. Ligand binding induces a conformational change of the receptor leading to dissociation from a chaperone complex containing heat shock proteins. Activated receptors dimerize and bind at estrogen response elements (EREs) in the promoters of E2-responsive genes. Ligand/receptor complexes bound to EREs results in the recruitment of coregulatory transcription factors to the promoter which modify target gene transcription (Belcher 2008).

Rapid signaling (non-genomic) mechanisms of ERα and ERβ activation can also occur

(Banerjee et al. 2014, Levin 2009, Belcher 2008). Rapid signaling through the ERs can either be

membrane-initiated or occur intracellularly. Activation of ERα increased phosphoinositide-3-

kinase (PI3K), extracellular signal-regulated kinases (ERK) 1/2 mitogen-activated protein

kinases (MAPK), and protein kinase B (PKB)/Akt through a G protein-coupled mechanism

(Banerjee et al. 2014, Kumar et al. 2007, Revankar et al. 2005, Razandi et al. 2003b). In the

cerebellum and cerebellar granule cell neurons, E2-induced neurotoxicity was mediated by rapid

24  

signaling actions of ERβ, involving a G protein- and protein kinase A (PKA)-dependent

mechanism. Treatment with E2 led to rapid activation of ERK1/2 MAPK and stimulated cell

death (Le and Belcher 2010, Belcher et al. 2005, Wong et al. 2003). In the female heart,

estrogens rapidly altered myocyte Ca2+ handling, leading to the promotion of arrythmogenesis

(Yan et al. 2011). These rapid signaling actions were mediated by an imbalance between the

stimulatory effects of ERβ and the suppressing actions of ERα (Belcher et al. 2012). The

protective effects of E2 against bone resorption may be due to rapid signaling actions as well,

based on results from studies using an E2 dendrimer conjugate incapable of inducing the nuclear

actions of ERα (Bartell et al. 2013). Another physiological action of ERα-mediated rapid

signaling is the increased production of nitric oxide (NO) within minutes of E2 treatment due to

activation of endothelial nitric oxide synthase (eNOS), which plays a crucial role in maintaining

vascular homeostatis and function (Wu et al. 2011, Wyckoff et al. 2001, Chen et al. 1999,

Lantin-Hermoso et al. 1997).

Because ERs are located intracellularly in the cytoplasm, recruitment to the membrane is

often necessary for these rapid signaling actions to occur. One such process is palmitoylation, or

the covalent attachment of fatty acids to cysteine residues (Meitzen et al. 2013). A Cys447Ala

mutation in ERα impaired localization to the plasma membrane, indicating Cys447 as the crucial

site for palmitoylation (Acconcia et al. 2005, Acconcia et al. 2004). A second palmitoylation site,

Cys451, was also implicated in the rapid signaling mechanisms of ERα (Adlanmerini et al.

2014). Recruitment of ERα to the plasma membrane also likely occurred through its interaction

with c-Src or caveolin-1 (Banerjee et al. 2014, Haynes et al. 2003, Razandi et al. 2003a).

A third membrane-associated G protein-coupled receptor that binds E2 and is expressed

in several tissue types is G protein-coupled receptor 30 (GPR30) or G protein-coupled estrogen

25  

receptor 1 (GPER1) (Carmeci et al. 1997, Kvingedal and Smeland 1997, Takada et al. 1997,

Owman et al. 1996). Activation of GPR30 by E2 induced rapid signaling pathways, usually

within minutes. GPR30 and ERα both activated ERK1/2 via epidermal growth factor receptor

(EGFR) transactivation (Razandi et al. 2003b, Filardo et al. 2000). While both receptors also

activate PI3K (Revankar et al. 2005), most cell types utilized GPR30 in vivo to activate PI3K

and MAPK (Prossnitz and Maggiolini 2009, Revankar et al. 2005). Another rapid signaling

mechanism was intracellular Ca2+ mobilization via activation of GPR30 (Revankar et al. 2005).

While GPR30 is not a nuclear hormone receptor, GPR30-mediated events still indirectly led to

transcriptional activation as well (Prossnitz and Maggiolini 2009). For example, several cell

types had increased expression of c-fos and Bcl-2 through E2-mediated GPR30 activation,

leading to increased proliferation (Prossnitz and Barton 2009, Prossnitz and Maggiolini 2009). It

was suggested that GPR30 signaling played a role in tumor proliferation and invasiveness in

breast, endometrial, and ovarian cancers and indicated poor survival (Smith et al. 2009, Smith et

al. 2007, Filardo et al. 2006, Filardo et al. 2002, Filardo et al. 2000).

The estrogen-related receptors (ERRs) are members of the NHR superfamily discovered

through a cDNA screening and have a similar sequence in the DBD as the human ERα (Giguere

et al. 1988). There are currently three types of human ERRs, named ERRα (ESRRA), ERRβ

(ESRRB), and ERRγ (ESRRG). These receptors are considered orphan receptors because the

endogenous ligand is unknown. They have a low sequence homology in the LBD with ERα;

therefore, they do not bind or respond to E2 (Huss et al. 2015). However, ERRs are

constitutively active based on dimerization partners and also bind exogenous estrogenic

compounds such as BPA or DES (Liu et al. 2014, Matsushima et al. 2007, Huppunen and

Aarnisalo 2004). Similar to ERs, ERRs bind to specific DNA sequences named estrogen-related

26  

receptor response elements (ERREs) as monomers, homodimers, or heterodimers (Huppunen and

Aarnisalo 2004, Gearhart et al. 2003). ERRs do not bind strongly to EREs; however, ERRs and

ERα do share some target genes and ERRs can modulate estrogen signaling by cooperation for

full gene activation or by antagonism (Zhang et al. 2006, Kraus et al. 2002, Vanacker et al. 1999,

Yang et al. 1996). Crystal structures showed that ERRs complex with a coactivator peptide from

peroxisome proliferator-activated receptor coactivator-1α (PGC-1α) and had a transcriptionally

active conformation in the absence of ligand (Kallen et al. 2004), suggesting that ligand binding

is not necessary for ERRs to bind to DNA and regulate transcription. Signaling through ERRs

was important in regulation of genes involved in oxidative phosphorylation, mitochondrial

biogenesis, fatty acid transport, and bone and cartilage formation. They were also involved in

regulating the stress response and the development of breast and prostate cancers (Misawa and

Inoue 2015, Bonnelye and Aubin 2013, Byerly et al. 2013, Huss et al. 2004, Schreiber et al.

2004).

1.8.3 Role of ERs in Uterine Structure and Reproductive Function

Estrogen receptors play a critical role in regulating numerous processes in the uterus.

However, of most importance to uterine structure and function is ERα. Studies using global ERα

knockout or uterine epithelial cell-specific ERα knockout models showed that ERα is required

for fertility because implantation and decidualization cannot occur without activation of the

receptor (Pawar et al. 2015, Hewitt and Korach 2003, Hewitt et al. 2002).

1.8.3.1 Postnatal Uterine Development

Expression and localization of ERα changed in the developing uterus after birth. Initially,

ERα was located in the stroma at low expression levels which progressively increased during

development. In rats, a gradual change in localization occurred during development, with ERα

27  

initially expressed in the luminal epithelium at postnatal day (PND) 7 but then most abundant in

the glandular epithelium at PND21 (Fishman et al. 1996). These changes in localization and

increases in expression occurred until puberty and the beginning of estrous cyclicity, at which

time their localization and expression was similar to an adult rodent.

Adenogenesis, or uterine gland development, is another process that required ERα

(Lubahn et al. 1993). Glands were not present in the uterus at birth in laboratory rodents or

humans. Starting at PND5 in rodents, epithelial invaginations in the uterus were observed that

formed glandular epithelium (Cooke et al. 2012a, Cooke et al. 2012b). Glands were not present

in the uterus until PND7 in mice and PND9 in rats (Cooke et al. 2013, Branham et al. 1985). On

PND15 in mice, a general organization of the adult uterus was established (Hu et al. 2004). In

humans, glandular epithelium differentiated from luminal epithelium to form tubular glands

throughout the stroma (Cooke et al. 2013).

1.8.3.2 Impact on Gene Expression

Expression of several genes in the uterus is dependent on ERα signaling. In the absence

of ligand, ERα still bound to chromatin structures, but ERα binding was greatly increased by the

presence of E2. Often, RNA polymerase II was located in similar binding sites, suggesting co-

localization in preparation for transcriptional activation (Hewitt et al. 2012). Over 1800 genes

were found to be regulated by E2 in the uterus (Leitman et al. 2012). In the cycling uterus,

clusters of genes involved in extracellular matrix (ECM), cell adhesion, proliferation, immune

response, coagulation, metabolism, and Wnt and hedgehog signaling were differentially

expressed in stages of the reproductive cycle, suggesting that their expression is dependent in

part on E2 activation of ERs (Yip et al. 2013).

28  

1.8.3.3 ERα in the Brain and Impact on Cyclicity

In the brain, ERα also influences uterine function through responses to circulating E2

levels and negative feedback mechanisms. ERα-expressing neurons in the arcuate nucleus were

involved in this negative feedback and normal cyclicity (Yeo and Herbison 2014). Inhibition of

GnRH release from GnRH neurons occurred through E2 binding to ERs (Radovick et al. 2012,

Sanchez-Criado et al. 2006). Using in situ hybridization, expression of kisspeptin (Kiss1) and

ERα in the hypothalamus was highly overlapping. Kiss1 is a protein that can directly regulate

GnRH release from GnRH neurons due to binding to and activating GPR54 located on GnRH

neurons (Navarro et al. 2004). Regulation of GnRH neurons by Kiss1 significantly impacted

estrous cyclicity and puberty by regulating LH and FSH expression (Witham et al. 2013, Navarro

et al. 2004). Other studies supported that regulation of GnRH release was mediated by E2

binding to ERα in Kiss1 neurons (Radovick et al. 2012, Cao and Patisaul 2011, Vida et al. 2010).

Because ERα expression and signaling in the brain was highly involved in regulating estrous

cyclicity, alterations in this signaling significantly impaired uterine function. Administration of

exogenous estrogenic compounds during neonatal development significantly decreased Kiss1

neuron density and signaling in the hypothalamus, leading to impaired reproductive function

(Patisaul et al. 2009, Bateman and Patisaul 2008).

1.8.3.4 Role of GPR30 in the Uterus

GPR30 is expressed in both the endometrium and myometrium of the uterus, with

expression levels varying during the reproductive cycle (Maiti et al. 2011, Kolkova et al. 2010,

Otto et al. 2009). However, GPR30 had little effect on uterine structure and fertility based on

knockout studies (Otto et al. 2009). In the endometrium, GPR30 activation negatively regulated

rapid signaling of ERα by inhibiting ERK1/2 phosphorylation and reducing uterine growth,

29  

suggesting that these signaling events are relevant for balancing ERα-mediated effects (Gao et al.

2011). Within the myometrium, GPR30 activation increased smooth muscle contraction in

response to oxytocin, implying an important role in labor (Maiti et al. 2011).

Progression of several uterine diseases is partially mediated by GPR30. In leiomyoma

samples, GPR30 expression was significantly increased compared to normal myometrium (Tian

et al. 2013). Proliferation and invasion of endometrial carcinoma cells occurred via activation of

GPR30 by E2 or hydroxytamoxifen and downstream signaling of the ERK/MAPK pathway (Du

et al. 2012, He et al. 2009, Vivacqua et al. 2006).

1.9 The Cycling and Pregnant Uterus

1.9.1 Circulating Hormones and Tissue Remodeling

The main function of the uterus is to protect and provide nourishment for a growing fetus.

The myometrium and endometrium are two distinct morphological and functional tissue layers.

The myometrium is divided into two layers composed of smooth muscle and vasculature and

surrounds the endometrium to provide structural support and protection. It can greatly expand

during pregnancy and contracts during labor (Thompson 2015). The endometrium consists of

lumen and gland structures lined by epithelial cells, which are connected by the lamina propria or

stroma and ECM (Figure 6). These structures are crucial for preparing the uterus for implantation

and fetal development and differentiate based on circulating E2 and P levels, also known as the

endometrial or menstrual cycle in humans and the estrous cycle in rodents (Buy and Ghossain

2013, Molina 2013).  

30  

 

Figure 6. Normal uterine structure in adult CD1 mouse. Uterus is divided into myometrium (Myo) and endometrium (Endo). Myo consists of inner myometrium (IM) and the outer myometrium (OM). Endo is made up of luminal and glandular structures and connected by stroma. Similar uterine structures are found in normal human tissue.

In humans, the entire reproductive cycle lasts on average 28 days, beginning with the

endometrium shedding in a process called menstruation. As described in Molina (2013),

menstruation occurs in the menstrual phase of the uterus if fertilization and implantation does not

occur. The menstrual phase is considered day 1 of the cycle. Menstruation also indicates day 1 of

the follicular phase, or maturation of the developing egg, in the ovary. During the beginning of

the follicular phase, sex hormone levels are low. Starting at day 3 of the follicular phase, E2

levels begin to gradually increase, peaking at day 13. The proliferative phase in the uterus occurs

between days 4 to 13 and is characterized by thickening of the endometrium in preparation for

implantation. Around day 14, ovulation occurs due to a rapid increase in LH and FSH and a peak

in E2 levels, signifying the end of the follicular phase. Following ovulation, the ovary enters the

luteal phase, in which the follicle that released the egg gradually matures to form a corpus

luteum. The luteal phase lasts from day 14 to day 26. A mature corpus luteum secretes P4,

31  

leading to the increased P4 levels observed after ovulation. Levels of LH, FSH, and E2 rapidly

decrease; however, a second, smaller peak in E2 is also observed during high P4 levels around

day 20. In the uterus, the secretory phase occurs from day 13 to day 24 and is characterized by an

inhibition of endometrial thickening by high P4 levels (Molina 2013, Hinson et al. 2010).

Morphological changes prepare the uterus for implantation, a process called decidualization

(Gardner and Shoback 2007). If implantation has not occurred at day 28, the uterus reenters the

menstrual phase and the thickened endometrium is shed (Molina 2013, Hinson et al. 2010).

Graphical representation of the human menstrual cycle is found in Figure 7.

Figure 7. Simplified representation of the human menstrual cycle. The menstrual cycle has an ovarian and endometrial component. The ovarian cycle is divided into the follicular and luteal phases. The endometrial cycle is divided into the menstrual (Mens), proliferative and secretory phases. The relative sex hormone levels are also graphically shown from Molina (2013).

A similar process occurs in rodents, but on a shorter time scale. While the menstrual

cycle in humans is roughly 28 days, the estrous cycle is only 4 days in rodents (Cora et al. 2015,

Goldman et al. 2007). The proestrus and estrus stages are similar to the proliferative phase in

humans in that the endometrium is thickening in preparation for implantation. Ovulation occurs

32  

at the end of the estrus stage. However, peaks in E2, LH, and FSH do not occur at the same time.

Instead, all three hormones peak between the proestrus and estrus stages, with FSH having a

second, smaller peak at the end of the estrus stage (McLean et al. 2012). Following ovulation, the

rodent enters metestrus if fertilization has not occurred. This stage is characterized by

endometrial degeneration and is similar to the luteal phase in humans (Hedrich and Bullock

2004). However, instead of menstruation like in humans, rodent tissue degenerates and

eventually regenerates. A second small peak in E2 occurs in metestrus, as well as gradual

increases in P4 that peaks in diestrus. The diestrus stage is characterized by a quiescent state in

the endometrium (McLean et al. 2012).

1.9.2 Role of Extracellular Matrix in Uterine Structure and Function

The extracellular matrix (ECM) is a group of interacting proteins and polysaccharides

that are secreted by surrounding cells to provide structural support and initiate biochemical and

biomechanical processes (Frantz et al. 2010). The ECM plays a dynamic and tightly regulated

role in maintaining structure of the cycling uterus. Regulation of collagen synthesis and

degradation is summarized in Figure 8. In the cycling uterus, expression of ECM proteins

involves several levels of regulation, almost all of which are differentially expressed in response

to circulating levels of E2 or P, highlighting that sex hormones are important for numerous

aspects of tissue remodeling during the menstrual or estrous cycles.

Accumulation of collagens involves a balance between hormonal control of collagen

gene expression and matrix metalloproteinase (MMP) expression and activity (Gaide

Chevronnay et al. 2012, Henriet et al. 2012). The collagen types I and III are primary collagens

expressed in mammalian uterus, with collagen types IV, V, and VI expressed variably during the

cycle, mainly in the stroma, basement membranes of glandular epithelium, and myometrium

33  

(Augsburger and Henzi 2008, Boos 2000, Tanaka et al. 1998, Stenback 1989, Aplin et al. 1988,

Karkavelas et al. 1988). In response to E2, synthesis of mRNA and protein for collagen subtypes

increased (Frankel et al. 1988, Dyer et al. 1980, Salvador and Tsai 1973).

Figure 8. Regulation of collagen synthesis and degradation. Shown is a simplified schematic of the major levels of control of transcriptional and posttranslational regulation of collagens and MMPs. In stromal and epithelial cells in the uterus, E2 binds to ERα, dimerizes, and translocates to the nucleus. Binding to EREs and recruitment of co-activators leads to transcription of collagen type 1 (pink) and type 3 (green) and proMMP2 (blue) and proMMP14 (purple). Once collagen mRNA is translated, it is released into the extracellular compartment. ProMMP14 is activated and localized to the plasma membrane, where it forms a complex with TIMP2 (orange). This complex interacts with proMMP2 to activate the enzyme via a cysteine-switch mechanism, which is then released into the extracellular compartment. MMP2 can be inhibited by TIMPs and α-macroglobulin.

Matrix metalloproteinases are a group of Ca2+-dependent collagen degrading enzymes

whose expression and activity are also tightly regulated by E2 levels during the menstrual or

estrous cycle (Gaide Chevronnay et al. 2012, Russo et al. 2009, Helvering et al. 2005, Rodgers et

al. 1994). They are divided into different subgroups based on their structure (Figure 9). For

34  

example, collagenases (MMP1, 8, and 13) differ from gelatinases (MMP2, 9) due to the lack of

fibronectin type II domains located within the catalytic domain. Substrates for each MMP also

differ. However, most collagens can be degraded by several different MMPs. For example,

collagen type I is a substrate for and degraded by MMP1, 2, 7, 8, 13, and 14 (Curry and Osteen

2003). The MMP9 and MMP2 proteins are key gelatinases present in both latent and active

forms in endometrial tissue of the uterus throughout the reproductive cycle (Gaide Chevronnay

et al. 2012). Other MMPs in the uterus include MMP1, 3, 7, 10, 11 and various membrane-

associated MMPs, which also have differential expression during the estrous or menstrual cycle

(Goffin et al. 2003, Goldman and Shalev 2003, Curry and Osteen 2001, Hulboy et al. 1997).

Initially, MMPs are transcribed and translated as quiescent proMMPs and activated in the

extracellular space by proteolytic cleavage. Regulation of MMP2 activity during the reproductive

cycle is in part controlled by MMP14, which activates proMMP2 by proteolytic cleavage via a

“cysteine switch” mechanism of action. A cysteine residue in the propeptide domain interacts

with Zn2+ in the catalytic domain, preventing cleavage of the propeptide domain and activation

of the MMP. Disruption of this interaction leads to opening of the cleavage site in the propeptide

domain and the active site in the catalytic domain and activation of the MMP (Gaide Chevronnay

et al. 2012, Murphy et al. 1999, Strongin et al. 1995, Sato et al. 1994, Springman et al. 1990, Van

Wart and Birkedal-Hansen 1990).

35  

Figure 9. Schematic representation of MMP structures. The common name and associated number for MMPs are shown. The MMPs contain several domains, including a signal peptide, propeptide, catalytic domain containing a Zn2+ binding site, hinge region, and hemopexin-like domain. The MMP is activated by cleavage of the signal peptide and propeptide. The gelatinases contain several fibronectin type II domains within the catalytic domain. A furin site that allows for intracellular activation can be found in MMP11 and the transmembrane MMPs. The transmembrane MMPs also contain a transmembrane linker and a cytoplasmic domain. The activity of MMPs is also tightly controlled and is inhibited by P4 or specific

inhibitory proteins. Attenuation of MMP14 expression by P4 led to decreased MMP2 activation

(Zhang et al. 2000). Tissue inhibitors of metalloproteinases (TIMPs) and α-macroglobulin (α-

MG) also inhibit MMPs in active form (Dong et al. 2002). Based on crystal structures, TIMPs

have an overall “wedge-like” shape that fits into the active site of MMPs, inhibiting ECM

components from being degraded (Nagase et al. 2006, Fernandez-Catalan et al. 1998, Gomis-

36  

Ruth et al. 1997). Inhibition by α-MG occurred by entrapping the MMP between the four

subunits of one α-MG molecule. The MMP/α-MG complex was then cleared by endocytosis

mediated by low density lipoprotein receptor-related protein (LRP) (Nagase et al. 2006,

Strickland et al. 1990). TIMPs-1, -2, and -3 were differentially expressed during the estrous or

menstrual cycle in response to cycling levels of sex hormones, increasing and decreasing at

similar time points to MMPs (Gaide Chevronnay et al. 2012, Goffin et al. 2003, Zhang and

Salamonsen 1997, Hampton and Salamonsen 1994, Sato et al. 1991). The fluctuating expression

levels prevented excess degradation of collagens and other ECM components by high levels of

MMPs. However, TIMP-2 also aided in activation of proMMP2 to MMP2, binding to proMMP2

and sequestering the quiescent protein near MMP14 (Butler et al. 1998). Overall, several levels

of regulation exist that control collagen synthesis and degradation in the cycling uterus.

1.9.3 Role of Extracellular Matrix during Pregnancy

Components of the ECM are highly involved in the various processes of a successful

pregnancy, including decidualization, implantation, placentation, and involution (Figure 10).

Serious complications or pregnancy failure may arise from aberrant ECM expression or activity

(Plaks et al. 2013, Singh et al. 2013, Pereza et al. 2012). During decidualization, endometrial

fibroblasts differentiated into decidual cells and denoted potential implantation sites

(Abrahamsohn and Zorn 1993). Surrounding these sites were collagen fibrils organized into a

basket-like structure comprised of collagens I, III, V, and VI (Carbone et al. 2006, Dziadek et al.

1995). Collagen IV and laminin were also expressed in the pericellular region of stromal cells in

the decidua (Iwahashi et al. 1996). Collagen diameter and thickness were greatly increased in the

decidua, while the collagen fibrils in the normal endometrium were unaffected (Spiess et al.

2007, Spiess and Zorn 2007, Teodoro et al. 2003, Iwahashi et al. 1996, Zorn et al. 1986).

37  

The overall amount of collagen in the decidua was increased, particularly collagen V,

which was expressed exclusively in the decidua. In humans, the ratio of collagen III/collagen I

was decreased and the ratio of collagen V/collagen I was significantly increased in the decidua

compared to normal endometrial tissue. Collagen V played a vital role in successful

decidualization and implantation, since a low collagen V/collagen I ratio was found in decidual

tissues in which a spontaneous abortion occurred (Iwahashi and Nakano 1998). It was

hypothesized that the ECM greatly increases in thickness and amount in order to reinforce the

decidual cells as they support the implanting and developing embryos, as well as forming a

barrier to prevent the invasion of immune cells and subsequent rejection of the embryo (Carbone

et al. 2006). The process of placentation and trophoblast invasion also involved alterations in

collagen accumulation, mainly collagens I, III, and IV (Kitaoka et al. 1996, Kitaoka et al. 1994).

During involution, the total amount of collagen rapidly decreased, reaching nulliparous levels by

day 5 in rats and 1-2 weeks in humans (Morrione and Seifter 1962, Harkness and Moralee 1956).

Figure 10. Schematic of decidualization and implantation in the human uterus. The embryo attaches to the luminal epithelium (LE) and begins pushing through the basement membrane (BM) to invade the underlying stroma. Simultaneously, stromal cells are differentiating into decidualized cells. Part of the trophoblast eventually attaches as the placenta. This figure is adapted from Sharkey and Macklon (2013) with the addition of labels denoting stages of embryo implantation and invasion.

38  

During decidualization, endometrial decidual cells exhibited an upregulation of

expression of TIMPs (Nuttall and Kennedy 1999), potentially to inhibit degradation of collagen

surrounding the implantation sites by the increased expression of MMP2 (Tsai et al. 2009).

Implantation and placentation occurred due to a balance between expression and activity of

MMPs and TIMPs. In order to enable invasion of trophoblast cells through the decidua, MMP2

and 9 degraded collagen IV in the basement membrane (Staun-Ram and Shalev 2005).

Expression of MMP2 and 9 and TIMPs was also found at implantation sites in the uterus and in

pre-implantation embryos (Fontana et al. 2012, Wang et al. 2003, Das et al. 1997). Co-

localization of MMP2, MMP14, and collagen IV mRNA was found in invading trophoblast cells,

as well as co-localization of MMP2 and MMP14 mRNA in decidual cells (Bjorn et al. 1997).

Human chorionic gonadotropin (hCG), a hormone indicative of pregnancy released by the

trophoblast, increased MMP2 and 9 secretion from trophoblast cells and decreased TIMP-1, -2,

and -3 expression in endometrial stromal cells to aid in trophoblast invasion and placentation

(Fluhr et al. 2008). During involution in the human uterus, the overall collagen content

significantly decreased (Morrione and Seifter 1962). Using a rodent model, mRNA expression of

Mmp2 and Mmp14 and activity of MMP2 were highly increased until five days postpartum. At

this time, Timp2 mRNA expression increased to inhibit MMP2 expression and activity (Manase

et al. 2006), which aid in explaining the significant decrease in collagen in the involuting human

uterus postpartum.

Preeclampsia, a serious and potentially life-threatening pregnancy complication, may be

promoted by aberrant expression of MMP2 and 9 to alter the ECM components of the

decidualized areas of the endometrium (Plaks et al. 2013, Lockwood et al. 2008). Increased

MMP9 and MMP2/TIMP-2 complex expression in serum or genetic polymorphisms in MMP2 or

39  

MMP9 gene promoters were correlated with spontaneous abortions (Nissi et al. 2013, Pereza et

al. 2012). Because of the associated increase in implantation failure, spontaneous abortions, or

pregnancy complications due to aberrant MMP expression, collagen and MMPs play distinct and

important roles in preparation of pregnancy, during pregnancy, and postpartum.

1.10 Uterine Disorders

1.10.1 Leiomyoma

1.10.1.1 Description, Symptoms, and Treatment of Leiomyoma

Leiomyoma, or uterine fibroids, are benign fibrotic growths in the myometrium that arise

from smooth muscle cells and are one of the most common benign pathologies to affect women

of reproductive age (Baird et al. 2003, Kempson and Hendrickson 2000). Some of the more

common symptoms in women with uterine fibroids are heavy and prolonged menstrual bleeding,

dysmenorrhea, pelvic pain or pressure, anemia, urinary incontinence, constipation, lower back

pain, compression of other pelvic organs, sexual dysfunction, infertility and recurrent pregnancy

loss (Commandeur et al. 2015). Treatment usually involves administration of GnRH agonists,

progestins, selective PR modulators, or oral contraceptives, implantation of levonorgestrel

intrauterine devices, or interventions such as myomectomy (removal of fibroids while preserving

the uterus), uterine artery embolization, and magnetic resonance (MR)-guided ultrasound

treatments. However, these treatments are often variable in effectiveness or ineffective all

together (Commandeur et al. 2015, Vilos et al. 2015, Gorny et al. 2014). In most instances,

hysterectomy is the only effective treatment for fibroids and is the most common cause for

hysterectomy in the United States (Commandeur et al. 2015, Vilos et al. 2015).

40  

1.10.1.2 Etiology of Leiomyoma

Hormonal and genetic factors play a role in the development of uterine fibroids. For

example, preadolescent and early adolescent and postmenopausal females have a low risk due to

the low circulating levels of sex hormones. Women taking oral contraceptives or that have had

multiple pregnancies also have a lower risk compared to normally cycling women (Commandeur

et al. 2015, Ross et al. 1986). Epidemiological studies have correlated having relatives with

leiomyoma as a risk factor for a woman to develop leiomyoma (Sato et al. 2002). Ethnicity also

plays a role, with African American woman developing fibroids more often and at a younger age

than Caucasian women (Baird et al. 2003).

Fibroids are clonal in origin and all cells in the mass arise from one cell, suggesting that

the parental cell has multipotent stem cell properties (Commandeur et al. 2015, Holdsworth-

Carson et al. 2014, Cai et al. 2007, Rogalla et al. 1995). Conditional knockout of β-catenin in the

embryo led to adipose gradually replacing smooth muscle cells in the Müllerian duct

mesenchyme (Arango et al. 2005). Cells in the uterus were immunopositive for α-smooth muscle

actin (α-SMA), ERα, and β-catenin and were located near cells positive for stem cell markers

and negative for several hematopoietic markers (Szotek et al. 2007). These studies provided

strong evidence that there exists a population of myometrial stem or progenitor cells, leading to

the theory that these stem/progenitor cells become dysregulated by a genetic mutation and

differentiate into actively dividing fibroid cells instead of smooth muscle cells (Bulun 2013). The

mediator complex subunit 12 (MED12) gene was mutated in over 70% of uterine fibroid samples

(Makinen et al. 2011), suggesting that mutations in MED12 may play a role in abnormal

differentiation of stem/progenitor cells.

