By Hala Kufaishi 2015

Vaginal Fibroblasts Derived From Premenopausal Women With and Without Severe Pelvic Organ Prolapse: Differential Characteristics and Effect of Mechanical Stretch

Hala Kufaishi

A thesis submitted in conformity with the requirements for the degree of Master of Science Institute of Medical Science University of Toronto

Vaginal Fibroblasts Derived From Premenopausal Women with and without Severe

Pelvic Organ Prolapse: Differential Characteristics and Effect of Mechanical Stretch

Master of Science, 2015 Hala Farid Kufaishi Institute of Medical Science, University of Toronto

ABSTRACT

Mechanical properties of connective tissue depend on appropriate cell-cell and cell- matrix interactions. Failure of the vaginal connective tissue integrity may cause the development of pelvic organ prolapse (POP). I hypothesized that (1) primary fibroblasts derived from vaginal tissue (VFs) of premenopausal women with severe POP display differential functional characteristics as compared to VFs derived from age-matched non-

POP women; (2) continuous mechanical loading of the POP VFs can directly influence the extracellular matrix (ECM) proteins and matrix remodeling factors they secrete. I demonstrated that POP-VFs show altered in vitro cellular characteristics vs. non-POP-VFs.

Moreover, mechano-responses of POP-VFs to static mechanical loading on collagen-coated plates showed a difference in the expression of ECM and cell adhesion proteins. These data indicate that risk factors that induce stretch of pelvic floor may result in defective vaginal tissue composition and subsequent POP development; which should be considered during mesh-augmented reconstructive surgery of the pelvic floor.

ACKNOWLEDGMENTS

The three years I have spent at the Lye Lab have been a truly memorable and rewarding experience. I have learned countless life lessons, met so many inspirational people, and discovered an inner confidence in myself that I did not know I had.

First and foremost, I would like to thank God, whose many blessings have made me who and where I am today. The following document summarizes three years’ worth of effort, frustration and achievement. However, there are several people with whom I am indebted for their contribution in the research, study and writing of this thesis. Thank you to my parents and in-laws for their continuous support and encouragement, and for babysitting the countless hours that I spent writing this dissertation. To my wonderful husband, Mohamad, thank you for supporting me through my many times of stress, excitement, frustration, and celebration. I am so grateful to my son, Yezen, for showing me that with perseverance and good time-management, raising an infant, writing exams, finishing up a thesis and getting accepted to a dream residency program is, in fact, possible.

It is with immense gratitude that I acknowledge the support and help of my Professor, Dr.

Stephen Lye. Thank you for the opportunity to undertake my thesis project at your lab as the honorary Urogynecology research student. I would also like to thank Dr. Oksana Shynlova for