41  

1.10.1.3 Regulation of Collagen Accumulation in Leiomyoma

The connection between fibrosis and alterations in collagen accumulation in uterine

fibroids is well established. During the proliferative phase, mRNA for collagen I and III was

overexpressed in uterine fibroids compared to surrounding tissue, suggesting that ECM

components are under hormonal control within the fibroid (Stewart et al. 1994, Brandon et al.

1993). This idea was supported by in vitro studies that treated normal myometrial and

leiomyoma cells with low levels of E2 and observed significant increases in collagen

biosynthesis and in vivo studies reporting that estrogen treatment promoted proliferation of

fibroids through the activation of fibroblasts (Luo et al. 2014, Zbucka et al. 2007). The

orientation and alignment of collagen fibrils in the fibroid were also altered, suggesting

differences in the amount of proteoglycan between the fibrils or alterations in the number of

cross links (Leppert et al. 2004). Expression of MMP2 and MMP14 and activity of MMP2 were

also increased in leiomyoma samples (Tsigkou et al. 2015, Wang et al. 2015, Bogusiewicz et al.

2007), potentially in an attempt to regulate the increased amount of collagen present in the

fibroid.

1.10.2 Endometriosis

1.10.2.1 Description, Symptoms, and Treatment of Endometriosis

Endometriosis is a benign estrogen-dependent gynecological condition characterized by

the presence and growth of ectopic endometrial glands and stroma, often associated with

inflammation and increased fibrosis. Ectopic tissue is most often found in the pelvic region or

peritoneum, but it is also found in the bowel, diaphragm, or pleural cavity. Prevalence of

endometriosis in the general population ranges from 6 to 10%; however, in women with

infertility or pain, prevalence increases to 35-50% (Giudice and Kao 2004). The most common

42  

symptom reported is chronic pelvic pain in 58% of women. Between 30 and 50% of women with

endometriosis are also infertile; however, the mechanism and causation are not well understood

(Hickey et al. 2014). Laparoscopy is the only tool to definitively diagnose a woman with

endometriosis due to the lack of biomarkers (Fassbender et al. 2015, Macer and Taylor 2012,

Kennedy et al. 2005). Treatments for endometriosis center mainly on pain management or relief.

Several accepted therapies include the use of GnRH analogues or levonorgestrel intrauterine

devices. Laparoscopic surgery is often used to ablate and excise the tissue or transect nerves.

These interventions significantly reduce pain and lead to increased pregnancy and live birth rates

(Brown and Farquhar 2015, Kennedy et al. 2005).

1.10.2.2 Role of Extracellular Matrix in Pathogenesis of Endometriosis

Retrograde menstruation, in which endometrial tissue is disseminated and implanted into

the peritoneal cavity via the fallopian tubes, is the most accepted theory for the pathogenesis of

endometriosis (Ahn et al. 2015, Sampson 1927). Between 75-90% of women undergo retrograde

menstruation; however, women that develop endometriosis are likely to have other genetic and

biochemical factors leading to development of the disease (Ahn et al. 2015). It is not clearly

understood how the endometrial tissue attaches to the peritoneum, whether it is directly to the

peritoneal mesothelium or to the underlying ECM (Adachi et al. 2011, Lucidi et al. 2005, Witz et

al. 2003, Dunselman et al. 2001, Nisolle et al. 2000, Sillem et al. 1999). Differences in

expression of adhesion molecules that are part of the ECM and ECM signaling, such as integrins

and cadherins, appear to play a role in the development of endometriosis (reviewed in Young et

al. (2013)). Numerous integrin subtypes were present in human peritoneal mesothelium and in

endometriotic lesions in a nude mouse model (Hull et al. 2008, Witz et al. 2000). Endometriotic

samples also had impaired expression and localization of several integrin receptors, suggesting a

43  

role in enhanced invasion (Giannelli et al. 2007, Klemmt et al. 2007, Sillem et al. 1999). E-

cadherin was expressed in healthy peritoneal and normal endometrial tissue (Jimenez-Heffernan

et al. 2004, Groothuis et al. 1999). An increased number of endometriotic lesions appeared to be

E-cadherin negative though (Gaetje et al. 1997). Increased expression of P-cadherin in

endometriotic tissue compared to normal endometrial tissue suggested that it also played a role in

increased cell adhesion (Chen et al. 2002).

Endometriotic tissue invasion was also facilitated by MMPs (Sharpe-Timms and Cox

2002). Expression of numerous MMPs, including MMP3, 9, 10, 12, and 13, was increased in

human samples and animal models. The most common MMP to increase was MMP9.

Interestingly, eutopic endometrial tissue in women with endometriosis also had increased

expression and activity of MMP9 compared to normal eutopic endometrial tissue. Expression of

TIMP-1 appeared to vary though (Pelch et al. 2010, Pino et al. 2009, Collette et al. 2006, Collette

et al. 2004). Increased MMP9 expression and decreased TIMP-1 expression was also found in

peritoneal fluid (Szamatowicz et al. 2002). These findings suggest that MMP9 may aid in

degrading the ECM in the peritoneum, allowing ectopic endometrial tissue to invade and implant

into the peritoneal cavity.

1.10.3 Pyometra

1.10.3.1 Description, Etiology, and Treatment of Pyometra

Pyometra is an inflammatory condition characterized by the accumulation of intraluminal

purulent material in the uterus and can be divided into open- and closed-cervix distinctions.

Open-cervix pyometra means that the purulent fluid can drain, while closed-cervix pyometra

means that the fluid remains trapped in the uterine cavity. Closed-cervix pyometra is an

emergency requiring rapid intervention to prevent systemic infection and death (Smith 2006). In

44  

humans, pyometra is extremely rare yet life threatening, with a 0.04% incidence rate in the

general population. The development of pyometra has been reported in children as young as four

months of age. In the elderly, this rate jumps to >13%, showing an age-dependent increase in

development (Barry 2015, Geggie and Walton 2006, Yildizhan et al. 2006). In humans, pyometra

often occurs due to impairment in natural drainage subsequent to damage from a malignant

disease such as cervical cancer and radiotherapy treatment (for example Vyas et al. (2009),

Yildizhan et al. (2006)). In veterinary medicine, pyometra is most common in intact canines,

with nearly 25% presenting with the disease before 10 years of age (Smith 2006, Egenvall et al.

2001), but pyometra is also found in small and captive large felids, livestock, mares, and even

rhesus monkeys, rhinoceros, and orca whales (Knudsen et al. 2015, Patisaul 2012, McCain et al.

2009, McCarthy et al. 1989, Ridgway 1979).

Treatment for pyometra varies with the severity of the condition. Complete

ovariohysterectomy is preferred in older animals, those with systemic inflammation, or animals

with closed-cervix pyometra (Smith 2006). Otherwise, the standard protocol is treatment with

antibiotics, fluids, and either prostaglandins or antiprogestins in an attempt to reverse progression

and save breeding capabilities (Fieni et al. 2014, Smith 2006).

A combination of hormonal, genetic, and inflammatory or infectious factors play a role in

the etiology of pyometra. However, not all rodent strains treated with estrogenic compounds will

develop pyometra. Treatment with estrogenic or progestogenic compounds, including

phytoestrogens, E2, DES, and contraceptives, in various species led to the development of the

disease (Asa et al. 2014, Patisaul 2012, Brossia et al. 2009, Gould et al. 2005, Pandey et al. 2005,

McCarthy et al. 1989, McLachlan et al. 1980, Burrows 1936). Brown Norway, but not Sprague

Dawley, ACI, or Fischer 344, rats developed pyometra after E2 treatment or DES exposure

45  

(Brossia et al. 2009, Gould et al. 2005, Pandey et al. 2005). Using quantitative trait loci (QTL)

mapping, key genetic variations between the Brown Norway, ACI, and Fischer 344 strains were

found in Eutr1 for E2-induced pyometra and Eutr2 for DES-induced inflammation and

pyometra, supporting that genetic factors play a role in the etiology of the disease (Gould et al.

2005, Pandey et al. 2005). Strain susceptibility can also be used to identify other endpoints that

are estrogen-responsive as well.

1.10.3.2 Role of Bacteria and the Immune System in the Development of Pyometra

During metestrus in the reproductive cycle, the uterus is primed for embryonic

implantation and the maternal immune system is decreased in order to allow the embryo,

considered a foreign body, to implant. However, bacterial infection in the uterus from the normal

flora in the vaginal or digestive tract is also highly likely to occur during this stage due to the

decreased immune response. In roughly 70% of cases in canines in which bacteria are present in

the uterine cavity, Escherichia coli was cultured with a high percentage displaying virulence

factors (Agostinho et al. 2014, Hagman 2012, Coggan et al. 2008, Hagman and Kuhn 2002).

Canines inoculated with E. coli at different estrous cycle stages only developed a severe

pyometric phenotype when inoculated in early metestrus (Tsumagari et al. 2005). In the bursa

surrounding the ovaries in canines, E. coli was present but there was also bacterial isolates of

strains of Staphylococcus and Streptococcus, among others (Rubio et al. 2014), suggesting that

infection from several different bacterial types can lead to the development of pyometra.

Antibiotics are often used to combat the bacterial infection; however, 50% of canines with

pyometra had multidrug resistance (Agostinho et al. 2014), suggesting that treatment with

antibiotics may not be useful and could lead to further antibiotic resistance.

46  

The presence of bacteria leads to an immune or inflammatory response in the uterus.

Toll-like receptors (TLRs) are important in regulating the innate and adaptive immune responses

in the host tissue in response to bacterial lipopolysaccharide (LPS) binding (Takeda et al. 2003,

Janeway and Medzhitov 2002, Hoshino et al. 1999). In canines with pyometra, mRNA and

protein expression of TLR2 and TLR4 were significantly increased in all cell types examined

(Chotimanukul and Sirivaidyapong 2012, 2011, Silva et al. 2010), potentially due to the

increased bacterial content in the uterus. This increased expression could also represent a change

associated with the development of pyometra, rather than be playing a causative role. Other

immune cells types (ie. granulocytes, macrophages, and leucocytes) were decreased in cows with

pyometra, while lymphocytes were increased (Brodzki et al. 2014). Expression of mRNA and

protein for pro-inflammatory interleukins (ILs), including IL-1α, IL-1β, and IL-6, were increased

in pyometric samples from affected animals (Brodzki et al. 2015, Voorwald et al. 2015,

Bukowska et al. 2014). Anti-inflammatory ILs, such as IL-4 and IL-10, were also increased in

animals with pyometra (Maciel et al. 2014), suggesting an attempt by the animal to combat the

effects of the present bacteria and developing disease. Alterations in cytokine production resulted

in differences in prostaglandin synthesis. Prostaglandin F2α synthase (PGFS) and prostaglandin

E2 synthase (PGES) mRNA levels were significantly increased in animals with pyometra,

leading to increases in prostaglandin E2 and F2α levels (Silva et al. 2010). It has been proposed

that several of the altered proteins could be potential predictive biological markers to detect

animals with pyometra or animals which could develop pyometra without invasive procedures

(Dabrowski et al. 2015, Voorwald et al. 2015, Dabrowski et al. 2013, Karlsson et al. 2012).

47  

1.10.3.3 Involvement of Sex Hormones and Collagen Accumulation in Pyometra

Alterations in sex hormone receptor levels and collagen accumulation were observed in

canines and felids with pyometra. Serum P levels were unaffected, but serum E2 levels were

greatly increased in both halves of diestrus and early anestrus (Ververidis et al. 2004). While

progesterone receptor (PR) expression was decreased in surface epithelium only, ERα expression

was decreased in all areas of tissue, including surface epithelium, stromal cells, normal glands,

cystic glands, and smooth muscle (Kunkitti et al. 2011, Misirlioglu et al. 2006, De Bosschere et

al. 2002). Within the cervix of canines with pyometra, the closed-cervix phenotype had a

significantly decreased percentage of collagen compared to open-cervix, while both had a

decreased percentage of collagen compared to normal cervical tissue (Linharattanaruksa et al.

2014). These decreases in collagen may be due to the substantial upregulation in MMPs and

associated activity (Voorwald et al. 2015, Hagman 2012, Chu et al. 2002).

1.10.4 Equine Endometrosis

1.10.4.1 Description, Etiology, and Treatment of Equine Endometrosis

Equine endometrosis is distinct from human endometriosis and is defined as an age-

related and irreversible degenerative uterine disease characterized by gland nest structures

surrounded by increased endometrial stromal and periglandular fibrosis (EPF). The disease can

be classified into Kenney categories I, IIA, IIB, and III based on disease severity, progression,

and amount of fibrosis and inflammation (Buczkowska et al. 2014, Hanada et al. 2014, Snider et

al. 2011, Evans et al. 1998, Ferreira-Dias et al. 1994, Ricketts and Alonso 1991, Kenney 1978).

When classifying equine endometrosis, several potential pathological changes are examined,

including the presence of inflammatory infiltrates, infection, fibrosis and gland nesting,

hypertrophy associated with fibrosis, lymphatic lacunae, cystic glands or nonglandular cysts,

48  

atrophy/hypoplasia, and glandular hyperplasia. Kenney category I is characterized by no

pathologic changes or changes that are slight and scattered. Category I mares do not have an

impaired ability to support a fetus to term. Kenney category II is characterized by more extensive

or severe inflammatory changes, increase in fibrosis of individual gland branches or gland nests,

presence of atrophy, and presence of lacunae but are not large enough to palpate. Mares in

category II have impairment in the ability to carry a foal to term. The severity of the fibrotic

changes distinguishes categories IIA and IIB, with category IIA having 10-35% of glands

affected and category IIB having 35-60% of glands affected. Kenney category III is

characterized by widespread fibrosis, severe and widespread inflammation, and severe lymphatic

lacunae. These pathologic changes greatly reduce fertility, with only about 10% of mares having

the ability to carry a foal to term (Buczkowska et al. 2014, Kenney 1978). The severity, or higher

category, of EPF is inversely correlated to successful conception and gestation and is associated

with increased rates of embryonic or fetal foal loss and increased susceptibility to infection

(Buczkowska et al. 2014, Kenney 1978). An endometrosis-like phenotype has also been reported

in canine biopsy samples (Christensen et al. 2012).

While it is unclear if genetic or hormonal factors are involved in the etiology of equine

endometrosis, the involvement of increased immune cell infiltration has been reported. Increased

neutrophil infiltration possibly occurred in response to bacterial challenge or non-infectious

stimuli, such as semen (Rebordao et al. 2014). This increase in inflammation led to the formation

of neutrophil extracellular traps (NETs) containing pro-inflammatory proteins that damaged

nearby tissue and contributed to the development of fibrosis (Rebordao et al. 2014).

No effective treatment for equine endometrosis has been developed. Intrauterine

infusions with 30% dimethyl sulfoxide (DMSO) in 0.9% saline decreased inflammatory cells and

49  

periglandular fibrosis in barren mares but fertility rates remained low (Ley et al. 1989).

However, follow-up studies to determine if the severity of endometrosis returned to that

observed pre-treatment were lacking. Recently, mesenchymal stem cell (MSC) therapy has been

proposed as a potential treatment for endometrosis due to their ability to give rise to several cell

types and their trophic mediation, including in inflammatory and fibrotic processes. The

differentiated cells then secrete growth factors, cytokines, and MMPs to degrade the fibrotic

components and inhibit the inflammatory response (van Poll et al. 2008, Caplan and Dennis

2006, Aggarwal and Pittenger 2005, Caplan 1991). A simple delivery method, similar to that

used for artificial insemination, using equine adipose-derived MSCs (eAD-MSCs) has been

developed and eAD-MSCs delivered through this technique are incorporated into and distributed

throughout uterine tissue (Mambelli et al. 2013). Mares with endometrosis treated with eAD-

MSCs exhibited decreases in vimentin, laminin, cytokeratin 18, and α-SMA staining as early as

Day 7 after treatment, as well as improved morphological characteristics. An increase in Ki-67

staining was present in control and experimental groups at Days 7, 21, and 60 after treatment, but

the experimental group exhibited larger increases at all three time points. However, a large

sample size was lacking, with only four mares treated with eAD-MSCs, and one mare with

advanced degeneration exhibiting no changes in protein expression with treatment (Mambelli et

al. 2014). While decreases in phenotypic markers were present, it was unknown if symptoms

were decreased or if fertility was improved. This method is limited by the local nature of

treatment, preventing progression or reoccurrence of endometrosis, and will require a suitable

systemic treatment method to be developed as well in order to fully abolish endometrosis from

the endometrium. Also, the treatment may not be useful on mares with highly advanced disease.

50  

Several questions still exist about etiology, progression, and treatment of equine endometrosis

without an experimental model that can be used for rapid and high powered experimentation.

1.10.4.2 Progression of Equine Endometrosis

Early stages of endometrosis were accompanied by periglandular-associated stromal cells

that actively produce collagen fibers (Buczkowska et al. 2014). A progression of the disease was

proposed based on alterations in proteins involved in proliferation and ECM and sex hormone

protein levels (Hoffmann et al. 2009b). After the early stages of fibrosis were underway, either

an active or inactive non-destructive phenotype developed (Table 2); however, the factors

regulating if the tissue becomes an active or inactive phenotype were unknown (Hoffmann et al.

2009b) . An active phenotype was characterized by the presence of metabolically active fibrotic

stromal cells and myofibroblasts, while an inactive phenotype was characterized by the

arrangement and contractibility of myofibroblasts (Hoffmann et al. 2009b). However, the

presence of myofibroblasts in general was indicative of advanced endometrosis (Walter et al.

2001).

To distinguish between the active and inactive phenotypes, immunohistochemical

examinations was performed, with the inactive phenotype having a mild decrease in Ki-67

antigen in fibrotic stromal cells compared to normal tissue while the active phenotype was not

changed. In glandular epithelia, the inactive phenotype had decreases in ERα and PR compared

to normal tissue while the active phenotype had increases in both receptors (Hoffmann et al.

2009b). Eventually, endometrosis progressed to an active or inactive destructive form. An active

destructive form resulted from an increase in myofibroblasts and ECM proteins and the inactive

destructive phenotype arose from mechanical stress (Hoffmann et al. 2009b). Based on an

immunohistochemical assessment, inactive destructive endometrosis again had a decrease in Ki-

51  

67 antigen in fibrotic stromal cells compared to normal tissue while the active destructive type

did not. Active destructive endometrosis had 50-80% of cells staining for fibronectin or

proteoglycane, while the inactive destructive phenotype had 20-40% of cells staining for

fibronectin and 10-20% of cells staining for proteoglycane. Finally, the amount of Ki-67 antigen

in the glandular epithelia was decreased in the active destructive phenotype compared to normal

tissue, while the inactive destructive phenotype had no change in the amount of Ki-67 antigen. In

the glandular epithelia, the active destructive phenotype displayed decreases in ER while the

active non-destructive phenotype displayed increases in ER (Hoffmann et al. 2009b). However,

this proposed progression was limited due to the potential bias when estimating the percentage of

positive cells and the staining intensity. Some of the proposed changes appeared too small to

quantify, so the distinction between active and inactive phenotypes might not be meaningful.

Table 2. Characteristics of Equine Endometrosis Phenotypes

Non-destructive Destructive Active Inactive Active Inactive

Fibrotic Stromal Ki-67 − ↓ − ↓

Glandular Epithelium Ki-67 − − ↓ −

PR ↑ ↓ ↓ ↓ ERα ↑ ↓ ↓ ↓

ECM Fibronectin 10-20% 20-40% 50-80% 20-40%

Proteoglycane 10-20% 10-20% 50-80% 10-20%

1.10.4.3 Characterization of Extracellular Matrix Alterations in Equine Endometrosis

Because fibrosis is highly involved in the development of endometrosis, varying

expression patterns of ECM proteins between the defined categories or phenotypes were

characterized. Periglandular-associated fibroblasts produced increased levels of collagen IV,

laminin, and fibronectin. These fibroblasts appeared to have also differentiated into

52  

myofibroblasts, based on their expression of α-SMA, tropomyosin, transforming growth factor-

β1 (TGF-β1), and 11β-hydroxysteroid dehydrogenase (11β-HSD2) (Ganjam and Evans 2006,

Walter et al. 2001). While synthesis of ECM proteins was increased in endometrosis, MMP2 and

tissue transglutaminase 2 (TG2), an enzyme involved in ECM protein cross linking, expression

was also increased (Walter et al. 2005). Because TG2 cross links are resistant to proteolytic

cleavage, the rate and level of collagen deposition was increased (Walter et al. 2005, Skill et al.

2004, Greenberg et al. 1991). While TG2 was degraded by an MMP2/MMP14 complex, it also

interacted with an intermediate form of MMP2 and decreased the rate of MMP2 maturation

(Walter et al. 2005, Belkin et al. 2004). This imbalance in collagen and MMP2 homeostasis may

lead to the increased fibrosis observed in affected, degenerating regions of the uterus.

1.10.4.4 Fetal Foal Loss

Fetal foal loss in mares with equine endometrosis may be due to alterations in the

endometrial environment, specifically expression patterns of interleukins and prostaglandins that

are involved in estrous cycle regulation and pregnancy in several farm animals (Franczak et al.

2010, Weems et al. 2006, Tanikawa et al. 2005). Myofibroblasts, which are present in advanced

endometrosis, increased cytokine production (Zhang et al. 1996). When comparing Kenney

categories II and III to category I in each stage of the estrous cycle, significant alterations in

prostaglandin-endoperoxide synthase (PTGS-2), PGFS, and PGES mRNA levels were found,

which correlated with alterations in prostaglandin F2α and prostaglandin E2 concentrations

(Szostek et al. 2012). Comparing the same categories within the stages of the estrous cycle,

significant differences in IL mRNA transcription were observed as well, which translated into

differences in protein expression. Specific IL proteins examined include IL-1α, IL-1β and the IL-

1 receptor subunits (IL-1RI and IL-1RII), and IL-6 and the IL-6 receptor (IL-6Rα) (Szostek et al.

53  

2013). Endometrial samples from each category were treated with either IL-1α, IL-1β, or IL-6 to

determine the influence of ILs on prostaglandin secretion and synthase mRNA transcription.

Each category exhibited distinctive alterations in patterns of secretion and transcription levels

(Szostek et al. 2013). The increased differentiation of fibroblasts into myofibroblasts in

categories II and III of endometrosis may be leading to altered expression in ILs and subsequent

altered prostaglandin synthase expression and prostaglandin concentrations. The abnormal

endometrial environment then resulted in decreased fertility and fetal loss.

Several proteins are normally secreted from endometrial glands during the estrous cycle

in fertile humans and animals. Aberrant expression of these proteins may indicate a reduction in

fertility in general and may be used to determine the differences between fertile and barren

mares. Uteroglobin (UG), uterocalin (UC), uteroferrin (UF), and calbindinD9k (CAL) are

important secreted proteins involved in a successful pregnancy. UG has anti-inflammatory

properties and prevents the maternal immune system from attacking trophoblastic cells during

implantation and placental growth (Mukherjee et al. 1994, Miele et al. 1990). UC is a carrier

protein that transports hydrophobic proteins and molecules to the conceptus (Suire et al. 2001,

Crossett et al. 1998). UF is primarily an iron transporter to the conceptus, but also is a

hematopoietic growth factor (Bazer et al. 1991, Ducsay et al. 1984, McDowell et al. 1982). CAL

is a Ca2+-binding transporter in the uterus (Wooding et al. 1996, Inpanbutr et al. 1994). In both

non-destructive and destructive endometrosis, expressions of UG, UC, and CAL were largely

decreased compared to unaffected tissue, while UF expression was increased (Lehmann et al.

2011, Hoffmann et al. 2009a). Decreases in secreted proteins essential for viable growth of a

fetus in endometrosis also played a significant factor in decreased fertility and fetal loss.

54  

1.11 Statement of Purpose

1.11.1 Objective and Rationale

To repeat, the overall objective of this dissertation research was to characterize the

endocrine disrupting effects of BPA or EE exposure in the uterus of two mouse strains. Two

exposure paradigms were used: 1) exposure in adulthood, during which time mice mated and

produced a litter, and 2) whole life exposure without breeding. Previous studies examined

incidence of pathologies after a much longer exposure time period than is proposed in this

dissertation research. Uterine immune responsiveness and fibrosis are components of several

human and animal uterine pathologies, and the effect of BPA or EE exposure on these endpoints

has not been studied.

1.11.2 Central Hypothesis and Specific Aims

The central hypothesis for this dissertation was that exposure to estrogenic EDCs, such as

BPA or EE, will influence E2-regulated uterine physiology, leading to an increased incidence of

uterine pathology. A second hypothesis was that a greater sensitivity in the C57Bl/6N strain to

the estrogenic actions of EDCs also increases the incidence of uterine pathology compared to the

CD1 strain.

The specific aims of this dissertation research included 1) investigating the endocrine

disrupting actions of EDCs in both exposure models on the development of uterine pathologies,

2) examining the effects of EDCs in both exposure models on the immune responsiveness in the

uterus, 3) investigating the ECM changes associated with EDC exposure in both models, and 4)

comparing the pathological, immune, and ECM alterations between the C57Bl/6N and CD1

strains.

55  

CHAPTER 2

METHODS

56  

2.1 BPA and EE Exposure

2.1.1 Animal Housing

All animal procedures were performed in accordance with protocols approved by the

University of Cincinnati Institutional Animal Care and Use Committee and followed

recommendations of the Panel on Euthanasia of the American Veterinary Medical Association.

C57Bl/6NHsd and Hsd:ICR (CD1) mice (Harland; Indianapolis, IN) aged six to seven weeks

were evenly distributed between the nine exposure groups, with five of a single sex occupying a

cage for an acclimation period of two weeks. During this period, general health and vaginal

cytology were monitored daily and body weight and food consumption were monitored weekly.

Animals were kept on a polycarbonate-free housing system with single-use polyethylene cages

and water bottles certified BPA-free (Innovive; San Diego, CA). Sanichip bedding (Irradiated

Aspen Sani-chip; PJ Murphy Forest Products Corp.; Montville, NJ) was used to prevent

contamination from mycoestrogens in corncob bedding (Thigpen et al. 2013). Sterile drinking

water containing <1% of oxidizable organics was produced from a water purification system

(Millipore Rios 16 with ELIX UV/Progard 2; Billerica, MA). A 14-hour on, 10-hour off light

cycle was maintained throughout the study. Diet and water were provided ad libitum.

2.1.2 Exposure

C57Bl/6N and CD1 mice were randomly assigned and divided equally between a control

group (0 ppm), five BPA groups (0.03, 0.3, 3, 30, or 300 ppm), and three EE groups (0.0001,

0.001, 0.01) as a positive control for estrogenic activity. In a pilot study, higher EE doses were

used (0.1 and 1.3 ppm) but were discontinued in later phases. Mice remained on diet from arrival

to necropsy. Offspring were exposed to BPA or EE from conception to necropsy (Figure 1).

Food consumption was recorded weekly. The amount of food eaten per cage was divided by the

57  

number of animals per cage and the number of days between food weight collections to estimate

the amount of food consumed per animal. Actual doses of BPA or EE were calculated based on

estimated food consumption. Animals were fed a defined casein-based phytoestrogen-free diet

(Product # D1010501, Research Diets, Inc.; New Brunswick, NJ) unsupplemented (control or 0

ppm) or supplemented with either BPA (2,2-bis(4-hydroxyphenyl)propane; CAS No. 80-05-7;

Lot 11909; USEPA/NIEHS standard) or EE (1,3,5(10)-estratrien-17α-ethinyl-3,17β-diol; CAS

No. 57-63-6; Batch No. H923; Steraloids Inc.; Newport, RI) that was incorporated

homogenously in the diet at each desired concentration. Final BPA concentrations (0.03, 0.3, 3,

30, and 300 ppm) resulted in doses of 0.004, 0.04, 0.4, 4, and 40 mg/kg/day. Concentrations of

EE (0.0001, 0.001, and 0.01 ppm) resulted in doses of 0.00002, 0.0002, and 0.001 mg/kg/day

(Kendziorski and Belcher 2015, Kendig et al. 2012, Kendziorski et al. 2012). Details of diet

composition are found in Table 1 and amounts of BPA or EE and dyes incorporated into the diets

are found in Table 2.

Figure 1. Exposure paradigm for sires, dams, and offspring in control and BPA or EE exposure groups.

58  

Table 1. Composition of Defined Control and Test Diets

Macronutrients Source gm% kcal% Ingredients g kcal

Protein Casein 18.1 20 Casein 200 800

Carbohydrate D-glucose 60.3 65 L-Cysteine 3 12

Corn Starch Maltodextrin

Corn Starch 346 1384

Fat Soy Oil 6.2 15 Maltodextrin 10 45 180

Fiber Cellulose 11.7 Dextrose 250 1000

Inulin 2.3 Cellulose, BW 200 125 0

Total ME (kcal/g)* 3.63 100 Inulin 25 25

Soybean Oil 70 630

Mineral Mix S10026 10 0

Dicalcium Phosphate 13 0

Calcium Carbonate 5.5 0

Potassium Citrate, 1

H2O 16.5 0

Vitamin Mix V10001 10 40

Choline Bitartrate 2 0

Total 1121.05 4071

*Metabolizable energy estimate based on the Atwater factors: protein: 4 kcal/g; fat: 9 kcal/g; carbohydrate: 4 kcal/g

Table 2. Amounts of EE, BPA, and Dyes Incorporated into Diets Control BPA EE 0 0.03 0.3 3 30 300 0.0001 0.001 0.01 0.1 1.3

17α-Ethinyl Estradiol (µg)

0 0 0 0 0 0 0.12 1.2 12 120 1470

Bisphenol A (µg) 0 33.7 337 3370 33700 337000 0 0 0 0 0

Red Dye #40,

FD&C (g) 0 0.04 0 0.04 0.025 0 0 0.05 0.025 0 0.05

Blue Dye #1, FD&C (g)

0 0.01 0.04 0 0 0.025 0.05 0 0.025 0.05 0

Yellow Dye #5, FD&C (g)

0.05 0 0.01 0.01 0.025 0.025 0 0 0 0 0

59  

2.1.3 Breeding

After the acclimation period, males and females within an exposure group were paired

single-housed for breeding and observed daily within the first three hours of the light-on cycle

for a copulation plug. Once a plug was observed, vaginal cytological assessment ceased and the

breeding pair was separated. If a copulation plug was not observed after two weeks, female body

weight gain was used as an indicator of pregnancy and the breeding pair was separated. Females

were observed twice daily for general health, signs of pregnancy, and parturition (designated as

PND0). Offspring were separated at PND22 and group housed ≤5 per sex per cage. The dams

were housed 5 per cage after separation from their offspring.

2.1.4 Vaginal Cytology

In order to examine the stage of reproductive cycle and to ensure female mice were

analyzed and sacrifice occurred during the estrus phase of the estrous cycle, vaginal cytology

was performed. Vaginal lavage was performed daily during the first three hours of the light-on

cycle using a 0.09% sterile saline solution and a fire polished glass pipette. Samples were

collected and a microscopic cytological assessment was completed. Each stage of the cycle had

different morphological features to the vaginal cells in the collected sample. Estrus is indicated

by the presence of cornified, non-nucleated epithelial cells. The presence of infiltrating immune

cells such as leukocytes and few epithelial cells with nuclear degeneration is indicative of

diestrus. Proestrus is indicated by the presence of round, nucleated epithelial cells (McLean et al.

2012).

Estrous stage identification determined by vaginal cytology was confirmed by

morphological assessment of the hematoxylin & eosin (H&E)-stained uteri for C57Bl/6N and

CD1 dams and offspring. Staining of uteri is described in Section 2.2.2. Mice in estrus and

60  

proestrus have a distended lumen, with the lumen being thicker in estrus, indicative of tissue

breakdown. Mice in diestrus have a thin lumen in the process of regeneration (Hedrich and

Bullock 2004). Mice that were classified as being in estrus or diestrus were used for all studies.

2.2 Assessment of Pathology

2.2.1 Necropsy and Tissue Preparation

Dams were sacrificed between 19 and 23 weeks of age, and offspring were sacrificed

~PND90. All animals were sacrificed in estrus, based on cytological assessment of vaginal cells.

Animals were evenly and randomly divided into a group in which all tissue would be frozen and

a group in which all tissue would be perfused. Randomization was based on the identification

number of the animal, with the first half being frozen and the second half being perfused. All

tissues were collected as part of a larger study (Kendig et al. 2012). Half of the study mice were

euthanized by rapid cervical dislocation to collect frozen tissue or CO2 asphyxiation to collect

perfused tissue. Tissues were quickly excised and excess fat and connective tissue were

removed. Tissues were weighed and placed in a 2 mL free standing microcentrifuge tube with

screw cap (Fisher Scientific; Waltham, MA) for freezing in a 100% ethanol/dry ice mixture.

Frozen tissue samples were stored at -80oC until used. For the other half of the study animals, the

chest cavity was opened and a cardiac puncture of the left ventricle was used in order to flush the

animal with heparinized saline and then perfuse with 10% neutral-buffered formalin (NBF;

Polysciences, Inc.; Warrington, PA). Tissues were collected, trimmed of excess fat, weighed, and

placed in properly sized cassettes and washed in multiple changes of 10% NBF and then washed

in six washes of 70% ethanol before tissue processing.

Tissues were processed using a Tissue Tek VIP 3000 Tissue Processor (Sakura/Miles

Scientific; Torrance, CA) using the following program: one wash in 70% ethanol for 20 minutes,

61  

two washes in 80% ethanol for 20 minutes each, two washes in 95% ethanol for 20 minutes each,

one wash in 100% ethanol for 20 minutes, two washes in 100% ethanol for 30 minutes each, two

washes in xylene for 30 minutes each, and three washes in heated paraffin under vacuum seal for

30 minutes each (58oC; Type 3 paraffin, Richard Allen Scientific; Kalamazoo, MI). After

processing, tissues were embedded in paraffin (Histoplast IM; Richard Allen Scientific;

Kalamazoo, MI) using a Shandon Histocentre 3 Embedding Station (Thermo Shandon;

Kalamazoo, MI). To embed uteri, a section of one uterine horn was cut and embedded

transversely to assess myometrial wall thickness. The rest of each uterus was placed

longitudinally in the same paraffin block.

2.2.2 Hematoxylin and Eosin Staining

Paraffin-embedded uteri from C57Bl/6N and CD1 dams and offspring were sectioned at a

5 µm thickness on an HM 325 Rotary Microtome (Thermo Shandon; Kalamazoo, MI) and placed

on SuperFrost Plus slides (Fisher Scientific; Waltham, MA). Slides were incubated at 60oC for

>20 minutes in a hybridization incubator (Fisher Scientific; Waltham, MA). Once cooled to

room temperature (RT), an H&E stain was performed in order to visualize the nuclei of cells

(hematoxylin) and the basic compartments or cytoplasm (eosin). Sections were deparaffinized in

three washes of xylene for 5 minutes each, rehydrated through a graded series of ethanol (two

washes in 100% ethanol for 3 minutes each, two washes in 95% ethanol for 1 minute each, and

one wash in 70% ethanol for 1 minute), and rinsed in running tap water for 5 minutes. Slides

were rinsed in dH2O, stained in Hematoxylin 2 (Cat# 7231; Richard Allen Scientific;

Kalamazoo, MI) for 30 seconds, and rinsed in running tap water. Slides were placed in Clarifier

2 (Cat# 7442; Richard Allen Scientific; Kalamazoo, MI) for 30 seconds and rinsed in running tap

water for 30 seconds. Slides were placed in Bluing Reagent (Cat# 7341; Richard Allen

62  

Scientific; Kalamazoo, MI) for 1 minute and rinsed in running tap water for 1 minute. Slides

were placed in 95% ethanol for 1 minute and counterstained with Eosin-Y (Cat# 7111; Richard

Allen Scientific; Kalamazoo, MI) for 2 minutes. Excess eosin was blotted with paper towel and

slides were dehydrated through three washes of 100% ethanol for 15 dips each. Slides were

placed in three washes of xylene for 15 dips each and coverslipped using Permount (Fisher

Scientific; Waltham, MA), which were allowed to dry overnight.

2.2.3 Myometrial Wall Thickness

Transverse sections of the H&E-stained uterine tube of C57Bl/6N and CD1 dams from

the preliminary study were measured in order to quantify the thickness of the inner and outer

myometrial layers. Using a Nikon Eclipse 55i microscope using a DS-Fi1 CCD camera

controlled with DigitalSight software, five random areas at 20x magnification of the outer edge

of the transverse section were imaged by an observer blinded to strain and exposure group.

Within each image, five measurements from the beginning of the inner myometrium to the outer

edge of the outer myometrium were collected using DigitalSight software. Using a stage

micrometer, the ratio of 60 µm/100 pixels was measured. Wall thickness measurements were

converted from pixels to micrometers (µm) using the following formula:

X = 60 µm (pixel measurement) 100 pixels

2.2.4 Histologic Assessment of Uterine Pathology

The H&E-stained uteri of C57Bl/6N and CD1 dams and offspring (PND21, PND49, and

~PND90) were assessed for distended glands, gland nests, cysts, and pyometra. Distended glands

are characterized by a thinning of the epithelial cells that make up the gland. Gland nests are

groups of glands that appear clumped together in an oval or circle and are surrounded by fibrotic

63  

stromal cells instead of being dispersed throughout the endometrium. Cysts appear as circular

holes in the endometrium surrounded by a thin layer of epithelial cells. The presence of the

pathology and, when applicable, the number of incidences were noted for each uterus. The

observer was blinded to strain, exposure, and generation.

2.3 Assessment of Collagen Synthesis and Degradation and Immune Response

2.3.1 Picrosirius Red Staining and Collagen Accumulation

Paraffin-embedded uterine sections for C57Bl/6N and CD1 dams and offspring were cut

at a 5 µm thickness on an HM 325 Rotary Microtome and put on SuperFrost Plus slides. Slides

were incubated at 60oC for >20 minutes in a hybridization incubator. Slides were deparaffinized

in three washes of xylene for 5 minutes each, rehydrated in graded series of ethanol (two washes

in 100% ethanol for 2 minutes each, two washes in 95% ethanol for 2 minutes each, and one

wash in 70% ethanol for 2 minutes), and rinsed in dH2O. Slides were stained with Weigert’s

hematoxylin (American MasterTech; Lodi, CA) for 8 minutes and rinsed with running tap water

for 5 minutes. Sections were placed in a 0.2% phosphomolybdic acid hydrate in water for 2

minutes, rinsed with dH2O for 30 seconds, and placed in a Picrosirius Red F3BA (1.4% 2,4,6-

trinitrophenol, 0.4% Direct Red 80) solution in water for 60 minutes. Slides were transferred to a

0.1 N hydrochloric acid solution for 2 minutes and placed in 70% ethanol for 45 seconds. Slides

were dehydrated (one wash of 95% ethanol for 15 dips and three washes of xylene for 15 dips

each) and washed in xylene three times for 15 dips each. Sections were coverslipped using

Permount.

2.3.2 Immunohistochemistry

Uteri sections were cut at a 5 µm thickness on an HM 325 Rotary Microtome and put on

SuperFrost Plus slides. Slides were incubated at 60oC for >20 minutes in a hybridization

64  

incubator. Slides were deparaffinized in three washes of xylene for 5 minutes each, rehydrated in

graded series of ethanol (two washes of 100% ethanol for 3 minutes each, two washes of 95%

ethanol for 1 minute each, and one wash of 70% ethanol for 1 minute), and rinsed in running tap

water for 5 minutes. Slides were immersed in 0.01 M citrate buffer (anhydrous citric acid, pH

6.0; Fisher Scientific; Waltham, MA) containing 0.05% Tween-20 at 100oC for 10 minutes and

allowed to cool for 30 minutes. Slides were washed twice in their respective wash solutions, with

20 dips per wash. The wash solution for MMP2, F4/80, α-SMA, and ERα was phosphate-

buffered saline (PBS, pH 7.6; 3.8 mM NaH2PO4 (Fisher Scientific; Waltham, MA), 16 mM

Na2HPO4 (Fisher Scientific; Waltham, MA), and 0.15 M NaCl (Fisher Scientific; Waltham,

MA)) and the wash solution for MMP14 was Tris-buffered saline with Tween-20 (TBST, pH

7.6; 20 mM Tris base (Fisher Scientific; Waltham, MA), 137 mM NaCl, 0.1% Tween-20 (Fisher

Scientific; Waltham, MA)). Endogenous peroxidase activity was blocked by immersing slides in

PBS containing 3% H2O2 (Fisher Scientific; Waltham, MA) for 10 minutes and rinsing in

running tap water for 5 minutes. Non-specific binding sites were blocked by 3% normal goat

serum (Vector Laboratories; Burlingame, CA) in PBS for one hour at RT in a humidified

incubator tray (Ciba Corning Diagnostics; Medfield, MA). Sections were then incubated

overnight at 4oC in a humidified incubator tray with rabbit polyclonal anti-MMP2 (3.33 µg/mL

(1:300); NB200-193, Novus Biologicals; Littleton, CO), rabbit monoclonal anti-MMP14 (0.5

µg/mL (1:250); ab51074, Abcam; Cambridge, MA), rat monoclonal anti-F4/80 (C1:A3-1; 4

µg/mL (1:250); ab6640, Abcam; Cambridge, MA), rabbit polyclonal anti-α-SMA (1 µg/mL

(1:200); ab5694, Abcam; Cambridge, MA), or rabbit polyclonal anti-ERα (MC-20; 1 µg/mL

(1:200); sc-542, Santa Cruz Biotechnology; Dallas, TX) Additional information regarding these

antibodies can be found in Table 3. Coverslips were removed in PBS and slides were washed

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three times in wash solution for 5 minutes each. Sections incubated in biotinylated goat anti-rat

(mouse adsorbed) (10 µg/mL (1:50); Vector Laboratories; Burlingame, CA) or biotinylated goat

anti-rabbit (30 µg/mL (1:50); Vector Laboratories; Burlingame, CA) diluted in 3% normal goat

serum in PBS for 1 hour at RT in humidified incubator trays. Additional information regarding

these antibodies can be found in Table 3. Slides were washed in wash solution three times for 5

minutes per wash. The avidin-biotin complex (ABC; Vector Laboratories; Burlingame, CA) was

prepared 30 minutes prior to use by adding 2 drops of avidin DH (reagent A) and two drops of

biotinylated horseradish peroxidase H (reagent B) to 5 mL of PBS. Sections were incubated in

ABC solution for 1 hour at RT in humidified incubator trays. Slides were washed in wash

solution three times for 5 minutes per wash. Sections were visualized using 0.05% 3’3-

diaminobenzidine (DAB; Vector Laboratories; Burlingame, CA). DAB was prepared

immediately before use by adding 2 drops of buffer (pH 7.5), 4 drops of DAB, and 2 drops of

H2O2 to 5 mL of dH2O. Visualization was stopped by placing the slides in PBS. After all of the

sections developed, slides were washed in tap water three times for 5 minutes per wash. Slides

were rinsed in dH2O for 15 dips, counterstained in Hematoxylin 2 for 30 seconds, and rinsed in

running tap water. Slides were placed in Clarifier 2 for 30 seconds and rinsed in running tap

water for 30 seconds. Slides were placed in Bluing Reagent for 1 minute and rinsed in running

tap water for 1 minute. Slides were dehydrated through one wash of 95% ethanol for 15 dips and

three washes of 100% ethanol for 15 dips each. Slides were placed in three washes of xylene for

15 dips each and coverslipped using Permount. Mouse spleen was used as positive control tissue

for F4/80 staining, mouse lung was used as positive control tissue for MMP2 and MMP14

staining, mouse heart was used as positive control tissue for α-SMA staining, and rat mammary

tumor was used as positive control tissue for ERα staining.

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The number of F4/80-positive cells in the uterus was quantified. Five random images at

40x magnification of myometrium and five random images at 40x magnification of endometrium

in which mostly stroma was visible were collected. For each image of endometrium and

myometrium, the total number of positively stained cells and the total number of stromal or

smooth muscle cells were counted using Photoshop CC and ImagePro Plus v4.5 (Media

Cybernetics; Rockville, MD). The number of positively stained cells was then divided by the

total number of cells and multiplied by 100 to generate the percent of F4/80-positive cells in each

mouse.

Table 3. Primary and Secondary Antibodies

Antigen Source Catalog # Conc IHC

(µg/mL)

Dilution IHC

Conc WB

(µg/mL)

Dilution WB

Species/Type Ref for

Specificity

MMP2 Novus NB200-193 3.33 1:300 0.25 1:4000 Rabbit

polyclonal

MMP14 Abcam ab51074 0.5 1:250 0.066 1:2000 Rabbit

monoclonal

Farahani et al.

(2012)

β-actin Cell

Signaling 8457 0.02 1:2000

Rabbit monoclonal

F4/80 Abcam ab6640 4 1:250 Rat

monoclonal

α-SMA Abcam ab5694 1 1:200 Rabbit

polyclonal

Farahani et al.

(2012)

ERα Santa Cruz

sc-542 1 1:200 Rabbit

polyclonal

Garcia-Falgueras

et al. (2011)

goat

anti-rabbit Vector BA-1000 30 1:50 Goat IgG

goat anti-rat

Vector BA-9401 10 1:50 Goat IgG

2.3.3 Quantitative RT-PCR

Frozen tissue of uterine horns ≤30 mg from CD1 mice exposed to 0 ppm and 30 ppm

BPA were homogenized for RNA isolation using an RNeasy Fibrous Tissue Mini Kit (Qiagen;

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Valencia, CA). Tissue was homogenized in 300 µL of Buffer RLT (RNA lysis buffer containing

guanidine thiocyanate). To the homogenized tissue, 590 µL of RNase-free water (Teknova;

Hollister, CA) and 10 µL of proteinase K were added, mixed, and incubated at 55oC for 10

minutes. Samples were then centrifuged at 10,000 x g for 3 minutes. The supernatant was

transferred to a new tube, 450 µL of 100% ethanol was added, and the tube was mixed. Of the

total amount of supernatant, 700 µL was transferred to an RNeasy Mini column and the column

was centrifuged for 15 seconds at 8,000 x g. The flow through was discarded and the step was

repeated until all of the supernatant was centrifuged. Next, 350 µL of Buffer RW1 (RNA wash

buffer 1 containing ethanol) was added to the RNeasy column, centrifuged for 15 seconds at

8,000 x g, and the flow through was discarded. In the column, 10 µL of DNase stock solution

was mixed with 70 µL of Buffer RDD (DNase wash buffer) and incubated for 15 minutes at RT.

After the incubation, 350 µL of Buffer RW1 was added to the column and centrifuged for 15

seconds at 8,000 x g. The flow through was discarded and 500 µL of Buffer RPE (wash buffer)

was added to the column. The column was centrifuged for 15 seconds at 8,000 x g and the flow

through was discarded. A second 500 µL of Buffer RPE was added to the column and

centrifuged for 2 minutes at 8,000 x g. The column was placed in a new 1.5 mL microcentrifuge

tube and 30-50 µL of RNase-free water was added to the column. The column was centrifuged

for 1 minute at 8,000 x g and repeated with the RNA eluate. RNA concentrations and potential

contamination were determined using a Nanodrop 2000 spectrophotometer (Thermo Scientific;

Waltham, MA). RNase-free water was used as a standard. Concentrations of RNA are given as

ng/µL. The 260/280 ratio can indicate protein contamination and should be around 2.0

(ThermoScientific 2011). The 260/230 ratio indicates salt, carbohydrate, phenol, EDTA, or

guanidine HCl contamination and should also be around 2.0-2.2 (ThermoScientific 2011). RNA

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concentrations and ratios for samples can be found in Table 4. Most 260/280 ratios were around

2.0, but the low 260/230 ratios suggest a high level of residual salt or guanidine. RNA was stored

at -80oC until further use.

Table 4. RNA Concentrations

Sample ID Concentration (ng/µL) 260/280 (nm) 260/230 (nm) 0 ppm 1102 74.1 2.07 0.82 1104 90.9 2.05 0.57 1106 40.5 2.05 0.43 1108 31.0 1.90 0.52 1110 53.1 2.02 0.76 1111 153.0 2.04 1.26 1112 137.0 2.02 1.42 1114 177.3 1.94 1.17

30 ppm BPA

1502 66.1 2.00 0.72 1504 65.7 2.02 0.55 1508 129.3 1.94 0.96 1509 150.1 1.67 0.82 1510 73.0 2.00 0.54 1511 41.2 2.02 0.31 1512 29.8 1.91 0.80 1513 200.9 2.07 1.14 1514 401.7 2.07 1.37

Uterine RNA was converted to cDNA using a High Capacity cDNA Reverse

Transcription Kit (Life Technologies; Grand Island, NY). A 2x RT master mix was prepared

based on the number of reactions. Per reaction, 2 µL of 10x RT buffer, 0.8 µL of 25x dNTP mix

(100 mM), 2 µL of 10x RT random primers, 1 µL of MultiscribeTM reverse transcriptase, and 4.2

µL of RNase-free water were mixed and kept on ice. In a domed PCR tube, 10 µL of the 2x RT

master mix and 10 µL of RNA sample were mixed by pipetting up and down twice. Tubes were

briefly centrifuged in a Centrifuge 5810R (Eppendorf; Hamburg, Germany) at 4,000 rpm at 4oC

and loaded into a PTC-150 Minicycler (MJ Research; Waltham, MA). To perform reverse

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transcription, the following conditions were used: 10 minutes at 25oC, 120 minutes at 37oC, 5

minutes at 85oC, and ≤24 hours at 4oC. The resulting cDNA was stored at -20oC until further use.

Predesigned TaqMan RT-PCR assays (Applied Biosystems; Grand Island, NY) were

used for the genes found in Table 5. PCR amplification was performed in a total volume of 20

µL containing 20x TaqMan expression assay primers (1 µL/reaction), TaqMan Universal Master

Mix (10 µL/reaction; Applied Biosystems; Grand Island, NY), RNase-free water (8 µL/reaction),

and 10 ng of cDNA (1 µL/reaction). Amplification was performed in triplicates on a Step One

Plus Real-Time PCR System (Applied Biosystems; Grand Island, NY) under the following fast

conditions: 20 seconds at 95oC for one holding stage, then 1 second at 95oC and 20 seconds at

60oC for the cycling stage (60 cycles). Relative expression was quantified using the ΔΔCt

method, using 18s (Mm03928990_g1) as the endogenous control for normalization.

Table 5. TaqMan Probe Sets Used for qRT-PCR Analysis

Gene Name Gene

Symbol ABI Assay

Entrez Gene

ID Unigene # NCBI Location Chromosome

collagen, type I, alpha 1

Col1a1 Mm00801632_gH 12842 Mm.277735 Chr.11: 94936270 - 94951867

collagen, type III, alpha 1

Col3a1 Mm00802296_g1 12825 Mm.249555 Chr.1: 45311538 - 45349706

matrix metallopeptidase 2

Mmp2 Mm00439498_m1 17390 Mm.29564 Chr.8: 92827292-92853420

tissue inhibitor of metalloproteinase 1

Timp1 Mm00441818_m1 21857 Mm.8245 Chr.X: 20870166 - 20874737

tissue inhibitor of metalloproteinase 2

Timp2 Mm00441825_m1 21858 Mm.206505 Chr.11: 118301061 - 118355411

tissue inhibitor of metalloproteinase 3

Timp3 Mm00441826_m1 21859 Mm.4871 Chr.10: 86300412 - 86349505

18s ribosomal RNA 18s Mm03928990_g1 19791

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2.3.4 SDS-PAGE and Immunoblotting

Frozen uterine tissue was homogenized in lysis buffer (20 mM Tris-HCl (Fisher

Scientific; Waltham, MA), 150 mM NaCl, 1 mM EDTA (Fisher Scientific; Waltham, MA), 1

mM EGTA (Fisher Scientific; Waltham, MA), 1% Triton X-100 (Bio-Rad; Hercules, CA),

cOmplete Protease Inhibitor Cocktail (EDTA-free; Roche; Basel, Switzerland), pH 7.5) and

centrifuged at 4,000 rpm for 10 minutes at 4oC on a Centrifuge 5810R (Eppendorf; Hamburg,

Germany). The supernatant was collected and stored at -20oC until further use.

A colorimetric DC protein assay (Bio-Rad; Hercules, CA) was used in order to quantify

the protein concentration of the uterine homogenates. A standard protein curve using bovine

serum albumin (Frac V) (BSA; Fisher Scientific; Waltham, MA) was generated by diluting 50

mg of BSA in 1 mL of lysis buffer. Serial dilutions were made for 1.5 mg/mL, 1.0 mg/mL, 0.5

mg/mL, and 0.25 mg/mL from the 50 mg/mL solution. Lysis buffer was used for 0.0 mg/mL. A

working reagent was prepared by combining 20 µL of reagent S with each mL of reagent A

needed to make reagent A’. Reagent A’ is needed to solubilize detergent in the lysis buffer. If

necessary, uterine lysates were diluted in lysis buffer. In a 1.5 mL microcentrifuge tube, 20 µL of

standard or lysate was combined with 100 µL of reagent A’ and 800 µL of reagent B. After a 15

minute incubation at RT, 230 µL of the reaction was transferred to a flat bottom 96-well plate

(Denville Scientific; South Plainfield, NJ) in triplicate. Absorbance at 750 nm was determined

using a SpectraFluor Plus spectrofluorometer with Magellan 3 software (Tecan; Männedorf,

Switzerland). Absorbance readings were converted to protein concentrations in Microsoft Excel

(Microsoft; Redmond, WA) by first averaging the absorbance readings for the 0 mg/mL standard

and subtracting this value from all readings. A standard curve was generated by averaging the

71  

values for each protein standard and linear regression analysis was used to calculate protein

concentrations for the uterine lysates.

Once protein concentrations were calculated, total protein (30 µg) was mixed 3:1 with

sample buffer (0.25 M Tris-HCl, 40% glycerol (Fisher Scientific; Waltham, MA), 8% SDS (Bio-

Rad; Hercules, CA), 0.01% bromophenol blue (Fisher Scientific; Waltham, MA), pH 6.8)

containing 0.04% β-mercaptoethanol (Fisher Scientific; Waltham, MA) and incubated at 96oC

for 5 minutes. Prestained SDS-PAGE protein standards of known molecular weights (Broad

Range; Bio-Rad; Hercules, CA) were used to determine relative molecular weights based on

mobility. Standards and protein lysates were loaded onto and resolved by electrophoresis on a

10% polyacrylamide gel (10% acrylamide (Fisher Scientific; Waltham, MA), 0.27%

bisacrylamide (Bio-Rad; Hercules, CA), 375 mM Tris base, 0.1% SDS, 0.025% ammonium

persulfate (APS; Bio-Rad; Hercules, CA), and 0.067% tetramethylethylenediamine (TEMED;

Bio-Rad; Hercules, CA), pH 8.8) with a 4% stacking gel (3.9% acrylamide, 0.1% bisacrylamide,

125 mM Tris base, 0.1% SDS, 0.0375% APS, and 0.1% TEMED, pH 6.8).

After electrophoresis, gels and Odyssey® nitrocellulose membranes (Li-Cor Biosciences;

Lincoln, NE) were placed in transfer buffer (48 mM Tris base, 39 mM glycine (Fisher Scientific;

Waltham, MA), 0.0375% SDS, 20% methanol (Fisher Scientific; Waltham, MA)) for 30 minutes

at 4oC. A wet transfer was then set up in the following order: black side of cassette, sponge, filter

paper, gel, membrane, filter paper, sponge, and white side of cassette. In between each layer, a

glass rod was used to expel of any bubbles. The black side of the cassette was placed facing the

black side of the Mini Trans-Blot Cell (Bio-Rad; Hercules, CA) for wet transfer. Transfer

occurred for 16 hours at 4oC at a constant amperage of 70 mA. Membranes were stained with

0.1% Ponceau S (Fisher Scientific; Waltham, MA) in 1% acetic acid (Acros Organics; Waltham,

72  

MA) to ensure protein transfer. Membranes were blocked with 5% non-fat dry milk (Blotting

Grade Blocker, Bio-Rad; Hercules, CA) in TBST for 1 hour at RT and then incubated overnight

at 4oC with either rabbit polyclonal anti-MMP2 (0.25 µg/mL (1:4000); NB200-193, Novus

Biologicals; Littleton, CO) or rabbit monoclonal anti-MMP14 (0.066 µg/mL (1:2000); ab51074,

Abcam; Cambridge, MA). A rabbit monoclonal anti-β-actin antibody (0.02 µg/mL (1:2000);

8457, Cell Signaling; Danvers, MA) was used as a loading control. Additional information

regarding these antibodies can be found in Table 3. Membranes were washed three times in

TBST for 5 minutes per wash and incubated with IRDye® 800CW goat anti-rabbit IgG (1:5000;

Li-Cor Biosciences; Lincoln, NE) for 1 hour at RT. Membranes were washed in TBST three

times for 5 minutes per wash and placed in TBS. Membranes were imaged and analyzed using an

Odyssey® CLx Infrared Imaging System (Li-Cor Biosciences; Lincoln, NE) and Image Studio

Software (Li-Cor Biosciences; Lincoln, NE).

2.3.5 Gelatin Zymography

Zymography is a molecular technique that uses electrophoretic methodologies to quantify

the activity of certain enzymes by adding a substrate to the gel. In this case, activity of MMP2

was quantified by using gelatin as a substrate. Methods for gelatin zymography were adapted

from previously described protocols (Toth et al. 2012, Belcher et al. 2005).

Frozen uterine tissue was homogenized in lysis buffer (25 mM Tris-HCl, 0.1 M NaCl,

1% Nonidet P-40 (Shell Chemicals; The Hague, Netherlands), cOmpleteTM Protease Inhibitor

Cocktail (EDTA-free), pH 7.5; Roche; Basel, Switzerland) and centrifuged at 4,000 rpm for 10

minutes at 4oC on a Centrifuge 5810R. The supernatant was collected and stored at -20oC until

further use.

73  

Protein concentrations of the uterine lysates were calculated using the colorimetric DC

protein assay, similar to the procedure used to quantify protein concentration for Western

blotting. The BSA standards were generated and lysates were diluted using lysis buffer specific

for zymography.

Once protein concentrations were calculated, 30 µg of total protein was mixed 3:1 with

sample buffer (0.25 M Tris-HCl, 40% glycerol, 8% SDS, 0.01% bromophenol blue, pH 6.8)

without the presence of a reducing agent. The lysate:sample buffer mixture was incubated at RT

for 15 minutes. Samples were loaded onto a 10% polyacrylamide gelatin-based zymography gel

(10% acrylamide, 0.27% bisacrylamide, 375 mM Tris base , 0.1% SDS, 0.025% APS, 0.067%

TEMED, 0.1% gelatin (Bio-Rad; Hercules, CA), pH 8.8) with a 4% stacking gel (3.9%

acrylamide, 0.1% bisacrylamide, 125 mM Tris base, 0.1% SDS, 0.0375% APS, and 0.1%

TEMED, pH 6.8). Following electrophoresis, zymograms were incubated in 2.5% Triton X-100

for 30 minutes at RT with gentle agitation in order to remove SDS from the samples and renature

the proteins. After renaturation, zymograms were rinsed in MilliQ H2O and incubated in

developing buffer (50 mM Tris-HCl, 0.2 M NaCl, 5 mM CaCl2 (Fisher Scientific; Waltham,

MA), pH 7.8) for 30 minutes at RT with gentle agitation. The developing buffer was decanted

and zymograms incubated in fresh developing buffer for 16 hours at 37oC in an isotemp

incubator (Fisher Scientific; Waltham, MA). Zymograms were stained with Coomassie Brilliant

Blue R-250 (Bio-Rad; Hercules, CA) dissolved in a 5% methanol/10% acetic acid solution for 3

hours at RT with gentle agitation. Zymograms were destained in a 5% methanol/10% acetic acid

solution for 15 minutes, 30 minutes, and 1 hour at RT with gentle agitation. Clear bands denoted

areas of proteolytic activity. Conditioned media from the human brain tumor cell line DAOY

(ATCC HTB-186) was used as a positive control for MMP activity (Belcher et al. 2009). In

74  

control experiments, the addition of EDTA (1 mM final concentration) to lysates or developing

buffer resulted in loss of proteolytic activity at the expected molecular weights. Gels were

imaged using an Alpha Innotech Multiimage II Gel Documentation System (Alpha Innotech

Corp.; San Leandro, CA) and quantified using Image Studio Lite Software.

2.4 Statistical Methods

When applicable, the observer was blinded to strain, exposure group, and generation. The

litter was the statistical unit used; one animal was selected from each litter as representative of

that litter. Pathology was assessed with statistical differences analyzed using Fisher’s exact test

or χ2 test for trend when assessing across PND. Myometrial wall thickness, gland nest density,

and percentage of F4/80-positive cell counts were analyzed using a two-way ANOVA and

Bonferroni post hoc test in dams and a one-way ANOVA and Dunnett’s multiple comparison test

in offspring. A Student’s t-test was used for analysis of mRNA and protein expression and

protein activity. C57Bl/6N offspring endpoints were not statistically analyzed due to a low

sample size for all groups. Outliers were removed based on Grubb’s test. Significance between

differences in values was defined as p<0.05. All data was analyzed using GraphPad Prism®

v5/v6 software (GraphPad; La Jolla, California).

75  

CHAPTER 3

Strain Specific Induction of Pyometra and Differences in Immune Responsiveness in Mice

Exposed to 17α-Ethinyl Estradiol or the Endocrine Disrupting Chemical Bisphenol A

Kendziorski JA, EL Kendig, RB Gear, and SM Belcher. 2012. Strain specific induction of pyometra and differences in immune responsiveness in mice exposed to 17α-ethinyl estradiol or the endocrine disrupting chemical bisphenol A. Reprod Toxicol. 34(1): 22-30. doi: 10.1016/j.reprotox.2012.03.001. PMID: 22429997

76  

Strain Specific Induction of Pyometra and Differences in Immune Responsiveness in Mice Exposed to 17α-Ethinyl Estradiol or the Endocrine Disrupting Chemical Bisphenol A

Jessica A. Kendziorski, Eric L. Kendig, Robin L. Gear and Scott M. Belcher*

Department of Pharmacology and Cell Biophysics University of Cincinnati College of Medicine

Cincinnati, OH, 45267-0575

*Corresponding Author: Scott M. Belcher Department of Pharmacology and Cell Biophysics University of Cincinnati College of Medicine 231 Albert Sabin Way; PO Box 670575 Cincinnati, OH, 45267-0575 Telephone: 513-558-1721 Fax: 513-558-4329 Email: scott.belcher@uc.edu

77  

3.1 Abstract

Pyometra is an inflammatory disease of the uterus that can be caused by chronic exposure to

estrogens. It is unknown whether weakly estrogenic endocrine disruptors can cause pyometra.

We investigated whether dietary exposures to the estrogenic endocrine disruptor bisphenol A

(BPA) induced pyometra. Pyometra did not occur in CD1 mice exposed to different dietary

doses of BPA ranging from 4.1 to >4000 µg/kg/day or 17α-ethinyl estradiol (EE; 1.2 to >150

µg/kg/day). In the C57Bl/6 strain, pyometra occurred in the 15 µg/kg/day EE and 33 µg/kg/day

BPA treatment groups. At the effective concentration of BPA, histological analysis revealed

pathological alterations of uterine morphology associated with a >5.3-fold increase in

macrophage numbers in non-pyometra uteri of C57Bl/6 mice exposed to BPA. These results

suggest that BPA enhances immune responsiveness of the uterus and that heightened

responsiveness in C57Bl/6 females is related to increased susceptibility to pyometra.

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3.2 Introduction

Inflammatory conditions of the uterus occur in specific strains of rats and mice receiving

extended exposures to estrogens (Burrows 1935). Pyometra is literally defined as “puss in the

uterus.” This potentially life-threatening infectious disease is characterized by inflammation and

intraluminal accumulation of purulent material in the uterus. Pyometra is clinically most well-

known to veterinary medical science, where it is a common occurrence in unspayed female dogs;

nearly 25% of bitches will be affected by pyometra before the age of 10 years (Smith 2006,

Egenvall et al. 2001). In addition to dogs, pyometra, while less common, is also a significant

disease in domestic cats and captive large felids, some livestock species, and laboratory animals

such as rabbits and some strains of mice and rats. In humans, pyometra is a relatively uncommon

condition with rare but significant mortality resulting from spontaneous uterine perforation or

rupture leading to septicemia. In the general population, it is estimated to account for about

0.04% of gynecological admissions; however, the incidence becomes increased to >13% in the

elderly (Yildizhan et al. 2006).

Hormonal factors, particularly the actions of estrogen on the uterus, play a central role in

the etiology of pyometra. Along with hormonal status and age, genetic factors related to cellular

infiltration of leukocytes into the uterus play a major role in regulating sensitivity to estrogen-

induced uterine inflammation and pyometra (Hagman et al. 2011, Gould et al. 2005, Roper et al.

1999). In sensitive strains of laboratory rats and mice, it is well established that chronic exposure

to estradiol or the highly efficacious non-steroidal estrogen, diethylstilbestrol (DES), can induce

pyometra (Gould et al. 2005, Pandey et al. 2005, McLachlan et al. 1980, Stone et al. 1979,

Gardner and Allen 1937, Zondek 1936, Burrows 1935). These findings suggest the possibility

that the highly prevalent environmental estrogenic endocrine disrupting chemical, bisphenol A

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(BPA), may have a similar effect. Bisphenol A is a synthetic monomer used in the production of

epoxy resin-based sealants and polycarbonate plastics. Measurable levels of BPA have been

found in >90% of humans tested at levels that affect fecundity, embryonic development and

carcinogenesis in experimental models (Ye et al. 2011, Richter et al. 2007, Calafat et al. 2005) .

The ability of endocrine disrupting chemicals such as BPA, that are considered weak estrogens

based on potency in a uterotrophic bioassay or binding affinity at the nuclear estrogen receptors,

to cause pyometra is essentially unknown. However, BPA can cause morphological changes in

the CD1 mouse uterus at doses considered to be safe in humans, suggesting mechanisms other

than those assessed by the rodent uterotrophic bioassay are involved with some actions of BPA

in the uterus (Markey et al. 2001). As part of a larger effort to determine the physiological

consequences of dietary exposure to the endocrine disrupting chemical BPA, we investigated

whether BPA might act in an estrogen-like fashion to induce pyometra in mice exposed to BPA

or 17α-ethinyl estradiol (EE) as a positive control for estrogenic activity in their diet.

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3.3 Materials and Methods

3.3.1 Animal husbandry and dietary exposures

All animal procedures were performed in accordance with protocols approved by the

University of Cincinnati Institutional Animal Care and Use Committee and followed

recommendations of the Panel on Euthanasia of the American Veterinary Medical Association.

Animals were maintained on a 14 h light, 10 h dark cycle and provided food and water ad

libitum in single-use polyethylene caging certified BPA-free (Innovive, San Diego California).

Sani-chip bedding (Irradiated Aspen Sani-chip; P.J. Murphy Forest Products Corp. Montville,

NJ) was used to avoid corncob-derived mycoestrogen exposure. Sterile drinking water was

generated by a dedicated water purification system (Millipore Rios 16 with ELIX UV/Progard 2)

that reduces organic contaminants to less than 1% of source levels. Drinking water was

dispensed from BPA-free polyethylene water bottles (Innovive, San Diego California).

A modified open standard diet (supplemental data table 1; Product #10010504 Research

Diets) was used to eliminate possible dietary phytoestrogen contamination. Test compounds

were directly incorporated into the extruded pellet diets that were unsupplemented (control) or

supplemented with 17α-ethinyl estradiol (EE; 1,3,5(10)-estratrien-17α-ethinyl-3,17β-diol, CAS

No. 57-63-6, Batch No. H923; Steraloids Inc., Newport, RI) or bisphenol A (2,2-bis(4-

hydroxyphenyl)propane; BPA; CAS 80-05-7; Lot 11909; USEPA/NIEHS standard) at desired

final concentrations (supplemental data table 1). For BPA, all three doses used were designed to

fall below the NOAEL (50 mg/kg/da). The high dose (30 ppm) was between the NOAEL and the

reference dose (50 μg/kg/da), the middle dose (0.3 ppm) approximated the reference dose, and

the low dose (0.03 ppm) fell below the reference dose (1982). Diets containing EE were

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estimated to be at or below an effective dose previously reported to decrease fecundity in mice

(1.3 ppm, 0.1 ppm and 0.01 ppm) (Thayer et al. 2001) .

Seventy (35 of each sex) C57Bl/6NHsd and seventy Hsd:ICR (CD1 Swiss) mice aged six

to seven weeks were received from Harlan Laboratories, Inc. (Indianapolis, IN). Males and

females from each strain were group housed five per cage with the same sex and evenly divided

among control or one of the six dietary treatments and maintained on those treatments from

arrival until necropsy. Mice were allowed to acclimate to housing and treatment for two weeks.

Body weight and food consumption were monitored weekly, while general health and vaginal

cytology were monitored daily. Following the acclimation period, males and females in the same

group were randomly paired in individual housing and observed daily within the first 3 hr of the

light cycle for copulation plug. Upon observation of a copulation plug, assessment of vaginal

cytology ceased, the male was removed, and females were individually housed and observed

twice daily. Dams were supplied with nestlets, monitored for general health, and observed for

parturition. Body weights of both males and females were monitored weekly until necropsy.

Dams and their offspring were housed together until postnatal day 21, at which time the

offspring were separated and dams were group housed with F0 females from the same treatment

group.

3.3.2 Monitoring of estrous

Progression of estrous cycles was monitored in F0 females by morphological analysis of

daily vaginal lavage prior to copulation. Briefly, approximately 200 µl of sterile NaCl solution

(0.09%) was used to flush the vulva using a fire polished pipette. Stage of the estrous cycle was

determined by microscopic assessment of the vaginal cell morphology in each lavage sample.

Samples with epithelial cells in various stages of nuclear degeneration and cytoplasmic shrinkage

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and polymorphonuclear leukocytes indicated diestrus; samples with nucleated endothelial cells

were indicative of proestrus; and samples with cornified, the presence of non-nucleated cellular

debris was characteristic of estrus.

3.3.3 Necropsy and tissue preparation

Female F0 mice were euthanized by CO2 asphyxiation and complete necropsy was

performed between the ages of 19 and 23 weeks. Digital images of animals, uteri, and ovaries

were captured as part of gross examination during necropsy (Nikon D3000 with 50 mM Nikkor

macro lens). Tissues were excised and cleaned of excess connective tissue and fat, blotted of

excess fluids, and weighed. Isolated tissues were fixed with several changes of 5%

paraformaldehyde or 10% neutral buffered formalin. Tissues were blocked by sectioning with a

sterile razor blade and washed several times in 70% ethanol prior to automated tissue processing

and embedding in paraffin (Histocenter 3; Thermo-Shandon Kalamazoo, MI). Microtome

sections were cut at 5 µm thickness from blocks at 4˚C and placed on positively charged slides

for hematoxylin and eosin (H&E) staining and immunohistochemistry. Standard H&E staining

was performed in order to examine tissue structure and morphology.

3.3.4 Immunohistochemistry

Paraffin sections were dewaxed and rehydrated through graded series of ethanol

concentrations into phosphate buffered saline (PBS, pH 7.4) and endogenous peroxidase blocked

by treatment with 3% H2O2 in PBS. After washing, heat-mediated epitope retrieval was

performed at 100oC in 10 mM citric acid with 0.05% Tween-20 (pH 6.0). Sections were

incubated at room temperature for >30 min with 3% normal goat serum in PBS, washed, and

then incubated overnight at 4˚C with 4 µg/mL of a macrophage specific F4/80 rat monoclonal

antibody (Cl:A3-1; Abcam Cambridge, Mass). Immunoreactivity was visualized with 0.05%

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3’3-diaminobenzidine by the avidin-biotin peroxidase complex method (Vector Laboratories).

Stained sections were lightly counterstained in hematoxylin for 30 seconds (Richard Allan;

Kalamazoo, MI) and mounted in Permount (Fisher Scientific, Fair Lawn NJ). Internal controls

for specificity of immunostaining included replacement of primary or secondary antibodies with

non-specific serum. Sections prepared from spleen served as positive controls. Microscopic

examination of immunostained material was carried out using a Nikon Eclipse 55i microscope

using a DS-Fi1 CCD camera controlled with Digital Sight software (Nikon Instruments, Inc.,

Melville, NY). For quantitative assessments of immunopositive cell numbers, five random 40x

fields of the myometrium and five random 40x fields of the endometrium were collected from a

single section for each animal. The number of immunopositive cells per field was manually

counted by an investigator blinded to treatment and strain. To account for possible region

specific differences in infiltration, the mean number of cells in each of five 40x fields for the

myometrium and the endometrium was determined for one section per individual. The sum of

those values was taken to be representative of the number of F4/80 positive cells for that

individual. Final photomicrograph graphics were generated and labeled using Adobe Photoshop

CS3 (Adobe Systems, Inc., San Jose, CA).

3.3.5 Statistical analysis

Analysis of weight and cell count data was performed using one-way ANOVA followed

by Tukey’s Multiple Comparison post-hoc test with Cramer’s correction or Dunnett’s multiple

comparison test using GraphPad Prism v5 software (GraphPad Prism®, GraphPad Software,

Inc., La Jolla, CA). Significance between differences in values was defined by a p-value of

<0.05.

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3.4 Results

3.4.1 Diet consumption

The amount of chow consumed by each female animal was determined immediately after

mating. From this consumption data, estimations of caloric intake and oral dose of EE and BPA

were calculated (Table 1).

For C57Bl/6 animals, the presence of 0.1 and 1.3 ppm EE in diet resulted in a significant

decrease in food consumption and the intake of significantly fewer calories per day as compared

to control diet. While EE dose dependently decreased consumption and caloric intake in CD1

animals, the difference in values for any one treatment group were not different from control. No

differences in consumption or caloric intake were detected in additional studies comparing

control diet and diet supplemented with 0.001 and 0.0001 ppm EE (data not shown). The

presence of BPA was not associated with any change in consumption or caloric intake for either

strain.

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Table 1. Estimated Food Consumption, Caloric Intake, and Dose

C57Bl/6 Control EE 0.01

ppm EE 0.1 ppm

EE 1.3 ppm

BPA 0.03 ppm

BPA 0.3 ppm

BPA 30 ppm

Food Consumption (g/mouse/day) 2.7 ± 0.4 2.2 ± 0.5 1.8 ± 0.5a 1.6 ± 0.2a 2.5 ± 0.4 2.5 ± 0.4 2.6 ± 0.4

Caloric Intake (kcal/mouse/day) 9.7 ± 1.5 7.9 ± 1.7 5.4 ± 1.9a 5.8 ± 1.0a 9.2 ± 1.5 9.2 ± 1.6 9.6 ± 1.4

Dose (μg/kg/day) 0.0 1.4 ± 0.4 14.7 ± 6.4 164.3 ± 49.5 4.0 ± 1.1 32.8 ± 10.3 4,168.0 ± 1,358.0

Sample Size 4 5 4 4 5 5 4

CD-1 Control EE 0.01

ppm EE 0.1 ppm

EE 1.3 ppm

BPA 0.03 ppm

BPA 0.3 ppm

BPA 30 ppm

Food Consumption (g/mouse/day) 4.2 ± 1.9 4.8 ± 1.8 4.0 ± 2.3 3.2 ± 2.0 4.6 ± 1.6 3.8 ± 0.8 4.7 ± 1.6

Caloric Intake (kcal/mouse/day) 11.2 ± 1.5 12.7 ± 2.0 11.1 ± 4.3 8.5 ± 2.6 12.3 ± 2.1 11.4 ± 3.7 13.2 ± 2.4

Dose (μg/kg/day) 0.0 1.2 ± 0.3 11.6 ± 3.6 158.8 ± 25.3 4.1 ± 1.1 41.7 ± 10.7 4182.0 ± 1218.0

Sample Size 5 5 4 4 5 5 5

Values represent mean of all food consumption measurements ± SD a: significantly different from control by one-way ANOVA (p < 0.01).

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3.4.2 Effects on body weight

Body weights were monitored from the onset of study until study completion when

animals were between 19 and 23 weeks of age. Not surprisingly, EE treatment dose dependently

impacted the body weight (Table 2). In the 1.3 ppm EE dose group, body weight was

significantly decreased at the end of the first study week. Lower concentrations of EE similarly

decreased body weight following an inverse correlation with dose (Table 2). There was also a

dose responsiveness observed in the onset of detectable differences in body weight of control and

EE dose groups. After the second week on treatment, both 0.1 and 1.3 ppm EE groups weighed

significantly less than controls. Those trends continued through the end of study. No difference

in either the amount or the rate of weight gained was observed in females of any of the BPA

groups (Table 1; Table 2). It thus appears that there is a dose dependent aversion to consuming

diet containing EE and that the sensitivity to EE diet was strain dependent.

After mating, body weight measurements for mated gravida and nulligravida females

were analyzed separately to avoid confounders on body weight related to metabolic

consequences of pregnancy, parturition, and lactation status. While impacts on body weight

related to treatment and dose remain evident during those later time points, the resulting

reduction in sample size prevented meaningful statistical analysis post copulation.

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Table 2. Female Body Weight C57Bl/6

Wks Control EE 0.01 ppm EE 0.1 ppm EE 1.3 ppm BPA 0.03 ppm BPA 0.3 ppm BPA 30 ppm

0 14.4 ± 0.6 14.1 ± 0.4 14.4 ± 0.4 14.7 ± 0.6 14.3 ± 0.8 13.7 ± 0.9 14.2 ± 0.9

1 15.5 ± 0.6 15.4 ± 0.9 14.0 ± 0.4 13.1 ± 0.4a 16.1 ± 0.8 15.8 ± 0.6 16.4 ± 1.8

2 16.5 ± 0.6 15.7 ± 0.9 14.7 ± 0.9a 13.2 ± 1.0a 16.6 ± 0.7 16.1 ± 0.7 16.4 ± 0.6

3 17.5 ± 1.0 16.3 ± 1.3 14.6 ± 0.7a 13.3 ± 0.7a 17.8 ± 0.9 17.3 ± 0.4 17.2 ± 0.3

Status gravida nulli

gravida gravida

nulli gravida

gravida nulli

gravida gravida

nulli gravida

gravida nulli

gravida gravida

nulli gravida

gravida nulli

gravida

4 17.9 ± 0.9 17.1 ± 0.4 15.2 16.3 ± 1.1 NA 14.7 ± 0.8 NA 12.8 ± 0.9 18.2 ± 0.6 17.0 ± 0.5 17.7 ± 0.4 17.2 ± 0.6 17.6 ± 0.3 16.4

5 19.8 ± 1.1 19.0 ± 0.8 15.9 17.3 ± 1.5 NA 15.4 ± 0.4 NA 13.1 ± 1.5 19.9 ± 0.8 18.1 ± 1.2 19.7 ± 0.7 18.8 ± 0.0 19.7 ± 0.6 17.6

6 26.4 ± 1.7 20.4 ± 0.2 17.1 19.2 ± 0.6 NA 15.4 ± 1.1 NA 13.1 ± 0.6 25.5 ± 0.2 19.3 ± 0.1 26.6 ± 2.6 20.1 ± 0.1 26.7 ± 3.0 18.9

7 24.3 ± 0.8 21.2 ± 1.1 18.2 20.2 ± 2.0 NA 16.2 ± 0.6 NA 13.5 ± 1.3 32.3 ± 8.5 21.4 ± 0.2 27.8 ± 6.1 21.7 ± 1.2 26.3 ± 4.7 19.6

CD-1

Wks Control EE 0.01 ppm EE 0.1 ppm EE 1.3 ppm BPA 0.03 ppm BPA 0.3 ppm BPA 30 ppm

0 20.1 ± 1.4 20.8 ± 0.6 19.9 ± 0.7 20.2 ± 1.7 20.3 ± 0.9 19.6 ± 1.3 19.3 ± 1.4

1 22.5 ± 1.2 24.0 ± 1.2 20.8 ± 1.7 18.6 ± 1.4a 23.3 ± 1.0 23.4 ± 2.7 23.7 ± 1.6

2 24.6 ± 1.5 26.6 ± 1.8 21.8 ± 1.7 18.7 ± 1.3a 25.4 ± 2.0 26.4 ± 4.2 27.0 ± 3.4

3 23.6 ± 1.0 25.0 ± 1.6 21.3 ± 1.3 18.3 ± 1.1a 24.0 ± 1.8 25.5 ± 3.2 25.6 ± 2.3

Status gravida nulli

gravida gravida

nulli gravida

gravida nulli

gravida gravida

nulli gravida

gravida nulli

gravida gravida

nulli gravida

gravida nulli

gravida

4 26.0 ± 1.3 NA 27.9 ± 1.5 26.4 ± 2.0 NA 22.5 ± 1.2 NA 19.0 ± 1.4 25.2 ± 2.3 27.3 ± 2.6 27.4 ± 3.0 25.6 ± 2.8 26.8 ± 0.5 29.7

5 33.6 ± 2.4 NA 35.7 ± 6.2 31.2 ± 0.5 NA 23.0 ± 1.3 NA 19.3 ± 1.3 32.3 ± 5.1 29.0 ± 3.2 32.1 ± 1.3 26.5 ± 2.3 34.3 ± 1.9 29.6

6 53.4 ± 5.2 NA 33.7 ± 3.4 32.9 ± 3.5 NA 24.1 ± 2.4 NA 19.4 ± 1.4 47.6 ± 13.3 31.8 ± 4.2 45.6 ± 8.2 27.6 ± 3.2 33.7 ± 1.6 32.7

7 35.5 ± 2.7 NA 36.0 ± 0.8 32.2 NA 29.1 ± 1.9 NA 19.9 37.1 ± 1.6 34.8 ± 4.5 38.5 ± 1.0 38.5 ± 3.9 36.5 ± 2.8 37.9

Values represent group mean ± SD; a Significantly different from Control by one-way ANOVA (p < 0.05). Weeks 4-7 of treatment are separated by those F0 females that produced a live litter (Gravida) and those that did not produce a live litter (Nulligravida).

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3.4.3 Fertility and fecundity

In both strains, diets containing 0.1 and 1.3 ppm EE resulted in no productive mating

(Table 3). This is considered likely to have resulted from insufficient nutritional intake, as is the

marked delay in mating and decreased percentage of mating that was also observed. In the

C57Bl/6 mice in the low dose EE group, a single productive mating occurred that resulted in a

small litter with 4 pups. In the low dose EE CD1 group, a modest reduction in fertility and

fecundity was also observed (Table 3). In contrast, no effect on fertility or fecundity in the

C57Bl/6 strain for any dose of BPA was observed (Table 3). In the CD1 strain 0.03 ppm (4

µg/kg/da) and 0.3 ppm (41 µg/kg/da) BPA treatment groups, an increase in latency of mating

and a decrease in productive mating was noted.

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Table 3. Measures of Fertility and Fecundity C57Bl/6

Control EE 0.01 EE 0.1 EE 1.3 BPA 0.03 BPA 0.3 BPA 30 No. Breeding Pairs 5 5 5 5 5 5 5 No. litters born 3 1 0 0 3 3 4 Mating index (%) 100 100 20 100 100 100 100 Fertility index (%) 60 20 0 0 60 60 80 Precoital time (da) 2.0 ± 0.8 3.0 ± 0.0 3.0 ± 0.0 3.8 ± 3.0 2.2 ± 0.8 2.2 ± 1.5 1.8 ± 0.5 Gestation (da) 19.0 ± 0.0 N/A N/A N/A 19.0 ± 0.0 18.7 ± 0.6 19.0 ± 1.0 Pups per litter 7.7 ± 1.2 4.0 0.0 0 8.0 ± 0.0 7.0 ± 0.0 6.8 ± 3.2 % Males 50.0 ± 10.1 50 N/A N/A 50.0 ± 17.68 40.0 ± 18.7 37.2 ± 10.2

CD1 Control EE 0.01 EE 0.1 EE 1.3 BPA 0.03 BPA 0.3 BPA 30

No. Breeding Pairs 5 5 5 5 5 5 5 No. litters born 5 3 0 0 3 3 4 Mating index (%) 100 80 60 40 80 80 100 Fertility index (%) 100 75 0 0 75 75 80 Precoital time (da) 2.6 ± 0.5 2.3 ± 0.5 7.2 ± 3.3 7.3 ± 6.5 4.2 ± 3.3 4.4 ± 3.1 2.2 ± 0.8 Gestation (da) 18.6 ± 0.5 18.5 ± 0.7 N/A N/A 19.0 ± 0.0 19.0 ± 0.0 19.0 ± 0.8 Pups per litter 8.4 ± 3.5 10.0 ± 3.0 N/A N/A 10.0 ± 4.0 10.0 ± 1.7 8.0 ± 4.8 % Males 34.9 ± 12.7 44.0 ± 18.7 N/A N/A 50.5 ± 29.3 45.4 ± 30.0 31.7 ± 19.9 Values represent mean ± SD where relevant; Mating index (%) = No. copulation plug positive females / No. females paired X 100; Fertility index (%) = No. females with live litters / No. copulation plug positive females X 100; Precoital time = days from introduction of mating pairs to observation of copulation plug.

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3.4.4 Necropsy and organ weights

Because of morbidity, the C57Bl/6 mice in the EE treatment groups were euthanized, and

necropsy was performed one to two weeks prior to necropsy of the control or BPA treated

groups. Similarly, one C57Bl/6 female was sacrificed prematurely (~wk 4) due to an undefined

neurological pathology; pyometra was not observed in that individual. Due to the small sample

size, variability of some endpoints, and pathology associated with differences in body weights,

statistical assessments of differences in these endpoints compared to controls are of limited value

in determining impact of treatments on gross organ weights. However, these values clearly

illustrate the dramatic strain specific differences observed in the uterine weights in response to

treatment with 0.1 ppm EE and 0.3 ppm BPA (Table 4). An average 23.5 fold increase in uterine

weight compared to controls was observed in the C57Bl/6 0.1 ppm EE treatment group. In

contrast, the uterine weights for the CD1 0.1 ppm and 1.3 ppm EE treated groups were not

different than the control groups. Spleen weights for the C57Bl/6 0.1 ppm EE treatment group

were significantly increased compared to controls, a result consistent with a substantial immune

response in females with affected uteri (p<0.05). With the exception of uterus and spleen weights

in each of the affected C57Bl/6 treatment groups, there were no additional changes in organ

weights.

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Table 4 Organ Weights C57Bl/6 EE (ppm) BPA (ppm)

Control 0.01 0.1 1.3 0.03 0.3 30

Sample Size 4 3 5 4 4 5 4

Liver 1.53 ± 0.32 1.07 ± 0.02 1.18 ± 0.28 1.61 ± 0.16 1.53 ± 0.15 1.34 ± 0.17 1.39 ± 0.22

Kidney 0.14 ± 0.01 0.12 ± 0.01 0.09 ± 0.01 0.09 ± 0.01 0.13 ± 0.01 0.13 ± 0.01 0.14 ± 0.01

Spleen 0.09 ± 0.00 0.07 ± 0.01 0.17 ± 0.11 0.11 ± 0.03 0.09 ± 0.01 0.09 ± 0.00 0.10 ± 0.02

Heart 0.19 ± 0.02 0.16 ± 0.04 0.10 ± 0.01 0.10 ± 0.01 0.16 ± 0.02 0.21 ± 0.06 0.19 ± 0.05

Brain 0.44 ± 0.01 0.45 ± 0.01 0.39 ± 0.01 0.39 ± 0.01 0.44 ± 0.01 0.46 ± 0.01 0.44 ± 0.01

Bladder 0.02 ± 0.01 0.02 ± 0.01 0.03 ± 0.01 0.04 ± 0.01 0.02 ± 0.01 0.03 ± 0.01 0.02 ± 0.01

Uterus 0.13 ± 0.03 0.16 ± 0.02 2.96 ± 0.59 0.63 ± 0.09 0.12 ± 0.02 1.21 ± 1.23 0.13 ± 0.01

CD1 EE (ppm) BPA (ppm)

Control 0.01 0.1 1.3 0.03 0.3 30

Sample Size 5 4 5 5 4 4 5

Liver 1.7 ± 0.2 2.0 ± 0.3 1.5 ± 0.3 1.8 ± 0.2 2.1 ± 0.4 1.6 ± 0.1 1.9 ± 0.4

Kidney 0.19 ± 0.02 0.20 ± 0.02 0.16 ± 0.02 0.12 ± 0.01 0.20 ± 0.03 0.17 ± 0.02 0.19 ± 0.02

Spleen 0.10 ± 0.02 0.12 ± 0.02 0.09 ± 0.02 0.11 ± 0.02 0.13 ± 0.01 0.09 ± 0.01 0.12 ± 0.03

Heart 0.20 ± 0.04 0.23 ± 0.03 0.13 ± 0.02 0.10 ± 0.01 0.22 ± 0.05 0.18 ± 0.03 0.21 ± 0.03

Brain 0.50 ± 0.03 0.50 ± 0.02 0.48 ± 0.03 0.45 ± 0.02 0.50 ± 0.03 0.49 ± 0.03 0.47 ± 0.03

Bladder 0.02 ± 0.00 0.02 ± 0.01 0.02 ± 0.01 0.02 ± 0.00 0.03 ± 0.00 0.02 ± 0.00 0.02 ± 0.00

Uterus 0.21 ± 0.04 0.23 ± 0.08 0.23 ± 0.09 0.30 ± 0.13 0.25 ± 0.06 0.34 ± 0.13 0.23 ± 0.09

Bold indicates p<0.05 compared to control values.

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3.4.5 Uterine Pathology: C57Bl/6 pyometra

During the course of the study, it was observed that four of the C57Bl/6 females in the

0.1 ppm EE group and one in the 0.3 ppm BPA group (none of which produced a live litter)

developed a moribund phenotype characterized by lethargy and highly distended abdomens.

Upon necropsy, the uterus of each of the affected animals was found to be grossly enlarged and

filled with purulent fluid, indicative of severe pyometra (Fig. 1A). No pyometra was observed in

any CD1 mice from any treatment group, indicating an increased sensitivity of estrogen-induced

pyometra for C57Bl/6 mice.

Histological examination of the pyometic uteri revealed thin remnants of the

myometrium and endometrium. The remaining cells of the endometrium were in a disorganized

and ulcerated layer that was highly necrotized (Fig. 1B-G). Immunostaining for the macrophage

specific antigen F4/80 revealed a high degree of macrophage infiltration associated with the

outer layer of myometrium and localized to the inner border of the endometrial-like cell layer

remnant (Fig. 1C; 1E; 1G).

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Figure 1. Histological and immunohistochemical assessments of C57Bl/6 uterus affected by severe pyometra. (A-E) Representative images of a pyometric uterus from a C57Bl/6 mouse in the 0.3 ppm BPA treatment group. (A) Shown is a photograph of the uterus dissected from the C57Bl/6 female in the 0.3 ppm BPA treatment group that developed pyometra. The luminal space of the uterus was filled with purulent material. The scale bar is 1 cm. (B) Representative photomicrographs of H&E stained sections of the affected uterus, showing extreme metaplasia and a high degree of necrosis of the endometrium, features characteristic of severe pyometra. (C) Shown is a representative section of tissue from the affected uterus stained with a monoclonal antibody raised against the macrophage specific antigen F4/80. (D) Shown are representative sections of a uterus from a C57Bl/6 female with pyometra from the 0.1 ppm EE treatment group stained with H&E or (E) stained with a monoclonal antibody raised against the macrophage specific antigen F4/80. Scale bar for paired images is 100 µm. (F) Shown are representative images from a section of a uterus from a C57Bl/6 female with pyometra from the 0.1 ppm EE treatment group stained with H&E or (G) stained with a monoclonal antibody raised against the macrophage specific antigen F4/80. Scale bars for images are 100 µm. OM, outer myometrium; IM, inner myometrium; E, endometrium.

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3.4.6 Uterine Pathology: effects on uterine morphology and macrophage infiltration

To investigate whether exposures to EE or BPA were associated with uterine pathology

and increased macrophage infiltration, even in the absence of severe overt pyometra, histological

comparison of uteri from C57Bl/6 females in the control group with those from the 0.3 ppm BPA

group without pyometra was performed. In the BPA treatment group, the general structural

composition of the uterus including the myometrium, luminal and glandular epithelium, and

endometrial stroma was intact. However, disorganization in these uterine structures was evident

(compare Fig. 2A with Fig. 2B; 2C). The myometrium contained two distinctive smooth muscle

layers each of proper thickness, and the endometrium contained groups of glands surrounding the

luminal epithelium. However, hypertrophic luminal epithelial cells were evident. In some cases,

cells of the stroma were tightly packed and disorganized (Fig. 2C). Cystic endometrial

hyperplasia (CEH) was evident in the uterus of one of the C57Bl/6 female in the 0.3 ppm BPA

group (Fig. 2C1). Compared to control (Fig. 2A2), a marked increase in F4/80 positive

immunostaining was observed in all uteri of the 0.3 ppm BPA treatment group (Fig. 2 B2; C2).

Quantitative assessment of F4/80 immunopositive cell numbers revealed a statistically

significant 5.3-fold increase in the number of macrophages present in the uterus of C57Bl/6 mice

from the 0.3 ppm BPA treatment group (Fig. 4). There were no detectable differences in the

density of immunopositive cells localized to the endometrium or the myometrium in either

control or 0.3 ppm BPA groups. Comparison of the numbers of F4/80 immunopositive cells in

uteri from primagravida and nulliparous F0 C57Bl/6 females without pyometra, revealed no

differences within a given treatment group.

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Figure 2. Effects of oral exposure to BPA on uterine morphology and macrophage infiltration in C57Bl/6 mice without overt pyometra. Shown are representative photomicrographs of uterine sections from a control female C57Bl/6 mouse (Panels A1-A4) and from two different animals in the 0.3 ppm BPA treatment group without symptomatic pyometra (Panels B1-B4; C1-C4). The uteri of BPA treated animals displayed metaplasia (C1, inset) throughout the various regions of the uterus with glandular disorganization and apparent luminal epithelial hyperplasia (C1, arrowhead). Cystic endometrial hyperplasia was also observed in one uterus (C1; denoted by *). Comparison of the density of F4/80 immunopositive cells present in sections from controls (Panels A2; A4) with those from the BPA treatment groups (Panels B2; B4 and C2; C4) demonstrates increased macrophage infiltration. Scale bars for each high magnification and low magnification photomicrograph is 100 µm.

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To assess whether heightened immune responsiveness with BPA and EE exposure was

specific to the uteri from C57Bl/6 animals that developed pyometra, we also assessed uterine

morphology and the numbers of macrophages present in uterine tissue from CD1 mice (Fig. 3A-

H). In contrast to uteri from C57Bl/6 animals, the overall morphology of the uterus was

generally normal for the CD1 females in the 0.3 ppm BPA (Fig. 3B; E) and 0.1 ppm EE (Fig. 3F-

H) treatment groups. However, CEH was detected in one of four CD1 females in the 0.3 ppm

BPA group, and three of four uteri in the 0.1 ppm EE group. Compared to the C57Bl/6 control

group, increased numbers of F4/80 immunopositive cells were evident in uterine tissues from

CD1 controls (Fig. 3A; D; Fig 4). Similar increases in macrophages were apparent for the 0.3

ppm BPA (Fig 3B; E) and 0.1 ppm EE treated females (Fig. 3F-H). Quantitative comparison of

the numbers of macrophages present in the uteri showed 2.7-fold higher F4/80 positive cells in

CD1 controls compared to C57Bl/6 control females (Fig. 4). Compared to CD1 controls, a

significant increase in the number of F4/80 positive cells was not observed in the uteri of CD1

mice in the 0.3 ppm BPA treatment group (Fig. 4).

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Figure 3. Effects of oral exposure to BPA on uterine morphology and macrophage infiltration in CD1 mice, which did not display pyometra. Shown are representative low and high magnification photomicrographs of uterine sections stained for F4/80 positive macrophages from female CD1 mouse in the control group (A; D), the 0.3 ppm BPA treatment group (B; E) or the 0.1 ppm EE treatment group (F-H). Control for specificity of immunostaining included replacement of primary antibodies with non-specific serum (C1). Immunostaining of sections prepared from spleen served as positive controls (C2). Scale bars represent 100 µm.

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Figure 4. Quantitative comparison of F4/80 immunoreactive cell numbers in the uterus of control and BPA treated C57Bl/6 and CD1 mice. Immunopositive macrophage cell numbers present in the myometrium and endometrium were counted by an observer blinded to treatment and strain. Values indicated are mean cell numbers present in one 40x field of the myometrium and one field of the endometrium of a single section per animal. Error bars represent the SEM. For C57Bl/6, n=4 animals, and for CD1, n=5 animals. The level of significance between values was assessed using a one-way ANOVA. * indicates the level of differences between values of same strain control group and BPA treated group are significant (P< 0.01); # indicates that the level of difference between values for each control group is significant (P< 0.01). No difference was observed between C57Bl/6 and CD1 mice treated with 0.3 ppm BPA.

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3.5 Discussion

It should first be pointed out that the intent of this study was to compare appropriate

concentrations of dietary estrogenic chemicals that would allow reproduction and not to

determine whether or not BPA had specific effects upon reproductive or other endpoints. As a

result, the observed changes in those endpoints (e.g. fertility and fecundity, organ weights, etc.)

must not be over interpreted. However, in the case where food consumption is markedly altered

and reproduction was completely blocked, some interesting and valuable insight to the impact of

these dietary estrogens is possible. A clear and significant differential sensitivity to estrogens

between the CD1 and C57Bl/6 strains was evident. This difference in sensitivity was dose

dependent and extended to an apparent aversion for consuming EE. That effect was dramatic and

statistically significant for two related, though independent, endpoint assessments (body weight

and food consumption), even in a study with limited statistical power. Additionally, the

development of pyometra, a well characterized uterine pathology related to estrogen exposure,

was found to occur in the C57Bl/6 strain but not in the CD1 strain. The relatively increased

sensitivity of the C57Bl/6 strain to impacts of estrogens on endpoints as varied as decreased food

consumption and a uterine immunopathology could be interpreted to suggest that the differential

sensitivity of these strains to estrogens is related to a common fundamental aspect of estrogen

signaling rather than isolated tissue-specific differences between strains. However, the apparent

trend toward decreased fecundity observed in the BPA exposed CD1 mice suggests there could

be increased reproductive sensitivities to some actions of BPA. The phenomenon of strain

differences in endpoint- and tissue-specific responses to estrogens in vivo has been extensively

supported in the EDC literature (Pepling et al. 2010, Brossia et al. 2009, Long et al. 2000,

Spearow 1999, Morozova 1991, Greenman et al. 1977). The effects observed here suggest about

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a 10-fold increase in estrogen sensitivity of the C57Bl/6 mice and support the notion of

differential sensitivity of these experimental animal models. However, statements regarding

strain specific estrogen sensitivity should take into account the specific endpoints analyzed.

In contrast to the relatively well-documented cases in veterinary medicine, estrogen-

induced pyometra has received relatively little attention from researchers investigating the

actions of endogenous estrogens and estrogenic endocrine disrupting chemicals. The first

reported observation of pyometra occurring in response to long periods of estrogen exposure in

laboratory rodents was made by Burrows (Burrows 1935). A similar study indicated that some

strains of rat remained resistant to follicular hormone-induced pyometra; a phenomenon not seen

in other species tested (Zondek 1936). Later studies demonstrated that estrogenic hormones

altered bacterial content of the murine uterus, suggesting that there could be a link between the

ability of estrogens to modify the structure of the uterus and the observed changes in the uterine

microbiome (Weinstein et al. 1943). Studies investigating the actions of the non-steroidal

estrogen, DES, clearly demonstrated strain specific differences in the susceptibility to developing

pyometra. Those strain-related differences may result from direct, estrogen receptor-mediated,

activation of the innate immune response (Calippe et al. 2010, Calippe et al. 2008, Hendry et al.

1997, McLachlan et al. 1980). During long-term DES exposure, Sprague-Dawley rats, but not

ACI rats, developed DES-induced pyometra (Stone et al. 1979). Subsequently, the Brown

Norway rat was identified as sensitive to developing pyometra, which allowed demonstration,

via quantitative trait loci mapping, that the strain specific differences in susceptibility to

developing estrogen-induced pyometra were, in part, genetically controlled (Gould et al. 2005).

The results of this study clearly demonstrate strain differences in susceptibility as dietary

exposure to approximately 15 µg/kg/day EE results in essentially quantitative induction of

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pyometra in C57Bl/6 mice, while the CD1 mice did not develop pyometra at that dose or even at

a 10-fold greater dose. Further, one of the five female mice that consumed diet containing 0.3

ppm BPA, resulting in exposure of about 33 µg/kg/day, developed pyometra. Again, no

pyometra was observed in any CD1 animals or in C57Bl/6 mice maintained on control diet

lacking estrogens. The development of pyometra in a BPA-treated animal is important in that it

is the first report of a weakly-estrogenic xenoestrogen inducing pyometra; this demonstrates that,

in regard to its pathological actions on the uterus, BPA is presumably acting as an estrogen

mimetic. This may be of concern in humans (especially sensitive subpopulations) given that

>90% of Americans tested have measurable levels of BPA in their urine (Ye et al. 2011, Calafat

et al. 2005). It is also notable that the dose of BPA that induced pyometra in this study was

below the dose considered safe for human exposure (50 μg/kg/day), which was based on a

NOAEL of 50 mg/kg/day from rodent dietary studies with a BPA concentration of 1000 ppm

(1982).

To assess the role of the immune response in the differential strain susceptibility to

develop pyometra with exposure to estrogens, we investigated the degree of uterine macrophage

infiltration in each strain. Regardless of treatment, we found high levels of macrophages in the

remnants of the necrotic myometrium and the endometrial-like cells of the pyometric uteri.

Based on those findings we hypothesized that BPA and EE were increasing the basal

inflammation in the uteri and reasoned that BPA-exposed C57Bl/6 mice without overt pyometra

would have an increased immune response, as indicated by increased presence of F4/80 positive

macrophages. Comparison of the numbers of macrophages present in the uteri of control CD1

and C57Bl/6 mice revealed a higher number of F4/80-staining macrophages in CD1 samples,

suggesting that there was an elevated basal level compared to C57Bl/6. However, there was a

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significant increase in macrophage counts in C57Bl/6 mice treated with 0.3 ppm BPA compared

to control with no increase seen in 0.3 ppm BPA treated CD1 mice. Along with increased

numbers of macrophages in the mice that did not develop pyometra, significant structural

differences were observed in the C57Bl/6, but not the CD1 strain. The C57Bl/6 mice exposed to

0.3 ppm BPA displayed stromal and glandular metaplasia, with no structural differences

observed in the corresponding CD1 treatment group. Those differences in uterine phenotype

further support the differential sensitivity to estrogens between the C57Bl/6 and CD1. Thus, it

seems likely that these immunologic and structural differences between the strains indicate a

potential key difference in susceptibility to developing overt pyometra which is related to the

immune response. While the basal (control) number of macrophages in the CD1 uterus was

significantly higher than that in the C57Bl/6 mice, the increase with BPA exposure was very

robust in the C57Bl/6 mice. It would appear that the magnitude of the uterine immune response

represents a possible reason for increased susceptibility in C57Bl/6 mice, and may also explain

the difference in structural response to BPA treatment in C57Bl/6 mice. Therefore, the

heightened responsiveness of the C57Bl/6 animals is considered a probable contributing factor

for the sensitivity of this strain to developing estrogen-induced pyometra. Previous studies in

“sensitive” strains of rat have identified quantitative trait loci associated with estrogen-induced

pyometra; however, there is no understanding of the mechanisms involved with sensitivity or

resistance to pyometra. This pathology may be related to interactions between the uterine

immune response and the uterine microbiome. The results here suggest that pyometra in the

C57Bl/6 strain might serve as a sensitive endpoint for understanding the mechanisms responsible

for the impact of estrogen and estrogenic endocrine disrupting chemicals on immunity. Further,

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the C57Bl/6 strain might serve as a useful model of sensitive subpopulations at risk for

developing immunological disorders related to exposures to estrogenic EDCs.

3.6 Acknowledgements

This work was supported by National Institute of Health grants R01 ES015145, RC2 ES018765,

T32 ES016646, and the University of Cincinnati Center for Environmental Genetics (P30-

ES06096).

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3.7 Supplemental Results

3.7.1 Uterine morphology: myometrial wall thickness

To characterize the effects of BPA or EE exposure on the myometrium, the thickness of

both outer and inner layers was measured in both strains and generations of mice. In adult CD1

mice, BPA exposure did not alter wall thickness (Figure 5A). However, EE exposure dose

dependently decreased wall thickness, with a significant decrease at 1.3 ppm EE (p<0.05; Figure

5B). In adult C57Bl/6N mice, the 0.03 ppm BPA exposure group exhibited a decreased wall

thickness that was not statistically significant, while increasing BPA exposure led to a dose

dependent increase in wall thickness (Figure 5A). A trend toward dose dependent decreases in

wall thickness in EE-exposed mice was also observed (Figure 5B). No C57Bl/6N mice in the 1.3

ppm EE exposure group were included due to the high incidence of pyometra in this group.

Strain had a significant main effect on wall thickness in the BPA-exposed (p=0.0008) and the

EE-exposed (p=0.0098) groups, with the 0.03 ppm BPA exposure groups in each strain being

significantly different (p<0.01).

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Figure 5. Myometrial wall thickness in mice across exposure groups and strain. Myometrial wall thickness for adult CD1 (A) and C57Bl/6N (B) mice are shown. Wall thickness measurements were collected as described in section 2.2.4 of the Methods chapter. Values indicated are the mean wall thickness (µm) per group and error bars represent the SEM. For adult CD1, n ≥ 4 animals. For adult C57Bl/6N, n ≥ 3 animals. Wall thickness for BPA- and EE-exposed groups was compared separately to the control group. The level of statistical significance between values in BPA-exposed adult mice was assessed using a two-way ANOVA, with exposure and strain as the main effects, and Bonferroni post hoc test. The 1.3 ppm EE exposure group was not included in two-way ANOVA analysis due to no C57Bl/6N mice being measured in this group. One-way ANOVA and Dunnett’s multiple comparison post hoc test were used to determine the statistical significance in the EE-exposed adult CD1 mice. * indicates a level of difference between values is significant (p<0.05); ** indicates a levels of differences between values is very significant (p<0.01).

3.7.2 Immune response in uteri of CD1 and C57Bl/6N mice

The percent of F4/80-positive cells was quantified in the endometrium and myometrium

of BPA- or EE-exposed adult CD1 and C57Bl/6N mice (Figure 6). In BPA-exposed adult mice,

endometrial F4/80-positive cells exhibited significant increases in both strains (Figure 6A), with

a significant main effect of exposure (p=0.0004) and a significant interaction between exposure

and strain (p=0.0308) based on two-way ANOVA. In the CD1 strain, a significant increase in

F4/80-positive cells occurred at 30 ppm BPA (p<0.001) compared to control. In the C57Bl/6N

strain, significant increases in F4/80-positive cells occurred at 0.03 ppm BPA (p<0.01), 3 ppm

BPA (p<0.01), and 30 ppm BPA (p<0.01) compared to control. No significant differences in

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endometrial F4/80 counts were observed in EE-exposed adult CD1 or C57Bl/6N mice, but all

groups appear increased compared to control in the C57Bl/6N strain (Figure 6B). However,

exposure had a significant main effect (p=0.0419). When quantifying myometrial F4/80-positive

cells in BPA-exposed mice (Figure 6C), a significant main effect of exposure (p=0.0199) and a

significant interaction between exposure and strain (p=0.0094) occurred. Adult CD1 mice again

had a significant increase at 30 ppm BPA (p<0.05) compared to control, similar to that observed

in endometrial cell counts. In the C57Bl/6N strain, significant increases occurred at 0.03 ppm

BPA (p<0.05), 3 ppm BPA (p<0.01), and 300 ppm BPA (p<0.05) compared to control.

Differences in percent of F4/80-positive cells between strains were observed in the control

(p<0.05) and 30 ppm BPA (p<0.01) groups. A similar difference between strains in the control

group was also observed when quantifying myometrial F4/80-positive cells in EE-exposed adult

mice (p<0.05; Figure 6D). Adult C57Bl/6N mice in the 0.01 ppm EE exposure group had a

significant increase in myometrial F4/80-positive cells compared to control (p<0.05; Figure 6D).

When quantifying F4/80-positive cells in EE-exposed adult mice, strain had a significant main

effect (p=0.0011).

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Figure 6. Quantification of F4/80-positive cells in BPA- or EE-exposed CD1 and C57Bl/6N adult mice. The percent of F4/80-positive cells was counted in the endometrium (A-B) or myometrium (C-D) of BPA- (A, C) or EE-exposed (B, D) adult CD1 and C57Bl/6N mice. Endometrial and myometrial F4/80-positive cells were quantified separately and normalized to either stromal or smooth muscle cells respectively. F4/80-positive and total cells were counted by an observer blinded to strain and exposure group. Values indicated are the percent of F4/80-positive cells per animal and error bars represent the SEM. For adult CD1, n≥7 animals except n=2 animals in the 3 ppm BPA and 0.001 ppm EE groups. For adult C57Bl/6N, n≥3 animals. The level of statistical significance between values was assessed using a two-way ANOVA, in which strain and exposure group were used as main effects, and Bonferroni post hoc test. * indicates a level of difference between values is significant (p<0.05); ** indicates a level of difference between values is very significant (p<0.01). *** indicates a level of difference between values is extremely significant (p<0.001).

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CHAPTER 4

Strain-Specific Induction of Endometrial Periglandular Fibrosis in Mice Exposed During

Adulthood to the Endocrine Disrupting Chemical Bisphenol A

Kendziorski JA and SM Belcher. 2015. Strain-specific induction of endometrial periglandular fibrosis in mice exposed during adulthood to the endocrine disrupting chemical bisphenol A. Reprod Toxicol, 58:119-130. doi: 10.1016/j.reprotox.2015.08.001. PMID: 26307436 

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Strain-Specific Induction of Endometrial Periglandular Fibrosis in Mice Exposed During Adulthood to the Endocrine Disrupting Chemical Bisphenol A

Jessica A. Kendziorski and Scott M. Belcher*

Department of Pharmacology and Cell Biophysics University of Cincinnati College of Medicine

Cincinnati, OH, 45267-0575 *Corresponding Author: Scott M. Belcher Department of Pharmacology and Cell Biophysics University of Cincinnati College of Medicine 231 Albert Sabin Way; PO Box 670575 Cincinnati, OH, 45267-0575 Telephone: 513-558-1721 Email: scott.belcher@uc.edu

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4.1 Abstract

The aim of this study was to compare effects of bisphenol A (BPA) on collagen

accumulation in uteri of two mouse strains. Adult C57Bl/6N and CD-1 mice were exposed to

dietary BPA (0.004-40 mg/kg/day) or 17α-ethinyl estradiol (0.00002-0.001 mg/kg/day) as effect

control. An equine endometrosis-like phenotype with increased gland nesting and periglandular

collagen accumulation was characteristic of unexposed C57Bl/6N, but not CD-1, endometrium.

BPA non-monotonically increased gland nest density and periglandular collagen accumulation in

both strains. Increased collagen I and III expression, decreased matrix metalloproteinase 2

(MMP2) and MMP14 expression, and increased immune response were associated with the

endometrosis phenotype in the C57Bl/6N strain and the 30 ppm BPA CD-1 group. The

association between the pro-collagen shift in increased collagen expression and decreased

MMP2 expression and activity implies that strain differences and BPA exposure alter regulation

of endometrial remodeling and contribute to increased fibrosis, a component of several human

uterine diseases.

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4.2 Introduction

Bisphenol A (BPA) is an endocrine disrupting chemical (EDC) released from

polycarbonate plastics and linings of food and beverage containers that contaminates the

consumed contents of the container (Cooper et al. 2011, Carwile et al. 2009, Krishnan et al.

1993). Consumption of BPA-contaminated foods and beverages is the major route of exposure in

humans, with additional exposure occurring through handling of thermal receipt paper,

application of dental sealants, and transfer from medical equipment (Ehrlich et al. 2014, Calafat

et al. 2009, Pulgar et al. 2000). Supporting widespread and prolonged human exposure to BPA,

measurable levels of BPA metabolites were found in over 93% of human urine samples in the

2003-2004 National Health and Nutrition Examination Survey (Calafat et al. 2005). Because of

the known estrogenic and androgenic endocrine disrupting activity of BPA, results from

numerous in vitro and in vivo studies have demonstrated that development and function of

reproductive tissues, including the uterus, were sensitive to BPA exposures (Caserta et al. 2014,

Peretz et al. 2014, Bredhult et al. 2007, Singleton et al. 2006, Schonfelder et al. 2004, Markey et

al. 2001, Papaconstantinou et al. 2000, Ashby and Tinwell 1998). Estrogenic activity of BPA in

the uterus has been established with the rodent uterotrophic assay, for example (Kendig et al.

2012, Laws et al. 2000), and numerous examples of changes in BPA-responsive gene expression

have been reported from studies in a variety of animal models (Calhoun et al. 2014, Hewitt and

Korach 2011, Hong et al. 2006, Naciff et al. 2002, Diel et al. 2000). Endocrine disrupting actions

of BPA have also been associated with pathological changes in the uteri of BPA-exposed rats

and mice (Kendziorski et al. 2012, Signorile et al. 2010, Newbold et al. 2009, 2007b).

Outside of the reproductive axis, we recently demonstrated that lifelong exposure to as

little as 0.004 mg/kg/day of BPA resulted in functional alterations of the collagen extracellular

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matrix (ECM) in the hearts of male and female CD-1 mice (Belcher et al. 2015). In the uterus,

the collagen ECM plays a dynamic structural and functional role in tissue remodeling of the

cycling uterus and during pregnancy (Gaide Chevronnay et al. 2012), whereas dysregulation of

ECM function and collagen accumulation contributes to endometrial adenocarcinoma, fibroids

(leiomyomas), and endometriosis (Kim et al. 2013, Klemmt et al. 2007, Jussila et al. 2004, Kao

et al. 2003, Stewart et al. 1994). In addition to being associated with pathology of the human

uterus, dysregulation of collagen accumulation and excessive fibrosis is a hallmark of equine

endometrosis, one of the most important causes of infertility in mares (Allen 1993) . Equine

endometrosis is an age-related and irreversible degenerative disease of the uterus characterized

by markedly increased endometrial stromal and periglandular fibrosis (EPF) associated with

“gland nest” structures and is unrelated to human endometriosis (Hanada et al. 2014, Snider et al.

2011, Evans et al. 1998, Ferreira-Dias et al. 1994, Kenney 1978). The severity of EPF is

inversely correlated to successful conception and gestation and is associated with increased rates

of embryonic or fetal foal loss and increased susceptibility to infection (Kenney 1978). The

etiology of EPF is unknown, in part due to the lack of suitable laboratory animal models to

facilitate experimental studies. The presence of EPF has also been noted in canines (Christensen

et al. 2012); however, it is unknown whether a similar pathology characterized by excessive

periglandular collagen accumulation and gland nest formation occurs in humans or laboratory

rodent models.

Accumulation of collagen in the uterus during the female reproductive cycle is a dynamic

and tightly regulated process involving hormonal control of collagen gene expression and matrix

metalloproteinase (MMP) expression and activity (Gaide Chevronnay et al. 2012). The types I

and III are primary collagens expressed in mammalian uterus, with lower levels of collagen types

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IV, V, and VI expressed variably during the cycle (Augsburger and Henzi 2008, Boos 2000,

Aplin et al. 1988, Karkavelas et al. 1988). 17β-Estradiol (E2) can increase mRNA and protein

synthesis of many collagen subtypes in the uterus (Frankel et al. 1988, Dyer et al. 1980, Salvador

and Tsai 1973). Estradiol also plays a critical role in the cyclic remodeling of the endometrium

through dynamic modulation of collagen degradation by tightly regulating MMP expression and

activity (Gaide Chevronnay et al. 2012, Russo et al. 2009, Helvering et al. 2005). The MMP9

and MMP2 proteins are key collagen degrading enzymes present in both latent and active forms

in endometrial tissue of the uterus throughout the reproductive cycle (Gaide Chevronnay et al.

2012). Regulation of MMP2 activity during the estrous cycle of rodents and the menstrual cycle

in humans is in part controlled by MMP14, which activates latent MMP2 (proMMP2) by

proteolytic cleavage (Gaide Chevronnay et al. 2012, Strongin et al. 1995, Sato et al. 1994).

Vascular smooth muscle cells respond to increasing levels of estrogen by induction of MMP14

protein expression and a corresponding increase in MMP2 activity (Grandas et al. 2009). Control

of collagen accumulation also involves inhibition of MMP activity through serum and tissue

inhibitors of metalloproteinases (TIMPs) which, in response to varying levels of E2 and

progesterone, are also differentially expressed during the phases of the estrous and menstrual

cycles (Gaide Chevronnay et al. 2012, Goffin et al. 2003, Zhang and Salamonsen 1997, Hampton

and Salamonsen 1994, Sato et al. 1991). TIMP-1, -2, and -3 are all expressed at high levels in the

uterus throughout the cycle, with TIMP-3 exhibiting the most dynamic spatio-temporal

expression profile (Gaide Chevronnay et al. 2012). While the effects of BPA on reproductive

tissues are well-studied, the impacts of exposures to BPA and other EDCs on collagens, MMPs

and the ECM of most tissues, including the uterus, are poorly understood. Evidence supporting

the possibility that BPA may alter MMP activity leading to dysregulation of collagen dynamics

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in the uterus is limited. However studies in developing mammary gland have identified an

exposure related change in ECM collagen content, and BPA can increase MMP2 and MMP9

expression in ovarian granulosa cells and ovarian cancer cell lines (Ptak et al. 2014, Dominguez

et al. 2008, Vandenberg et al. 2007b).

There are well known differences in the sensitivity of rodent strains to pathologic effects

of estrogenic compounds, including E2, diethylstilbestrol (DES) and BPA. Differences in

sensitivity of a variety of hormonally regulated responses, among them male reproductive

development, prolactin release, oocyte development, and expression of estrogen receptors (ERs)

in uterine epithelium, have been characterized (Pepling et al. 2010, Long et al. 2000, Spearow

1999, Steinmetz et al. 1997, Bigsby et al. 1990, Greenman et al. 1977). Strain-specific responses

to estrogen-like EDCs vary and are dependent on the tissue of interest and the phenotypic or

physiological response being examined. Even substrains derived from the C57Bl/6 line have

been shown to have important genetic and phenotypic differences (Simon et al. 2013, Mekada et

al. 2009). Numerous strain-specific differences in sensitivity of uterine and female reproductive

responses to estrogenic compounds have been reported in regards to uterine weight alterations,

development of pyometra, immune cell infiltration, and onset of puberty, including findings from

our previous work which demonstrated that C57Bl/6N mice were more sensitive to the actions of

17α-ethinyl estradiol (EE) and BPA than CD-1 mice (Kendziorski et al. 2012, Roper et al. 1999,

Griffith et al. 1997, Greenman et al. 1977, Drasher 1955). Based on that differential sensitivity

we hypothesized that the sensitivity to developing collagen-related pathology of the uterus would

likewise differ between the CD-1 and C57Bl/6N strains and that dietary BPA exposure would

alter the regulation of collagen. To examine whether BPA differentially altered collagen

accumulation in the adult uterus, uterine pathology was analyzed in young adult female

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C57Bl/6N and CD-1 mice that were exposed for 12-15 weeks to dietary BPA at doses extending

below the accepted safe exposure level in humans (0.05 mg/kg/day) to the defined no observed

adverse effect level (NOAEL) of 50 mg/kg/day (Kendig et al. 2012, Kendziorski et al. 2012,

FAO/WHO 2011, IRIS 1988). To characterize the differential impacts of these BPA exposures,

histological examination was performed to examine collagen-related uterine pathology, including

presence of EPF, gland nest formation, and collagen accumulation, with phenotypes compared

across exposure groups and between strains. Immunohistochemistry was also used to compare

differences in the distribution of MMP2, MMP14, F4/80, α-smooth muscle actin (α-SMA), and

ERα in the uterus. MMP expression and activity were also assessed to examine whether changes

in collagen degradation were contributing to strain- and exposure-related differences in collagen

accumulation.

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4.3 Materials and Methods

4.3.1 Animal Husbandry and Necropsy

All animal procedures were performed in accordance with protocols approved by the

University of Cincinnati Institutional Animal Care and Use Committee and followed

recommendations of the Panel on Euthanasia of the American Veterinary Medical Association.

Six to seven week old C57Bl/6NHsd and Hsd:ICR (CD-1 Swiss) mice were received from

Harlan Laboratories, Inc. (Indianapolis, IN), group housed five per cage and randomly assigned

to control or dietary exposure groups. Animals were housed in a polycarbonate-free housing

system with single-use BPA-free polyethylene cages and water bottles (Innovive, San Diego,

CA). Sanichip bedding (Irradiated Aspen Sani-chip; PJ Murphy Forest Products Corp, Montville,

NJ) was used to prevent contamination from mycoestrogens in corncob bedding. Sterile drinking

water containing <1% of oxidizable organics was produced from a water purification system

designed to eliminate organic chemical contamination (Millipore Rios 16 with ELIX

UV/Progard 2, Billerica, MA). Animals were fed a defined casein-based phytoestrogen-free diet

(Product # D1010501, Research Diets, Inc.; New Brunswick, NJ) unsupplemented (control or 0

ppm) or supplemented with either BPA (2,2-bis(4-hydroxyphenyl)propane; CAS No. 80-05-7;

Lot 11909; USEPA/NIEHS standard) or EE (1,3,5(10)-estratrien-17α-ethinyl-3,17β-diol; CAS

No. 57-63-6; Batch No. H923; Steraloids Inc.; Newport, RI) that was homogenously

incorporated in the diet at desired concentrations. Final dietary BPA concentrations (0.03, 0.3, 3,

30, and 300 ppm) resulted in doses of 0.004, 0.04, 0.4, 4, and 40 mg/kg/day, and concentrations

of EE (0.0001, 0.001, and 0.01 ppm) resulted in doses of 0.00002, 0.0002, and 0.001 mg/kg/day

(Kendig et al. 2012, Kendziorski et al. 2012). Following a 2 week acclimation to the BPA-free

system, study animals were maintained on assigned control or test diet ad libitum until necropsy

at 19 to 23 weeks of age, for a total exposure period of 12 to 15 weeks. During this time, animals

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were bred and those that produced a litter were included in this study. Progression of estrous

cycles was monitored by morphological analysis of vaginal lavage and necropsy was performed

while in estrus. Estrous stage was confirmed by histological assessment of uterine tissues.

4.3.2 Histology and Immunohistochemistry

Tissue isolation, fixation and preparation were described in detail previously (Kendig et

al. 2012, Kendziorski et al. 2012). Formalin fixed tissues were blocked for sectioning with a

sterile razor blade and washed several times in 70% ethanol prior to automated tissue processing

and embedding in paraffin (Histoplast IM; Richard Allen, Kalamazoo, MI). Uteri were placed in

the block longitudinally to examine the entire length of the uterine horn, as well as the dorsal and

lateral sides of the uterus. Microtome sections were cut at 5 µm thickness from blocks at 4˚C and

placed on positively charged slides for hematoxylin and eosin (H&E) staining and

immunohistochemistry. Standard H&E staining was performed in order to examine tissue

structure and morphology and to conduct histopathological analysis. For immunohistochemistry,

sections were dewaxed in xylene and rehydrated through graded series of ethanol concentrations

into phosphate buffered saline (PBS) for MMP2, F4/80, α-SMA, and ERα immunostaining or

Tris-buffered saline with 0.1% Tween 20 (TBST) for MMP14. Heat-mediated antigen retrieval

was performed at 100oC in 0.01M citrate buffer (pH 6.0). Endogenous peroxidase was blocked

with 3% H2O2 in PBS. Sections were incubated at room temperature (RT) in 3% normal goat

serum in PBS for 1 hour and then incubated overnight at 4oC with either rabbit polyclonal anti-

MMP2 (3.33 µg/mL (1:300); NB200-193, Novus Biologicals; Littleton, CO), rabbit monoclonal

anti-MMP14 (0.5 µg/mL (1:250); ab51074, Abcam; Cambridge, MA), rat monoclonal anti-F4/80

(C1:A3-1; 4 µg/mL (1:250); ab6640, Abcam; Cambridge, MA), rabbit polyclonal anti-α-SMA (1

µg/mL (1:200); ab5694, Abcam; Cambridge, MA), or rabbit polyclonal anti-ERα (MC-20; 1

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µg/mL (1:200); sc-542, Santa Cruz Biotechnology; Dallas, TX). Immunoreactivity was

visualized with 0.05% 3’3-diaminobenzidine by the avidin-biotin peroxidase complex method

(Vector Laboratories; Burlingame, CA). Sections were counterstained in Hematoxylin 2 (Richard

Allen; Kalamazoo, MI) for 30 seconds, dehydrated, and coverslipped using Permount (Fisher

Scientific; Fair Lawn, NJ). Mouse lung tissue was used as a positive control for MMP2 and

MMP14, mouse spleen tissue was used as a positive control for F4/80, mouse heart tissue was

used as a positive control for α-SMA, and rat mammary tumor was used as a positive control for

ERα. Negative staining controls included replacement of primary antibodies with non-specific

serum. Stained sections were examined on a Nikon Eclipse 55i microscope using a DS-Fi1 CCD

camera controlled with Digital Sight Software (Nikon; Melville, NY). Final figures were

generated using Adobe Photoshop (San Jose, CA).

4.3.3 Picrosirius Red Stain and Collagen Assessment

Paraffin-embedded uterine sections were stained with Picrosirius Red (Polysciences;

Warrington, PA) to visualize total collagen (red) with bright field illumination and by

polarization birefringence to assess the thickness and packing density of collagen fibers (Dayan

et al. 1989, Junqueira et al. 1979, Junqueira et al. 1978). Briefly, tissue sections were

deparaffinized, rehydrated, and stained for 8 minutes with Weigert’s hematoxylin (American

MasterTech; Lodi, CA). Stained sections were rinsed with tap water for 5 minutes, incubated for

2 minutes in 0.2% phosphomolybdic acid hydrate, and rinsed in deionized H2O for 30 seconds.

Slides were then stained for 1 hour in Picrosirius Red F3BA solution (1.3% 2,4,6-trinitrophenol,

0.4% Direct Red 80), transferred to 0.1 N hydrochloric acid solution for 2 minutes, washed in

70% ethanol for 45 seconds, dehydrated and then coverslipped. Stained sections were examined

on a Nikon Eclipse 80i microscope equipped with a polarized light attachment and a DS-Fi1

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CCD camera controlled with Digital Sight Software (Nikon; Melville, NY). Final figures were

generated using Adobe Photoshop.

4.3.4 Quantitative RT-PCR

Frozen uterine tissue ≤30 mg from CD-1 control and 30 ppm BPA exposure groups were

homogenized for RNA isolation using an RNeasy Fibrous Tissue Mini Kit (Qiagen; Valencia,

CA). Uterine RNA was converted to cDNA using a High Capacity cDNA Reverse Transcription

Kit (Life Technologies; Grand Island, NY). Predesigned TaqMan RT-PCR assays (Applied

Biosystems; Grand Island, NY) were used for the following genes: Col1a1 (Mm00801632_gH),

Col3a1 (Mm00802296_g1), Mmp2 (Mm00439498_m1), Timp1 (Mm00441818_m1), Timp2

(Mm00441825_m1), and Timp3 (Mm00441826_m1). PCR amplification was performed in a

final volume of 20 µL containing 1x TaqMan expression assay primers, Universal Master Mix

(Applied Biosystems; Grand Island, NY), and 10 ng of cDNA. Amplification was performed in

triplicates on a Step One Plus Real-Time PCR System (Applied Biosystems; Grand Island, NY)

under the following fast conditions: 20 seconds at 95oC for one holding stage, then 1 second at

95oC and 20 seconds at 60oC for the cycling stage (60 cycles). Relative expression was

quantified using the ΔΔCt method, using 18s as the endogenous control for normalization.

4.3.5 SDS-PAGE and Immunoblotting

Uterine lysates from frozen tissue were generated by homogenization in 20 mM Tris-HCl

(pH 7.5) with 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, and 1% Triton containing protease

(Roche; Nutley, NJ) and phosphatase (Sigma-Aldrich; St. Louis, MO) inhibitors. Protein

concentrations of cleared lysates were determined using the Bio-Rad Dc protein assay (Bio-Rad;

Hercules, CA). Protein lysate samples (30 µg) were mixed 3:1 with sample buffer (0.25 M Tris-

HCl, 40% glycerol, 8% SDS, 0.01% bromophenol blue, pH 6.8) containing 0.04% β-

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mercaptoethanol and incubated at 96oC for 5 minutes. Protein was loaded onto 10%

polyacrylamide gel and resolved by electrophoresis. Proteins were electrotransferred to an

Odyssey® nitrocellulose membrane (Li-Cor Biosciences; Lincoln, NE) for 16 hours at 4oC.

Membranes were stained with 0.1% Ponceau S in 1% acetic acid to confirm homogenous protein

transfer, destained in water and then blocked with 5% non-fat dry milk (Blotting Grade Blocker,

Bio-Rad; Hercules, CA) in TBST for 1 hour at RT. Membranes were incubated overnight at 4oC

with either rabbit polyclonal anti-MMP2 (0.25 µg/mL (1:4000); NB200-193, Novus Biologicals;

Littleton, CO) or rabbit monoclonal anti-MMP14 (0.066 µg/mL (1:2000); ab51074, Abcam;

Cambridge, MA). Blots were coincubated with a rabbit monoclonal anti-β-actin antibody (0.02

µg/mL (1:2000); 8457, Cell Signaling; Danvers, MA) for normalization of loading. Membranes

were washed and incubated with IRDye® 800CW goat anti-rabbit IgG (0.2 µg/mL (1:5000); Li-

Cor Biosciences; Lincoln, NE) for 1 hour at RT. Membranes were imaged using an Odyssey®

CLx Infrared Imaging System (Li-Cor Biosciences; Lincoln, NE) and immunoreactive bands

were visualized and analyzed with Image Studio Lite Software (Li-Cor Biosciences; Lincoln,

NE).

4.3.6 Gelatin Zymography

Methods for gelatin zymography were adapted from previously described protocols (Toth

et al. 2012, Belcher et al. 2005). Frozen uterine tissue was homogenized in 0.025 M Tris-HCl

(pH 7.5), 0.1 M NaCl, 1% Nonidet P-40 containing EDTA-free Protease Inhibitor Cocktail

(Roche; Nutley, NJ) and cleared by centrifugation at 4oC. Protein concentration was determined

using the Bio-Rad Dc protein assay (Bio-Rad, Hercules, CA). Equal amounts of total protein (30

µg) were mixed 3:1 with sample buffer (0.25 M Tris-HCl, 40% glycerol, 8% SDS, 0.01%

bromophenol blue, pH 6.8) and incubated at RT for 15 minutes. Protein was resolved onto native

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10% polyacrylamide gel containing 0.1% gelatin. Following electrophoresis, gels were washed

with gentle agitation in 2.5% Triton X-100 at RT for 30 minutes, rinsed in dH2O, and then

washed for an additional 30 minutes at RT with gentle agitation in developing buffer (0.05 M

Tris-HCl (pH 7.8), 0.2 M NaCl, 0.005 M CaCl2). Washed gels were then incubated in fresh

developing buffer at 37oC for 16 hours, stained with 0.5% Coomassie Brilliant Blue R-250 in 5%

methanol and 10% glacial acetic acid at RT for 3 hours, and then destained at RT for consecutive

washes of 15 minutes, 30 minutes, and 1 hour with 5% methanol and 10% acetic acid.

Conditioned media from the human brain tumor cell line DAOY (ATCC HTB-186) was used as

a positive control for MMP activity. In control experiments, the addition of EDTA (1 mM final

concentration) to lysates or developing buffer resulted in loss of proteolytic activity at the

expected molecular weights. Gels were imaged using an Alpha Innotech Multiimage II (Alpha

Innotech; San Leandro, CA) and optical density of cleared bands denoting areas of proteolytic

activity were quantified using Image Studio Lite Software.

4.3.7 Statistical Analysis

Pathology was assessed by an observer blinded to exposure and strain with statistical

differences analyzed using Fisher’s exact test. Gland nest density was analyzed using a two-way

ANOVA. Differences between control and exposure groups were analyzed using Dunnett’s

multiple comparison test and a Student’s t-test was used for analysis of mRNA and protein

expression and protein activity. Significance between differences in values was defined as

p<0.05. All data was analyzed using GraphPad Prism® v5/v6 software (GraphPad; La Jolla,

California).

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4.4 Results

4.4.1 Uterine Pathology: Gland Nests and Periglandular Fibrosis in C57Bl/6N and CD-1 Mice

A pathological assessment of a longitudinal section of the uterine horn between the

oviduct and the cervix was conducted. The number of animals presenting with each pathology is

summarized in Table 1. The uteri of control C57Bl/6N and CD-1 mice were characterized by a

low incidence of uterine cysts. Uterine cysts were defined as hyperplastic glands surrounded by a

thin layer of cuboidal epithelial cells. Distension of the glands and increased stromal

proliferation was commonly observed (Table 1).

Table 1. Incidence of Uterine Pathology in CD-1 and C57Bl/6N Mice at 19-23 Weeks of Age

BPA (ppm) EE (ppm)

CD-1 0 0.03 0.3 3 30 300 0.0001 0.001 0.01

n= 12 9 10 10 8 10 10 10 10

Gland Nests 1 (8%) 0 2 (20%) 0 6 (75%)a 3 (30%) 4 (40%) 0 0

Distended Glands 7 (58%) 7 (78%) 6 (60%) 6 (60%) 6 (75%) 8 (80%) 8 (80%) 8 (80%) 8 (80%)

Cysts 1 (8%) 1 (11%) 2 (20%) 2 (20%) 1 (13%) 0 0 0 0

C57Bl/6N

n= 7 7 7 4 6 3 4 3 6

Gland Nests 5 (71%) 6 (86%) 5 (71%) 4 (100%) 5 (83%) 3 (100%) 3 (75%) 2 (67%) 4 (67%)

Distended Glands 6 (86%) 5 (71%) 4 (57%) 3 (75%) 4 (67%) 2 (67%) 1 (25%) 2 (67%) 2 (33%)

Cysts 0 2 (29%) 3 (43%) 2 (50%) 0 1 (33%) 1 (25%) 1 (33%) 1 (17%)

a p = 0.0044 by two-tailed Fisher's exact test

Gland nests, characterized by multiple uterine glands surrounded by collagen producing

stromal cells with resulting periglandular fibrosis and marked collagen accumulation (Fig. 1

arrows), were frequently observed in C57Bl/6N control and in each of the C57Bl/6N BPA and

EE exposure groups (Table 1; Fig. 1). Compared to control, an increased intensity of red/yellow

and decreased green polarization birefringence, characteristic of increased collagen fiber

thickness and density, was associated with periglandular regions of stromal cells in the 30 ppm

BPA and 0.01 ppm EE exposure groups in C57Bl/6N mice (Fig. 1B; D; F).

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Figure 1. Effects of oral exposure to BPA or EE on uterine morphology and collagen accumulation in C57Bl/6N mice. Shown are representative photomicrographs of H&E- and Picrosirius Red-stained uterine sections from a control C57Bl/6N mouse (A-B) and from a mouse exposed to 30 ppm BPA (C-D) or 0.01 ppm EE (E-F). Similar uterine locations are highlighted in each panel. Arrows designate gland nests. Scale bar represents 50 µm.

In contrast to C57Bl/6N controls, only a single gland nest-like structure was observed in

one uterus from the CD-1 control group (Table 1). A significant increase in gland nests (Table 1;

Fig. 2C, arrows) and extensive periglandular fibrosis was observed in the CD-1 30 ppm BPA

exposure group (Fig. 2D) compared to control (Fig. 2A). In contrast to controls, intense

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red/yellow polarization birefringence, forming net-like patterns in areas between gland nest

structures, was characteristic of the CD-1 30 ppm BPA exposure group (Fig 2D). Gland nests

were not present in the phenotypically estrogenized uteri of CD-1 mice exposed to 0.01 ppm EE

(Fig. 2E-F), and notably lower levels of red/yellow and higher levels of green polarization

birefringence were observed, suggesting a less densely packed collagen (Fig. 2F).

Figure 2. Effects of oral exposure to BPA or EE on uterine morphology and collagen accumulation in CD-1 mice. Shown are representative photomicrographs of H&E- and Picrosirius Red-stained uterine sections from a control CD-1 mouse (A-B) and from a mouse exposed to 30 ppm BPA (C-D) or 0.01 ppm EE (E-F). Similar uterine locations are highlighted in each panel. Arrows designate gland nests. Scale bar represents 50 µm.

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Figure 3. Quantification of gland nest density in the uteri of control and BPA-exposed CD1 and C57Bl/6N mice. Comparison of gland nest density between the control groups of CD1 and C57Bl/6N mice (A), control and BPA-exposed CD1 mice (B), and control and BPA-exposed C57Bl/6N mice (C). The number of gland nests was counted by an observer blinded to strain and exposure group. Values indicated are the number of gland nests per mm2 per animal. Error bars represent the SEM. For C57Bl/6N, n≥3 animals and for CD1, n≥8 animals. The level of statistical significance between values was assessed using a two-way ANOVA, in which exposure and strain were the main effects. * indicates a level of differences between values is significant (p<0.05); ** indicates a level of differences between values is very significant (p<0.01). 

 

The density of gland nests in the endometrium for each strain and exposure group was

calculated (Figure 3). Two-way ANOVA of gland nest density indicated significant main effects

of both strain [F(1,77) = 26.99, (p<0.0001)] and exposure [F(5,77) = 4.411, (p=0.0014)] with no

significant interaction between factors [F(5, 77) = 1.493, (p=0.2019)]. The mean density of gland

nest in the endometrium of control C57Bl/6N uteri (M = 0.895 ± SD 0.927) was 9 times greater

than the density observed in the CD-1 strain (M = 0.101± SD 0.349; Fig. 3A). Gland nest density

of neither C57Bl/6N [F (3, 15) = 1.568, (p=0.2385)] nor CD-1 uteri [F (3, 38) = 0.6947,

(p=0.5610)] was affected by EE exposure. Exposure to 30 ppm BPA significantly increased

endometrial gland nest density in the CD-1 strain (Fig. 3B) and significant increases in density

were detected in 0.03 ppm BPA group and in the two highest BPA groups in the C57Bl/6N strain

(Fig. 3C). The increased density detected in the lowest exposure group and the dose dependent

increase in gland nest density observed across the 0.3 to 300 ppm dose range is suggestive of a

non-monotonic dose response relationship. The significant increase in gland nest density in the

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30 ppm BPA CD-1 mice correlated with the increased number of animals observed to have gland

nests in Table 1. In C57Bl/6N mice, however, gland nest density also increased in response to

BPA whereas the number of animals with gland nests was not changed (Table 1).

Figure 4. Relative expression levels and activity of mRNA and proteins involved in collagen synthesis and degradation. Quantitative RT-PCR analysis between control (0) and BPA-exposed (30) CD-1 mice for Col1a1, Col3a1, Mmp2, Timp1, Timp2, and Timp3 (A). The relative differences in gene expression between control and exposed groups were determined by the ΔΔ cycle threshold method with specific TaqMan assays. Representative Western blots for proMMP2 and MMP2 (B) and proMMP14 and MMP14 (C) are shown for control CD-1 and C57Bl/6N mice and 30 ppm BPA-exposed CD-1 mice. Relative proMMP2 and MMP2 protein expression levels (D) and activity (E) and protein expression levels of proMMP14 and MMP14 (F) are shown. Protein expression levels are based on Western blot analysis and activity is based on gelatin zymography as described in Methods. Results are expressed as mean of group ± SEM. Expression and activity of BPA-exposed CD-1 mice and control C57Bl/6N mice is graphed in relation to control CD-1 results for all experiments. Control CD-1 mice were compared to BPA-exposed CD-1 mice and control C57Bl/6N mice separately. For qRT-PCR, n≥4 animals. For protein expression and activity, n≥4 animals for CD-1 mice and n≥3 animals for C57Bl/6N mice. The level of statistical significance for differences between mean values of control and BPA-exposed groups was determined by Student’s t test for all experiments and is indicated by * (p<0.05).

4.4.2 Analysis of Collagen and MMP mRNA Expression

The extensive periglandular fibrosis and increased levels of collagen observed in the CD-

1 30 ppm BPA exposure group were suggestive of increased collagen expression and/or

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alterations in regulation of collagen fibril accumulation. The relative expression levels for

Col1a1, Col3a1, Mmp2, Timp1, Timp2, and Timp3 mRNA in uterine tissues from CD-1 mice in

the control (0) and 30 ppm BPA (30) exposure groups were quantified using qRT-PCR (Fig.

4A). Compared to control, significant increases in Col1a1 (p=0.0379) and Col3a1 (p=0.0161)

and a significant decrease (p=0.0162) in Mmp2 mRNA expression levels were detected in uteri

from CD-1 mice in the 30 ppm BPA exposure group. While Timp1 was not altered by BPA

exposure, Timp2 was significantly decreased (p = 0.0013) in the 30 ppm BPA exposure group

compared to control. Because of its broad inhibitory spectrum and more variable regulation in

the uterus (Gaide Chevronnay et al. 2012), Timp3 expression was also compared in the uterus of

control and 30 ppm BPA exposure groups and was found to be unchanged by BPA exposure.

4.4.3 Analysis of Protein Expression and Activity of MMPs

Western blot analysis was used to determine if protein expression of MMP2 (Fig. 4B)

and MMP14 (Fig. 4C) in uteri from the C57Bl/6N control group and CD-1 30 ppm BPA

exposure group were decreased compared to unexposed CD-1 controls, because these groups

displayed significant differences in gland nest formation and collagen accumulation compared to

CD-1 control animals. Relative to unexposed CD-1 controls, decreased proMMP2 and MMP2

protein levels were detected in both CD-1 30 ppm BPA and unexposed C57Bl/6N groups (Fig.

4B). Quantification found the decrease in expression of proMMP2 (t(9) = 3.014, p=0.0073) and

MMP2 (t(8) = 2.397, p=0.0073) in CD-1 30 ppm BPA and proMMP2 (t(7) = 3.404, p=0.0057)

and MMP2 (t(6) = 1.971, p=0.0481) in C57Bl/6N controls was significant (Fig. 4D). The

decreased expression of MMP2 was reflected by relative decreases in MMP2 gelatinase activities

of the CD-1 30 ppm BPA exposure group and C57Bl/6N control group as compared to

unexposed CD-1 controls (Fig 4E). The relative decreases in proMMP2 (t(10) = 5.486,

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p=0.0003) and MMP2 (t(6) = 4.610, p=0.0037) activity in the CD-1 30 ppm BPA exposure

group and proMMP2 (t(9) = 5.789, p=0.0003) and MMP2 (t(5) = 6.357, p=0.0007) activity in

C57Bl/6N controls were significant (Fig. 4E). As expected (Gaide Chevronnay et al. 2012), the

relative activity of MMP9 in C57Bl/6N and CD-1 uterus were not significantly different (t(4) =

0.0906, p=0.9322). Decreased levels of proMMP14 and MMP14 protein levels were also

detected in both CD-1 30 ppm BPA and unexposed C57Bl/6N groups (Fig. 4C). Expression of

proMMP14 in CD-1 30 ppm BPA (t(9) = 2.056, p=0.0350) and C57Bl/6N control (t(7) = 4.367,

p=0.0016) groups was significantly decreased. While it appeared that relative expression of

MMP14 was also decreased in the C57Bl/6N control group (t(7) = 1.481, p=0.0910), results

were not statistically significant due to the high levels of variance associated with quantifying the

more diffusely migrating MMP14 immunoreactive band observed in the CD-1 controls (Fig. 4C;

F).

4.4.4 Immunolocalization of MMP2 and MMP14 in the Endometrium

In the endometrium of the CD-1 control group, intense positive staining for MMP2 was

found throughout the stroma, as well as in the cytoplasmic compartment of luminal and

glandular epithelial cells (Fig. 5A). Intense staining of stromal cells was more variable with

moderate to high intracellular staining and more diffuse staining in extracellular regions (Fig.

5A). Expression did not appear altered in stroma closer to the glandular or luminal structures.

In C57Bl/6N controls, the cell specific staining pattern in the endometrium was similar,

but comparatively less intense (Fig. 5B). Immunoreactivity for MMP2 was also weaker in the

cytoplasm of glandular and luminal epithelial cells, reflecting the overall decreased expression

levels of MMP2 in the C57Bl/6N uterus. Intracellular MMP2 staining of stromal cells was

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generally weak and further reduced to background levels in the periglandular stroma surrounding

the gland nest (Fig. 5B; arrows).

Immunoreactivity for MMP14 was weak to moderate in the intracellular compartment of

stromal cells in endometrium of both CD-1 and C57Bl/6N controls. In the CD-1 endometrium,

mainly scattered stromal cells surrounding glands were MMP14 immunopositive, and glandular

and luminal epithelial cells were immunonegative (Fig. 5F). In the C57Bl/6N endometrium,

intracellular immunostaining was weak to moderate and more diffusely granular in appearance.

Notably less intense periglandular staining was typical (Fig. 5G; arrow). Weak to moderate

immunostaining was detectable in the borders of luminal epithelial cells.

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Figure 5. Strain differences in the immunohistochemical localization of MMP2 and MMP14 in the uteri of control CD1 and C57Bl/6N mice. Shown are representative photomicrographs of uterine sections stained for MMP2 in CD1 (A) and C57Bl/6N (B) mice and for MMP14 in CD1 (F) and C57BL/6N (G) mice. Arrows in B and G point to periglandular areas with decreased staining. Control for specificity of immunostaining included the replacement of primary antibody with non-specific goat serum for MMP2 (C) and MMP14 (H). Mouse lung tissue was used as a positive control for immunostaining for MMP2 (D) and MMP14 (I), with the replacement of primary antibody with non-specific goat serum for MMP2 (E) and MMP14 (J) used to control for specificity of staining. Scale bar represents 50 µm.

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Figure 6. Immunohistochemical characterization of endometrosis-like phenotype between BPA exposure groups or strain. Shown are representative photomicrographs of uterine sections from 0 ppm CD-1 mice (A-C), 30 ppm BPA CD-1 mice (D-F), and 0 ppm C57Bl/6N mice (G-I) stained for F4/80 (A, D, G), α-SMA (B, E, H), or ERα (C, F, I). Control for specificity of immunostaining included the replacement of primary antibody with non-specific goat serum for F4/80 (A, inset), α-SMA (B, inset), and ERα (C, inset). As a positive control for immunostaining, mouse spleen tissue was used for F4/80 (D, inset), mouse heart tissue was used for α-SMA (E, inset) and rat mammary tumor was used for ERα (F, inset). Scale bar represents 50 µm.

4.4.5 Immunohistochemical Characterization of an Endometrosis-Like Phenotype

Immunohistochemistry using antibodies against F4/80, α-SMA, and ERα was performed

on uterine sections from the 0 ppm CD-1, 30 ppm BPA CD-1, and 0 ppm C57Bl/6N groups to

further characterize the effects of BPA exposure or strain on the observed endometrosis-like

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phenotype (Fig. 6). An increase in F4/80 positive macrophages, indicative of a generally

increased immune response, surrounding gland nests and throughout the stroma was found in the

30 ppm BPA CD-1 exposure group (Fig. 6D) compared to the 0 ppm CD-1 group (Fig. 6A). In

the 0 ppm C57Bl/6N group, higher levels of F4/80 positive macrophages were observed in the

stroma surrounding gland nests (Fig. 6G) as compared to the stroma surrounding glands in the 0

ppm CD-1 group. No staining for α-SMA was observed in periglandular stromal cells

surrounding gland nests in the 30 ppm BPA CD-1 exposure group (Fig. 6E) or the 0 ppm

C57Bl/6N group (Fig. 6H). Endothelial cells surrounding blood vessels in the stroma were α-

SMA immunopositive as expected. Staining for ERα was localized to the nuclear regions in the

glandular epithelium or stromal cells and relative levels did not appear altered by BPA exposure

(Fig. 6F) or strain (Fig. 6I) compared to 0 ppm CD-1 (Fig. 6C).

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4.5 Discussion

Bisphenol A is an EDC with known pathophysiologic activity in the uterus; little

however is known about the relationship between BPA exposure and fibrosis, a common feature

of several uterine pathologies. In this study, the presence of periglandular-associated stromal

cells and fibrosis surrounding clusters or “nests” of endometrial glands, a hallmark of equine

endometrosis, was found to commonly occur in the uterus of C57Bl/6N mice. To our knowledge,

this is the first report of an endometrosis-like phenotype in the murine uterus. The histological

features of the observed EPF and gland nest phenotype suggests that the C57Bl/6N strain may be

a useful “off the shelf” experimental model for understanding the etiology and pathogenesis of

progressive increases in periglandular fibrosis and chronic endometrial degeneration in horses.

The decreased reproductive efficiencies and loss of fertility caused by equine endometrosis in

mares suggests that the endometrosis-like phenotype may play a role in the decreased average

litter size characteristic of C57Bl/6N (6.5 pups) compared to CD-1 mice (11 pups) (Harlan

2009a, 2009b, Evans et al. 1998). Whereas age is the most important factor related to onset of

equine endometrosis in mares (Hanada et al. 2014, Snider et al. 2011), the observed increases in

density and incidence of gland nests, as well as increased periglandular fibrosis resulting from

BPA exposure in both strains of mice suggest that genetic and endocrine disruptive effects

contribute to alterations in stromal collagen accumulation and the development of endometrosis.

The horse and the mouse have distinct similarities and differences in regards to uterine

biology that need to be taken into account when considering the mouse as a model to study a

disease mainly found in equine populations. Adenogenesis in both species continues postnatally

and can be inhibited by progesterone treatment (Cooke et al. 2012b, Filant et al. 2012,

Gerstenberg and Allen 2000). In both species, adenogenesis results in simple tubular gland

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structures (Gray et al. 2001); however, glands eventually branch in mares (Gerstenberg and

Allen 2000), whereas they remain simple tubular structures in mice. Regarding estrous cyclicity,

the two species display distinct patterns. While mice are polyestrous throughout the year,

cyclicity in mares is seasonal, occurring from late April to August. Outside of those months,

mares remain in a state of anestrus (Aiello 2014, Sendel 2010). In order for parturition to occur

in both species, glands and their secretions are crucial for blastocyst implantation,

decidualization, and fetal survival as implantation does not occur in mares until day 35

postovulation (Filant and Spencer 2014, Gray et al. 2001), with implantation occurring on day

4.5 in mice (Wang and Dey 2006). Postpartum, the uterus involutes and returns to normal

cyclicity within ten days for both species (Stewart et al. 2011, Gray et al. 2001, Gomez-Cuetara

et al. 1995). Differences aside, the important similarities in uterine biology between the two

species suggest that the mouse may be a useful model to study equine disease.

In response to BPA exposure, gland nest density in both mouse strains examined here

showed a non-monotonic dose response relationship commonly observed for EDCs. These

complex dose-response relationships can be a result of several separate, yet simultaneously

acting, mechanisms of action (Vandenberg et al. 2012). Estrogens and BPA are known to act

with numerous receptor mediated pathways, either through classical nuclear receptor or rapid

signaling mechanisms (Cookman and Belcher 2014, Belcher 2008). While BPA has estrogen-

like activity at ERα and ERβ at nM concentrations (Wetherill et al. 2007), it can also bind to and

inhibit the androgen and thyroid hormone receptors at µM concentrations (Teng et al. 2013,

Moriyama et al. 2002, Paris et al. 2002, Sohoni and Sumpter 1998) to cause oppositional effects

at low and high dose ranges. BPA can also elicit multiple responses through binding of a single

receptor, as shown in developing neurons and rat cardiomyocytes (Liang et al. 2014, Belcher et

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al. 2005) and partial agonist-like activity of BPA at ERα in the uterus is well established. For

example BPA can increase progesterone receptor (PR) expression in the immature rat uterus, but

also antagonizes E2-induced stimulation in PR expression (Gould et al. 1998). Further study is

required to elucidate the nature of the multiple regulatory mechanisms responsible for the

differential effects of BPA on gland nest density that were observed at low and high BPA doses,

and the nature of the baseline differences observed between the CD-1 and C57Bl/6N strains of

mice.

Excessive collagen accumulation and fibrosis in endometrial tissue is associated with the

progression of numerous diseases of the human uterus, most notably leiomyomas. Leiomyomas,

or uterine fibroids, are one of the most common benign pathologies to affect women of

reproductive age (Baird et al. 2003). The connection between fibrosis and alterations in ECM

signaling in uterine fibroids is well established (Leppert et al. 2004). During the proliferative

phase, mRNA for collagen I and III is overexpressed in uterine fibroids compared to surrounding

tissue, suggesting that ECM components are under hormonal control within the fibroid (Stewart

et al. 1994, Brandon et al. 1993). Exposure to BPA and the development of leiomyomas has also

been experimentally established, since exposure led to the presence of leiomyomas in murine

uteri at 18 months of age (Newbold et al. 2007b), and in vitro studies in human UL uterine

fibroid cells found that BPA increased proliferation and increased mRNA and protein expression

of ERα, IGF-1 and VEGF (Shen et al. 2014). The marked increase in collagen accumulation

induced by BPA in our study further supports the possibility that BPA exposure may contribute

to fibrotic uterine diseases and defined dysregulation of collagen accumulation dynamics is a

plausible mechanism of action. It is notable that the doses of BPA that led to alterations in

collagen accumulation observed here are below the defined NOAEL for BPA and that increases

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in gland nest density in the C57Bl/6N strain occurred at exposures below what is considered safe

for humans (1982).

Proteins involved in collagen synthesis and degradation are transcriptionally controlled

throughout the estrous or menstrual cycle (Gaide Chevronnay et al. 2012, Dyer et al. 1980). To

assess the relationship between BPA exposure and changes in collagen synthesis and

degradation, we first quantified mRNA expression of representative genes involved in these

processes at an exposure level (30 ppm) in CD-1 mice that led to clear increases in gland nest

formation and collagen accumulation. While it is known that E2 can increase mRNA levels of

collagens and MMPs in the uterus (Helvering et al. 2005, Frankel et al. 1988), exposure to BPA

resulted in increased Col1a1 and Col3a1 expression and decreased Mmp2 expression. In relation

to collagen synthesis, BPA acted similarly to E2 (Frankel et al. 1988). However, BPA had an

effect opposite of E2 on genes involved in collagen degradation, a finding in support of BPA

having antagonist-like actions that are interfering with the normal function of E2 in the uterus.

These findings point toward an imbalance between collagen synthesis and degradation leading to

an overall fibrotic phenotype.

Protein expression and activity of MMPs were quantified in order to assess the effect of

exposure and strain. Exposure to BPA significantly decreased the expression and activity of

latent and active forms of MMP2 and expression of the latent form of MMP14. The C57Bl/6N

strain also had significantly decreased latent and active MMP2 expression and activity and latent

MMP14 expression compared to the CD-1 strain. These results coincide with the mRNA

expression results, supporting that BPA decreased Mmp2 mRNA expression leading to a

decrease in protein expression. Decreased expression of proMMP14 and a trend toward

decreased expression of the active form add another layer of complexity to the dysregulation of

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collagen degradation in the BPA-exposed CD-1 uteri or the control C57Bl/6N uteri. A lower

level of MMP14 may also lead to a decrease in activation of MMP2, which would contribute to

an overall increase in collagen accumulation in the uterus.

The diffuse MMP2 immunostaining in the stromal compartment is similar to previous

observations in equine endometrosis (Walter et al. 2005). One difference between equine

endometrosis and the pathology observed in the C57Bl/6N and CD-1 strains was a lack of

MMP2 localization to the glandular and luminal epithelial cells in the mouse endometrium. Also

similar to equine endometrosis (Rebordao et al. 2014), an increased immune response associated

with gland nests was observed, which may have a role in the development of the phenotype. In

equine endometrosis, the periglandular stromal cells can differentiate into myofibroblasts that

express α-SMA (Walter et al. 2001). Differences in ERα staining and localization may also

indicate an active or inactive phenotype (Hoffmann et al. 2009b). We did not observe α-SMA

staining in the periglandular stromal cells or differences in ERα staining and localization, further

supporting the interpretation that gland nests present in these young adult mice represent an early

stage of pathology prior to loss of fertility. Thus, the C57Bl/6N mouse may serve as a small

animal model for examining the etiology and progression of equine endometrosis and also may

be useful in determining the role of EDCs in this disease.

We previously found that life-long BPA exposure resulted in sex-specific pathologic

changes in the ECM of the CD-1 mouse heart (Belcher et al. 2015). Along with those sex-

specific responses, differential responses to BPA exposure based on strain or genetic background

have also been reported (Kendziorski et al. 2012, Drasher 1955). In the uterus, BPA or EE can

significantly increase immune responses in C57Bl/6N mice resulting in development of

pyometra, yet this pathology was not observed in CD-1 mice (Kendziorski et al. 2012). Related

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strain-specific responses to BPA have also been reported in other estrogen-responsive tissues

(Wadia et al. 2007, Long et al. 2000, Steinmetz et al. 1997). In this study, a distinct strain

difference in the response to BPA was observed, as well as differences between the control

groups of mice in each strain. Strain differences may help to explain the etiology of

endometrosis. C57Bl/6N control mice already exhibit a high level of gland nests and

periglandular fibrosis, signifying a potentially unrecognized underlying genetic component to

progression of this phenotype, whereas BPA exposure in the CD-1 strain was required to induce

gland nest development through mechanisms presumably involving dysregulation of collagen

accumulation. Potential differences in baseline levels of ovarian hormones, ER and PR

expression, or cellular responsiveness to E2 or progesterone could be playing a role in the

differential development of an endometrotic-like phenotype observed in the CD-1 and C57Bl/6N

strains (Pepling et al. 2010, Roper et al. 1999, Griffith et al. 1997, Bigsby et al. 1990).

In conclusion, BPA may act as a partial ER agonist to inhibit actions of E2 and result in

collagen dysregulation and potentially fibrosis in the uterus. Up until now, little was known

about the impact of BPA exposure on the regulation of collagen in the uterus. This study informs

that BPA exposure increases gland nest formation and stromal and periglandular collagen

accumulation in both CD-1 and C57Bl/6N mouse strains. In relation to collagen synthesis, BPA

acts similarly to E2 by increasing mRNA expression levels of Col1a1 and Col3a1. However,

BPA exposure has the opposite effect on collagen degradation as would be expected from an

exposure to E2. Mmp2 mRNA expression, protein expression levels of MMP2 and MMP14 and

activity of MMP2 were decreased in BPA-exposed CD-1 mice and control C57Bl/6N mice

compared to control CD-1 mice. These decreases in MMP expression and activity may result in

decreased collagen degradation and lead to an accumulation of collagen in the stromal and

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periglandular compartments of the endometrium. Over time, this accumulation may lead to

fibrosis, a component of several uterine diseases that affect humans and animals, suggesting that

exposure to EDCs may play a role in the development or progression of those diseases. This

study is the first report of an endometrosis-like phenotype in the murine uterus, as well as the

first report of an EDC leading to the presence of this phenotype in any animal model. Based on

these results, the C57Bl/6N strain may serve as an experimental animal model for studying the

etiology and progression of equine endometrosis, as well as determining the effects of exposures

to EDCs on this disease and other uterine pathologies.

4.6 Acknowledgements

This work was supported by research grants R03ES023098 and RC2ES018765 from the National

Institute of Environmental Health Sciences (NIEHS) and a predoctoral fellowship awarded to

JAK from the Albert J. Ryan Foundation. The authors would like to acknowledge and thank Dr.

Vinicius Carreira for his assistance with pathological assessment, Ms. Robin Gear for her

guidance on qRT-PCR, Dr. Eric Kendig, Ms., Susie Christie, and Ms. Dana Buesing for their

planning and execution of the oral BPA/EE exposure study, and Dr. Burns Blaxall for his

generous gift of the α-SMA antiserum.

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CHAPTER 5

Effects of Bisphenol A and 17α-Ethinyl Estradiol in Nulligravida Uterus

of CD1 and C57Bl/6N Mice: Whole Life Exposure

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5.1 Introduction

The aims of this study were to examine the uteri of CD1 and C57Bl/6N mice with whole

life exposure to BPA or EE for increases in pathology, collagen dysregulation, and immune

responsiveness. Based on studies in adult mice exposed to BPA or EE (Kendziorski and Belcher

2015, Kendziorski et al. 2012), it was hypothesized that these endpoints would be significantly

and adversely altered in both strains by BPA or EE exposure. Myometrial wall thickness in CD1

mice was not affected by BPA or EE exposure though. Significant age- and exposure-related

increases in distended glands and gland nests were observed in CD1 mice. It appeared that BPA

and EE slightly increased collagen accumulation in the endometrium of CD1 mice; however,

gland nest density was not increased in BPA or EE exposure groups. Significant increases in

macrophage accumulation in the CD1 uterus were observed in BPA and EE exposure groups.

In humans, BPA exposure is ubiquitous and begins in utero. The main route of BPA

exposure in humans occurs orally by ingesting contaminated food or beverages contained in

polycarbonate plastics or food cans with epoxy resin linings, with additional exposure occurring

through handling of thermal receipt paper, application of dental sealants, and transfer from

medical equipment (Ehrlich et al. 2014, Calafat et al. 2009, Carwile et al. 2009, Pulgar et al.

2000, Krishnan et al. 1993). Fetuses and newborns have indirect exposure via placental transfer

to amniotic fluid (Edlow et al. 2012, Nishikawa et al. 2010, Ikezuki et al. 2002, Yamada et al.

2002). Greater than 93% of humans have detectable urinary levels of BPA and metabolites

(Calafat et al. 2008).

Uterine weights for CD1 mice in this study were significantly increased in all EE

exposure groups, and CD1 mice in the 30 ppm BPA group had an increased uterine weight as

well. Vaginal opening (VO) and days to cornification were not altered by BPA or EE exposure

(Kendig et al. 2012). A prenatal or neonatal exposure to BPA led to morphological alterations in

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epithelial cell proliferation or apoptosis and expression of the ERs and PR later in life (Vigezzi et

al. 2015, Mendoza-Rodriguez et al. 2011, Markey et al. 2005). Exposure to BPA and E2

increased myometrial thickness (Papaconstantinou et al. 2000), suggesting that proliferation and

growth of the myometrium are under estrogenic control. Increased incidence of cystic

endometrial hyperplasia, adenomyosis, and leiomyoma were found in CD1 mice prenatally

exposed to BPA (Newbold et al. 2009, 2007b). In the uterus of adult BPA-exposed mice, mice

display differences in the immune response between strains (Kendziorski et al. 2012), prompting

an examination of the effects of whole life exposure on the immune response in the uterus. Due

to the observed increase in leiomyoma and an equine endometrosis-like phenotype in BPA-

exposed mice, as well as significant alterations in collagen synthesis and degradation in the adult

uterus (Kendziorski and Belcher 2015, Newbold et al. 2007b), potential alterations in collagen

accumulation were also studied.

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5.2 Methods

5.2.1 Exposure

As described in section 2.1.1 of Methods, mice were housed in a polycarbonate-free

caging system with excess estrogenic compounds controlled. Adult CD1 and C57Bl/6N mice

were divided equally between control, BPA, and EE groups and exposed through diet to known

concentrations of BPA (0.03, 0.3, 3, 30, and 300 ppm) or EE (0.0001, 0.001, and 0.01 ppm).

Offspring were exposed from gestation day 0 under either PND21, 49, or 90 either indirectly via

placental transfer or directly via diet (Figure 1). Females were sacrificed in estrus when possible

and tissues were collected for to examine morphology and potential pathologies.

Figure 1. Exposure paradigm for sires, dams, and offspring in control and BPA or EE exposure groups.

5.2.2 Myometrial Wall Thickness

Tissue preparation and H&E staining were described previously in sections 2.2.1 and

2.2.2 of Methods. Transverse sections of the H&E-stained uterine tube from CD1 offspring were

measured for myometrial wall thickness. Five random areas at 20x magnification of the outer

edge of the transverse section were imaged by an observer blinded to exposure group. Within

each image, five measurements from the beginning of the inner myometrium to the outer edge of

the outer myometrium were collected. Using a stage micrometer, the ratio of 60 µm/100 pixels

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was measured. Wall thickness measurements were converted from pixels to micrometers (µm)

using the following formula:

X = 60 µm (pixel measurement) 100 pixels

5.2.3 Uterine Pathology

The H&E-stained uteri of C57Bl/6N and CD1 offspring (PND21, PND49, and ~PND90)

were assessed for distended glands, gland nests, and cysts. Distended glands were characterized

by a thinning of the glandular epithelial cells. Gland nests were characterized as groups of glands

that appear clumped together in an oval or circle surrounded by fibrotic stromal cells. Cysts

appeared as circular holes in the endometrium surrounded by a thin layer of epithelial cells. The

presence of the pathology and, when applicable, the number of incidences were noted for each

uterus. The observer was blinded to strain and exposure group.

5.2.4 Assessment of Collagen Accumulation and Immune Response

Picrosirious Red staining was used to visualize collagen accumulation in the uterus of

CD1 offspring, as described in section 2.3.1 of Methods. Immunohistochemistry using a rat

monoclonal anti-F4/80 (C1:A3-1; 4 µg/mL (1:250); ab6640, Abcam; Cambridge, MA) antibody

was performed to visualize and quantify the presence of macrophages in the uterus, as described

in section 2.3.2 of Methods. To quantify the number of F4/80-positive cells in the uterus, five

random images at 40x magnification of myometrium and five random images at 40x

magnification of endometrium in which mostly stroma was visible were collected. For each

image of endometrium and myometrium, the total number of positively stained cells and the total

number of stromal or smooth muscle cells were counted using Photoshop CC and ImagePro Plus

v4.5 (Media Cybernetics; Rockville, MD). The number of positively stained cells was then

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divided by the total number of cells and multiplied by 100 to generate the percent of F4/80-

positive cells in each mouse.

5.2.5 Statistical Analysis

When applicable, the observer was blinded to strain and exposure group. The litter was

the statistical unit used; one animal was selected from each litter as representative of that litter.

Pathology was assessed with statistical differences analyzed using Fisher’s exact test or χ2 test

for trend when assessing across PND. Significance of differences in percentage of F4/80-positive

cells was determined after arc sin transformation. Myometrial wall thickness, gland nest density,

and percentage of F4/80-positive cell counts were analyzed using a one-way ANOVA and

Dunnett’s multiple comparison test. C57Bl/6N offspring endpoints were not statistically

analyzed due to a low sample size for all groups. Outliers were removed based on Grubb’s test.

Significance between differences in values was defined as p < 0.05. All data was analyzed using

GraphPad Prism® v5/v6 software.

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5.3 Results

5.3.1 Uterine Morphology: Myometrial Wall Thickness

The myometrium is made up of two smooth muscle layers involved in protecting the

endometrium and supporting pregnancy and labor by inducing uterine contractions (Thompson

2015). Alterations in myometrial wall thickness may adversely impact these functions. To

determine if BPA or EE exposure affected myometrial thickness, the thickness of both outer and

inner layers was measured in CD1 mice. No differences in wall thickness were observed in BPA-

or EE-exposed CD1 offspring with a whole life exposure (Figure 2). The lack of differences in

myometrial thickness suggests that alterations in uterine morphology by BPA or EE begin at the

cellular level and gross differences are not observed until later in life.

Figure 2. Myometrial wall thickness in CD1 mice across exposure groups. Myometrial wall thickness for CD1 offspring is shown. Wall thickness measurements were collected as described in section 5.2.2. Values indicated are the mean wall thickness (µm) per group and error bars represent the SEM. n ≥ 3 animals. Wall thickness for BPA- and EE-exposed groups was compared separately to the control group. One-way ANOVA and Dunnett’s multiple comparison post hoc test was used to determine the level of statistical significance for BPA-exposed offspring and Student’s t test was used to determine the level of statistical significance in EE-exposed offspring. Statistical significance was set at p < 0.05.

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5.3.2 Uterine Pathology

The presence of distended glands, cysts, and gland nests were noted in the uteri of CD1

mice at PND21, 49, and 90 (Table 1). The presence of gland nests in the uteri of C57Bl/6N

offspring is also summarized in Table 1. At PND21, no animals presented with cysts or gland

nests. Distended glands were absent from control animals and were observed at low frequency in

the 0.03, 30, and 300 ppm BPA groups and the 0.0001 and 0.01 ppm EE groups. At PND49, a

low frequency of mice with distended glands was observed in control and most BPA and EE

exposure groups. In the BPA groups, the number of mice with distended glands followed a non-

monotonic response curve, with the highest incidence at 3 ppm BPA (63%). Cysts were observed

in one CD1 mouse in the 0.3 ppm BPA group and the 0.001 ppm EE group. The frequency of

mice with gland nests in the 0.03 ppm BPA exposure group (75%) significantly increased

compared to the control group (22%; p = 0.0445). All other BPA exposure groups and the 0.0001

and 0.001 ppm EE exposure groups appeared to have a higher frequency of mice with gland

nests compared to the control group as well.

Figure 3 shows representative images of uterine morphology for control (Figure 3A), 30

ppm BPA (Figure 3D), and 0.01 ppm EE (Figure 3G). At PND90, the incidence of mice with

distended glands was high in control (70%) and all BPA (50-85%) and EE (44-75%) groups. At

least one mouse in each BPA or EE group developed a cyst, except for 300 ppm BPA and 0.001

ppm EE (frequency <20%). Gland nests were present in control (20%) and all BPA and EE

groups, with the number of mice at or above 50% in the 0.3 and 300 ppm BPA exposure groups.

In the C57Bl/6N strain at PND90, gland nests were present in almost every animal. These results

provide insight into experiments related to collagen accumulation and normal uterine aging.

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When examining trends across PND in the CD1 strain, the number of mice with

distended glands significantly increased over time in the control (p = 0.0046), 0.03 ppm BPA (p

= 0.0287), 3 ppm BPA (p = 0.0208), 30 ppm BPA (p = 0.0287), 300 ppm BPA (p = 0.0025),

0.0001 ppm EE (p = 0.0273), and 0.001 ppm EE (p = 0.0068) groups. The time-dependent

observed increase in distended glands serves as an internal control due to the increase being well

known (Chapin et al. 2008). The presence of dilated or distended glands also regularly fluctuates

throughout the menstrual or estrous cycle (Rubin and Strayer 2012). No trends were found for

cyst formation over time. The number of mice presenting with gland nests clearly increased over

time, especially between PND21 and PND49. However, between PND49 and PND90, the

number of mice with gland nests did not increase further. Some exposure groups even had a

decrease in the number of mice with gland nests, the most drastic change being in the 0.03 ppm

BPA group (75% at PND49 vs 27% at PND90). In the 30 ppm BPA group, trends for increasing

the number of mice with gland nests over time approached significance (p = 0.0566).

5.3.3 Collagen Accumulation

A slightly increased intensity and dispersal of red/yellow birefringence, indicative of

increased collagen fiber thickness and density, was associated with stromal cells in the BPA-

exposed CD1 offspring (Figure 3F) compared to control (Figure 3C). This increase in stromal

cell-associated red/yellow birefringence was also observed in EE-exposed CD1 offspring, as well

as a slight increase in periglandular-associated green birefringence, suggestive of less densely

packed periglandular collagen (Figure 3I). The observed increase was much lower than that

observed in the adulthood exposure model though (Kendziorski and Belcher 2015).

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Table 1. Uterine Pathology in CD1 and C57Bl/6N Offspring BPA (ppm) EE (ppm)

PND21 CD1 0 0.03 0.3 3 30 300 0.0001 0.001 0.01

n= 5 6 5 7 7 5 5 6 5

Distended Glands 0 (0%) 1 (17%) 0 (0%) 0 (0%) 1 (14%) 1 (20%) 1 (20%) 0 (0%) 2 (40%)

Cysts 0 (0%) 0 (0%) 0 (0%) 0 (0%) 0 (0%) 0 (0%) 0 (0%) 0 (0%) 0 (0%)

Gland Nests 0 (0%) 0 (0%) 0 (0%) 0 (0%) 0 (0%) 0 (0%) 0 (0%) 0 (0%) 0 (0%)

PND49 CD1

n= 9 8 11 8 5 10 10 5 7

Distended Glands 2 (22%) 1 (13%) 4 (36%) 5 (63%) 2 (40%) 2 (20%) 2 (20%) 1 (20%) 2 (29%)

Cysts 0 (0%) 0 (0%) 1 (9%) 0 (0%) 0 (0%) 0 (0%) 0 (0%) 1 (20%) 0 (0%)

Gland Nests 2 (22%) 6 (75%)a 6 (55%) 5 (63%) 2 (40%) 6 (60%) 5 (50%) 2 (40%) 1 (14%)

PND90 CD1

n= 10 11 12 10 14 13 8 11 9 Distended Glands 7 (70%)b 7 (64%)b 6 (50%) 6 (60%)b 9 (64%)b 11 (85%)b 6 (75%)b 7 (64%)b 4 (44%)

Cysts 0 (0%) 2 (18%) 1 (8%) 1 (10%) 2 (14%) 0 (0%) 1 (13%) 0 (0%) 1 (11%)

Gland Nests 2 (20%) 3 (27%) 6 (50%) 3 (30%) 6 (43%)c 7 (54%) 3 (38%) 4 (36%) 3 (33%)

PND90 C57Bl/6N

n= 2 2 3 2 2 1 0 1 1

Gland Nests 1 (50%) 2 (100%) 3 (100%) 1 (50%) 2 (100%) 1 (100%) 0 0 (0%) 1 (100%) a p=0.0445 by one-tailed Fisher's exact test b p<0.05 by χ2 test for trend across all PNDs for increases in distended glands c p=0.0566 χ2 test for trend across all PNDs for increases in gland nests

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Figure 3. Effects of oral BPA or EE exposure on uterine morphology and collagen accumulation in CD1 offspring at PND90. Shown are representative photomicrographs of H&E- and Picrosirius Red-stained uterine sections from a control CD1 mouse (A-C) and from a mouse exposed to 30 ppm BPA (D-F) or 0.01 ppm EE (G-I). Similar uterine locations are highlighted in each panel. Arrows indicate gland nest structures. Scale bars represent 50 µm.

5.3.4 Quantification of Gland Nest Density

Quantification of gland nest density in the endometrium of both strains at PND90 was

performed (Figure 4). In the C57Bl/6N strain, the mean density of the control group (M = 1.138

± 1.138) was ~37 fold greater than the mean density of the control group in the CD1 strain (M =

0.031 ± 0.020). However, due to the low sample number in the C57Bl/6N strain, the results

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cannot be statistically compared (Figure 4A). Both strains had an observed non-monotonic dose

response to BPA exposure. In the CD1 strain, gland nest density was increased at 0.3, 30, and

300 ppm BPA compared to control (Figure 4B), which correlates with increases in the number of

mice with gland nests summarized in Table 1. However, the increases were not statistically

significant. In the C57Bl/6N strain, the two lowest BPA concentrations displayed increased

gland nest density compared to control, while 3 and 300 ppm BPA exposure led to greatly

decreased gland nest density (Figure 4C). A dose-dependent increase in gland nest density was

observed in EE-exposed CD1 mice, but the densities were not significantly altered compared to

control (Figure 4B).

Figure 4. Quantification of gland nest density in the endometrium of BPA- or EE-exposed CD1 and C57Bl/6N mice at PND90. Gland nest density between the control groups of each strain (A), control and BPA- or EE-exposed CD1 mice (B), and control and BPA- or EE-exposed C57Bl/6N mice (C) were compared. Gland nests were counted by an observer blinded to strain and exposure group. Values indicated are the number of gland nests per mm2 per animal and error bars represent the SEM. For CD1, n≥7 animals, and for C57Bl/6N, n≥1 animal. The level of statistical significance between values was assessed using a one-way ANOVA and a Dunnett’s multiple comparison post hoc test.

5.3.5 Immune Response

In the endometrium of BPA-exposed CD1 offspring, a significant increase in F4/80-

positive cells at 30 ppm BPA (p < 0.05) compared to control was quantified in the CD1 strain. A

non-monotonic dose response to BPA exposure and subsequent endometrial F4/80-positive cell

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counts was also present (Figure 5A). No significant differences were observed in the

myometrium of BPA-exposed offspring (Figure 5B). When comparing the endometrial and

myometrial F4/80-positive cell counts, the 30 ppm BPA exposure group was significantly

different between the two tissue types (p < 0.05; Figure 5). In the endometrium of EE-exposed

offspring, the 0.0001 ppm EE group had a significant increase in F4/80-positive cells (p < 0.013)

compared to control in CD1 offspring (Figure 5A). Similar patterns of F4/80-positive cell counts

in the endometrium of EE-exposed offspring were observed in the myometrium of EE-exposed

offspring (Figure 5B). The 0.0001 ppm EE exposure group had a significant increase in F4/80-

positive cells (p < 0.01) compared to control in CD1 offspring (Figure 5B). The 0.001 and 0.01

ppm EE-exposed CD1 offspring appeared to have decreases in the percent of F4/80-positive cells

compared to the control group.

Figure 5. Quantification of F4/80-positive cells in BPA- or EE-exposed CD1 offspring. In CD1 offspring, endometrial (A) or myometrial (B) F4/80-positive cells were counted in BPA- (gray bars) or EE-exposed (black bars) mice. Endometrial and myometrial F4/80-positive cells were quantified separately and normalized to either stromal or smooth muscle cells respectively. F4/80-positive and total cells were counted by an observer blinded to exposure group. Values indicated are the percent of F4/80-positive cells per animal and error bars represent the SEM. n ≥ 4 animals. The level of statistical significance between values was assessed using a one-way ANOVA and Dunnett’s post hoc test. * indicates a level of difference between values is significant (p<0.05); ** indicates a level of difference between values is very significant (p<0.01).

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5.4 Discussion

Bisphenol A is a compound with known endocrine disrupting activity in the uterus.

Through studies on adult CD1 and C57Bl/6N mice, differences in responses between mouse

strains to BPA exposure in regards to development of pyometra and an equine endometrosis-like

phenotype, collagen dysregulation, and immune response were reported (Kendziorski and

Belcher 2015, Kendziorski et al. 2012). In this study, the development of various pathologies

including an equine endometrosis-like phenotype, collagen accumulation, and immune response

to whole life BPA exposure in the offspring of CD1 mice were examined. While C57Bl/6N

offspring were examined for pathology and gland nest density, a small sample size prevented

analysis.

The myometrium is made of distinct outer and inner smooth muscle layers that provide

support for the uterus during pregnancy and parturition. During development, the layers can be

seen as early as PND10 in mice, with the adult configuration observed around PND15. By

PND42, the entire uterus has grown in size and complexity but retains its configuration

(Mehasseb et al. 2009). Treatment or exposure to estrogenic compounds has varying effects on

alterations in myometrial development and thickness. In young adult rats, myometrial thickness

is increased 24 hours after one dose of E2 (Kang et al. 1975), similar to results from E2-treated

or BPA-exposed ovariectomized adult mice (Papaconstantinou et al. 2000). However, prenatal

BPA exposure decreases the volume of myometrium (Markey et al. 2005) or perinatal estradiol

benzoate treatment speeds up uterine development without structural alterations (Mehasseb et al.

2009). These studies indicate that the developmental time point at which exposure occurs is

important for the end result. In this study, whole life BPA or EE exposure had no effect on

myometrial thickness. While morphology and pathology of the endometrium were examined at

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earlier time points (PND21 and 42), normal myometrial structure was not investigated. It is

possible that myometrial development was sped up, as was observed in Mehasseb et al. (2009).

However, even earlier time points than PND21 are necessary to examine to determine if

development did occur at an earlier time point.

Previous studies examining pathology in the rodent uterus prenatally or neonatally

exposed to BPA assess the tissue at 16-18 months of age (Newbold et al. 2009, 2007b). This

study assessed pathology at much younger ages, PND21, 49, and 90. As expected, few

pathological alterations were found at PND21 because endometrial development does not finish

until ~PND15 (Cooke et al. 2013). However, at PND49, at least one CD1 mouse presented with

distended glands or gland nests in control and all BPA or EE exposure groups. The frequency of

CD1 mice with distended glands further increased at PND90, suggesting that this pathological

alteration is a natural occurrence of development, aging, and cyclicity (Rubin and Strayer 2012).

The frequency of CD1 mice with gland nests remained fairly constant between PND49 and 90

and did not further increase with BPA or EE exposure, potentially indicative of gland nest

formation also being a natural occurrence of development. However, when gland nest density

was quantified, the occurrence of gland nests was almost negligible in the CD1 strain in all

exposure groups. The increased frequency of mice with gland nests was more than likely not

physiologically relevant. Offspring in the C57Bl/6N strain had a higher gland nest density than

the CD1 strain, similar to what was observed in the adult exposure model (Kendziorski and

Belcher 2015). When assessing collagen accumulation in the CD1 strain, BPA or EE exposure

did appear to shift stromal collagen to a more sheet-like pattern and a thicker fibril, while

periglandular collagen was largely unchanged. Because transcription of collagen types is

estrogenically regulated (Frankel et al. 1988, Dyer et al. 1980), BPA or EE may be increasing

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transcription and leading to increased collagen accumulation without a high incidence of gland

nests.

In both the whole life and adulthood exposures, the number of F4/80-positive cells

significantly increased in the 30 ppm BPA exposure group of CD1 mice (Figure 6 in Chapter 3),

suggesting that time of exposure does not matter in regards to alterations in immune

responsiveness in the uterus. However, other endpoints examined, including collagen

accumulation, gland nest density, and pathology, were not significantly altered in the whole life

exposure model as they were in the adulthood exposure model. Other than time of exposure, the

main difference is the mating, pregnancy, and parturition that occurred in the adult exposure

model. This difference between exposure models highlights the lack of understanding of the

relationship between estrogenic EDC exposure and potential alterations in the response to drastic

changes in the uterine environment that occur during mating and pregnancy. The presence of

semen or sperm induces an inflammatory response in the uterus (Bromfield 2014, Robertson et

al. 2013, Schuberth et al. 2008, Sharkey et al. 2007); however, it is unknown if the presence of

semen alters collagen regulation. Regulation of collagen and other ECM proteins is significantly

altered during pregnancy though (Tsai et al. 2009, Carbone et al. 2006, Bjorn et al. 1997,

Iwahashi et al. 1996), so the combination of these events may be important for creating an

environment sensitive to potential long-term endocrine disrupting effects.

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CHAPTER 6

SUMMARY AND CONCLUSIONS

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6.1 Significance

This dissertation research highlighted the impact of BPA and EE exposure on alterations

in immune responsiveness and fibrosis, along with the development of uterine pathologies such

as pyometra and an equine endometrosis-like phenotype (Kendziorski and Belcher 2015,

Kendziorski et al. 2012). Changes in immune responsiveness and ECM regulation are also

components of human uterine pathologies, including leiomyoma and endometriosis, and

important factors in infertility, suggesting that exposure to estrogenic EDCs is a contributing

factor to the development of uterine diseases and reproductive dysfunction.

Pyometra is a uterine disorder caused in part by hormonal factors. Exposure to estrogenic

compounds such as E2 and DES led to the development of this disease (Brossia et al. 2009,

Gould et al. 2005, Pandey et al. 2005). This dissertation research was the first report of BPA

exposure leading to the development of pyometra in any animal model (Kendziorski et al. 2012),

supporting that BPA acts as an estrogenic EDC in the uterus. In this study, the oral exposure in

intact female mice can be translated to humans due to the relevant route of exposure and dosing.

A second aspect to the studies presented in this dissertation was related to susceptibility to

pyometra development. Only BPA- and EE-exposed C57Bl/6N mice developed the pathology,

suggesting that genetic variation in humans could also impact sensitivity to estrogenic EDCs and

subsequent development of uterine pathologies. Genome regions involved in susceptibility to E2-

or DES-induced pyometra were known (Gould et al. 2005, Pandey et al. 2005), suggesting a

QTL involved in BPA-induced pyometra may be regulating susceptibility as well.

The observed alteration in immune responsiveness in both the adulthood and whole life

exposure model suggests that BPA exposure may have a detrimental effect on the immune-

related functions in fertility. Successful implantation depends greatly on the contributions of the

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varying immune cells present in the uterus (Gnainsky et al. 2014). Aberrant recruitment and

presence of immune cells in the uterus that occurs with BPA exposure may significantly reduce

the probability of successful implantation. Increased immune cell infiltration and macrophage

activation are also highly involved in the growth of endometriotic lesions (Ahn et al. 2015),

suggesting that BPA exposure can play a role in the progression and severity of uterine disorders

based on altering the immune environment in reproductive tissue.

The results presented in this dissertation highlighted that BPA exposure in adulthood led

to increased collagen accumulation in the uterus of mice due to dysregulation of collagen

synthesis and degradation. In adult CD1 mice, BPA directly increased collagen I and III

expression via transcriptional activation and directly and indirectly decreased MMP2, MMP14,

and TIMP2 expression and MMP2 activity. Exposure groups with observed increases in stromal

and periglandular collagen also had increases in gland nest density, another hallmark of equine

endometrosis (Kendziorski and Belcher 2015). These findings suggest that BPA exposure factors

in the progression of fibrotic human and veterinary uterine diseases by altering collagen

synthesis and degradation, leading to the observed fibrotic phenotype. Estrogens increased the

transcription of collagens and MMPs during the physiological cycling of sex hormones (Gaide

Chevronnay et al. 2012, Frankel et al. 1988). Exposure to BPA led to further transcription of

collagens, increasing the overall levels of collagen. However, BPA exposure did not increase

MMP expression levels, as would be expected with E2 treatment, suggesting that BPA acted

through a different mechanism than E2 to alter collagen degradation. The combination of

increased collagen synthesis and decreased collagen degradation led to fibrosis in the uterus,

potentially increasing the severity of present uterine diseases as well.

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Although collagen synthesis and degradation in the uterus are under estrogenic control

(Gaide Chevronnay et al. 2012, Henriet et al. 2012), no previous studies had been conducted

comparing and contrasting the differences in collagen accumulation between different mouse

strains. Due to the prevalence of gland nest structures and periglandular fibrosis in the C57Bl/6N

uterus, part of this dissertation research examined the differences in collagen accumulation

between the C57Bl/6N and CD1 strains (Kendziorski and Belcher 2015). Differences in

expression of proteins involved in regulating collagen accumulation between the strains are

summarized in Table 1. There was a general decrease in MMPs in the C57Bl/6N strain, leading

to an overall increase in collagen accumulation.

Table 1. Comparison of Relative Expression and Activity of Collagens and MMPs in Mouse Strains

  Expression Activity   Collagen proMMP2 MMP2 proMMP14 MMP14 proMMP2 MMP2

CD1 ‐  −  −  −  −  −  − 

C57Bl/6N ↑ ↓ ↓ ↓ ↓ ↓ ↓

Collagen dysregulation combined with increases in gland nest density are characteristics

of equine endometrosis (Evans et al. 1998, Kenney 1978). Further characterization of the stromal

fibroblasts surrounding gland nests in the equine endometrosis-like phenotype resulted in an

increase in F4/80 staining, but ERα and α-SMA staining in the stroma and glandular epithelium

were unaffected (Kendziorski and Belcher 2015, Hoffmann et al. 2009b, Walter et al. 2001). The

endometrosis-like phenotype observed in the C57Bl/6N strain appeared to be an early stage of

the progressive pathology, suggesting that aging and repeated mating cycles may lead to more of

the destructive degenerative phenotypes observed in equine populations. Under control

conditions, the C57Bl/6N mouse may be useful as an experimental model to study the etiology,

progression, and treatment of equine endometrosis.

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When comparing the two exposure models, mice with a whole life exposure to BPA or

EE did not develop significant alterations in collagen accumulation, increased gland nest density,

or pathology. Other than exposure, distinguishing characteristics were mating, gravid status, and

parturition, with mice in the adulthood exposure having delivered one litter and mice in the

whole life exposure model never mating and were nulligravid. This difference between exposure

models highlights the lack of understanding of the relationship between estrogenic EDC

exposure and potential alterations in the uterine environment that occur during mating and

pregnancy.

6.2 Limitations

While each technique is not without its limitations, the quantification of F4/80-positive

cells could have been performed differently using flow cytometry or more specific antibodies for

in situ hybridization. These techniques would have allowed for distinguishing between M1 and

M2 macrophages, which are differentially activated by various stimuli and have varying roles in

the periphery and tissues (Martinez and Gordon 2014). The techniques could also have been used

to characterize the presence and abundance of other immune cells, such as leukocytes,

neutrophils, and B and T cells. This knowledge would allow for a better understanding of the

specific types of immune cells regulated by BPA or EE exposure and potentially identify early

stages of immune alterations before the induction of pyometra. However, the choice to use a

general macrophage marker (F4/80) was made to examine an initial general immune response.

Further characterization of the specific immune cells involved in this response could be

performed using more sophisticated techniques, such as flow cytometry.

Loss of tissue integrity was another limitation to these studies that needs to be

considered. Depth in the uterine tissue was an important variable that needed to be controlled and

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known. When tissue sectioning, the depth was recorded and sections were collected at a similar

level in the uterus for all animals. However, it was possible that there was still variability

between samples, which could play a role in differences observed between samples. Loss of

tissue integrity was also an important factor in RNA isolation and protein homogenization. While

temperature of the tissue was kept as cold as possible, activity of degrading enzymes could still

have affected integrity of the tissue, leading to potential decreases in the amount of RNA or

protein isolated.

While the endometrium, specifically stromal cells surrounding glandular epithelium, was

the main localized site of interest in the uterus, both the myometrium and endometrium were

contained in protein homogenates used to quantify protein expression and activity. Laser capture

microdissection would be an appropriate technique to isolate the desired cell types of interest.

However, the significant decreases in MMP protein expression and activity observed in BPA-

exposed CD1 mice or control C57Bl/6N mice compared to control CD1 mice suggested that the

decreases in these proteins was so robust between the groups that it was unnecessary to isolate

specific cell types. Also, immunolocalization of MMP2 and MMP14 supported that expression

of these proteins is decreased in periglandular stromal cells.

6.3 Future Directions

With these novel findings regarding immune responsiveness and the development of

immune-related uterine pathology though come new questions regarding the impact of BPA

exposure and genetic variation in these processes. While it was known that estrogenic

compounds, now including BPA, can lead to the induction of pyometra, specific genes and

proteins involved in its etiology are still partially unknown. As described, mapping of

overlapping QTLs named Eutr1 and Eutr2, which include large chromosomal areas containing

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several genes, has been performed for E2- and DES-induced pyometra in rat strains (Gould et al.

2005, Pandey et al. 2005). A third QTL in mice involved in eosinophil infiltration, Est1 (Griffith

et al. 1997), also overlaps with Eutr1 and Eutr2. This overlapping region between three QTL

may contain genes involved in the development of pyometra. There are four genes in the

overlapping region, those encoding IL11Rα, Ccl19, Ccl21a, and Ccl21c, all of which are

involved in the immune system. The impact of exposure to EDCs on expression of these genes

and translated proteins need to be characterized to determine the role of these genes of interest in

immune responsiveness.

Of these four genes, only IL11Rα has been extensively studied in the uterus. In human

endometrium, IL11Rα was found in both the stroma and the apical surface of epithelial cells

(Karpovich et al. 2003) and was expressed in decidual tissue in the rat uterus peri-implantation

(Li et al. 2001) and glandular epithelial cells in canines during pregnancy (Guo et al. 2009). It

was important for proper decidualization and placentation in humans and animals (Menkhorst et

al. 2009, Paiva et al. 2009, Karpovich et al. 2005, Ain et al. 2004, Dimitriadis et al. 2003,

Bilinski et al. 1998, Robb et al. 1998). In women with infertility or recurrent miscarriages, there

were significant decreases in IL-11, the ligand for IL11Rα (Dimitriadis et al. 2007, Linjawi et al.

2004), while women with endometriosis and associated infertility had an absence of IL11Rα

staining in the glandular epithelium (Dimitriadis et al. 2006). Increased IL11Rα staining was also

found in human endometrial carcinoma samples, suggesting that aberrant signaling may play a

role in progression of the disease (Yap et al. 2010). It is currently unknown if IL11Rα is involved

in the induction of pyometra though or whether exposures to estrogens or estrogenic compounds

such as BPA directly or indirectly interact with the receptor. Located within the promoter are

three EREs to which an activated ERα dimer could bind, leading to potential activation or

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repression of the receptor or its downstream targets (SABiosciences 2012). To determine how

estrogenic EDCs, including BPA and EE, affect Il11ra expression, targeted ChIP assays could be

used. Also, sequencing of the Il11ra promoter can identify single nucleotide polymorphisms

(SNPs) that can impact differences in gene expression between the CD1 and C57Bl/6N strains

and aids in explaining the sensitivity between strains to estrogenic compounds. The sensitivity

between strains can be correlated to differences in sensitivities between humans to estrogenic

EDCs. Identification of SNPs can also aid in defining the etiology of pyometra and lead to the

development of a treatment for the disease.

Research for this dissertation determined that BPA exposure altered collagen

accumulation by increasing collagen synthesis and decreasing collagen degradation, leading to

an overall fibrotic phenotype. However, the mechanism of action of BPA that led to the

alterations in collagen synthesis and degradation remains unknown. Under physiological

conditions, E2 activated ERα and led to transcriptional upregulation of collagens and MMPs

(Gaide Chevronnay et al. 2012, Frankel et al. 1988). Exposure to BPA also increased

transcription of collagens, suggesting that BPA was also activating ERα to upregulate collagen

synthesis. However, the effects of BPA and E2 on MMPs and proteins involved in collagen

degradation were opposite, suggesting a different mechanism of action of BPA compared to E2.

One possibility is that BPA may be acting as a partial or inverse agonist on ERα or be

antagonizing the effects of E2. A partial agonist is a ligand that binds to a receptor at the active

site but produces a submaximal response (Golan et al. 2012). An inverse agonist is a ligand that

binds to the same receptor target as an agonist but the induced pharmacological response is in the

opposite direction (Golan et al. 2012). It is possible that BPA was still binding to ERα but had a

decreased efficacy compared to E2, leading to decreased expression and activity of MMPs.

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Partial agonist-like and antagonist activity of BPA at ERα in the uterus has been reported. For

example, BPA increased PR expression in the immature rat uterus, but also antagonized E2-

induced stimulation in PR expression (Gould et al. 1998).

A second possibility is that BPA was binding to another receptor target altogether. At nM

concentrations, BPA bound to and activated ERα and ERβ (Li et al. 2013, Wetherill et al. 2007,

Matthews et al. 2001, Kuiper et al. 1998). However, BPA also had strong binding to ERRγ and

GPR30 at similar concentrations (Okada et al. 2008, Thomas and Dong 2006). Other receptor

targets for BPA included AR and THR, at which BPA bound and inhibited at µM concentrations

(Teng et al. 2013, Moriyama et al. 2002, Paris et al. 2002, Sohoni and Sumpter 1998). Binding of

BPA to any of these receptors could potentially elicit the observed alterations in collagen

degradation and decreased expression and activity of MMPs. To determine this, a luciferase

assay containing the MMP2 promoter could be used. Treatment with BPA or E2 would help

answer which receptor BPA acts through to alter collagen degradation. ChIP experiments can

also be performed to determine if ERα or other receptor targets of BPA are bound to the

promoters of Mmp genes. These experiments would help to narrow down through which receptor

BPA is acting to induce the observed alterations in collagen degradation. However, translocation

and signaling of nuclear receptors also relies on the recruitment of coactivators or corepressors.

Experiments designed to identify the specific coregulators involved in Mmp transcription would

greatly expand knowledge on the differential activation of these genes by BPA.

The mechanisms behind gland nest formation and progression of gland nest severity is an

aspect of equine endometrosis that needs to be further explored. Because equine endometrosis is

irreversible, understanding the etiology and progression could help to develop treatments to

either prevent or halt progression. The mare would then have a better chance of carrying a fetus

165  

to term. To the best of knowledge, it is unknown if gland nest formation occurs in humans too.

However, gland nest formation and associated periglandular fibrosis could also play a role in

infertility in humans. Again, understanding the etiology of periglandular fibrosis may lead to

treatments for fibrosis in the human uterus and increased fertility. Because the C57Bl/6N strain

already exhibits increased incidence of gland nests and stromal and periglandular fibrosis, this

strain could potentially be a useful small animal experimental model to study the etiology and

progression of equine endometrosis.

A third future direction for this dissertation research would be to determine the effects of

alterations in collagen accumulation on uterine physiology. In particular, it would be important

to examine fertility, including decidualization, implantation, uterine stretch, and parturition.

While collagen fibrils are involved in decidualization and MMPs degrade these fibrils during

implantation (Carbone et al. 2006, Staun-Ram and Shalev 2005, Dziadek et al. 1995,

Abrahamsohn and Zorn 1993), the increased expression of collagens and the decreased

expression and activity of MMPs observed in this research could significantly decrease fertility

due to the inability of the uterus to undergo decidualization and implantation. Alterations in

collagen accumulation could have a significant detrimental impact on fertility and may help to

explain the increases in infertility in women with fibrotic uterine diseases.

When comparing CD1 mice in the whole life and adult exposure models, there was a

clear lack of effect on the development of fibrotic uterine diseases in the whole life exposure

model, as well as only a slight increase in collagen accumulation. However, the 30 ppm BPA-

exposed CD1 group in both models was shown to have increases in immune responsiveness

(Kendziorski and Belcher 2015). These findings suggest that physiological changes in the uterine

166  

environment during mating, pregnancy and parturition are important for the development of

fibrotic uterine pathologies and should be further investigated.

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LIST OF PUBLICATIONS AND ABSTRACTS

Publications

JA Kendziorski and SM Belcher. 2015. Effects of whole life exposure to bisphenol A or 17α- ethinyl estradiol in uterus of nulligravida CD1 mice. Data in Brief.

JA Kendziorski and SM Belcher. 2015. Strain-specific induction of endometrial periglandular fibrosis in mice exposed during adulthood to the endocrine disrupting chemical bisphenol A. Reprod Toxicol, 58: 119-130. doi: 10.1016/j.reprotox.2015.08.001.  

EL Kendig, DR Buesing, SM Christie, CJ Cookman, RB Gear, ER Hugo, JA Kendziorski, KR Ungi, K Williams, SN Kasper, and SM Belcher. 2012. Estrogen-like disruptive effects of dietary exposure to bisphenol A or 17α-ethinyl estradiol in CD1 mice. Int J Toxicol, 31(6): 537-50. doi: 10.1177/1091581812463254

JA Kendziorski, EL Kendig, RB Gear, and SM Belcher. 2012. Strain specific induction of pyometra and differences in immune responsiveness in mice exposed to 17α-ethinyl estradiol or the endocrine disrupting chemical bisphenol A. Reprod Toxicol, 34(1): 22-30, doi: 10.1016/j.reprotox.2012.03.001

Abstracts

J Kendziorski and S Belcher. Strain-Specific Induction of Endometrial Periglandular Fibrosis in Mice Exposed During Adulthood to the Endocrine Disrupting Chemical Bisphenol A. Oral Presentation. 49th Annual Albert J. Ryan Fellowship Symposium. North Conway, NH. May 15, 2015.

J Kendziorski and S Belcher. Exposure to Bisphenol A Increases Gland Nest Formation and Fibrosis in the Murine Uterus. College of Medicine Graduate Student Poster Forum. University of Cincinnati, Cincinnati, OH. March 6, 2015.

J Kendziorski and S Belcher. Exposure to Bisphenol A Increases Gland Nest Formation and Fibrosis in the Murine Uterus. Graduate Student Expo and Poster Forum. University of Cincinnati, Cincinnati, OH. March 6, 2015.

J Kendziorski and S Belcher. Sensitivity to estrogenic actions of bisphenol A in the uterus of two mouse strains. Oral presentation. 48th Annual Albert J. Ryan Fellowship Symposium. North Conway, NH. May 17, 2014.

J Kendziorski and S Belcher. Bisphenol A blocks the adverse growth effects of in utero arsenic exposure. Poster. University of Cincinnati Graduate Student Poster Forum. Cincinnati, OH. February 28, 2014.

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J Kendziorski, E Kendig, R Gear, and S Belcher. Dietary bisphenol A increases the uterine immune response and induces pyometra in mice. Poster. NIEHS BPA Grantee Research Meeting. NIEHS Research Triangle Park, NC. January 28-29, 2013.

J Kendziorski, E Kendig, R Gear, and S Belcher. Dietary bisphenol A increases the uterine immune response and pyometra in mice. Oral presentation. Society of Toxicology 2012 Annual Meeting. San Francisco, CA. March 11-15, 2012.

E Kendig, C Lo, C Cookman, S Christie, D Buesing, R Gear, J Kendziorski, K Ungi, and S Belcher. The effects of oral Bisphenol A exposure in the developing CD-1 mouse. Poster. ENDO 2011: The Endocrine Society’s 93rd Annual Meeting and Expo. Boston, MA. June 4- 7, 2011.

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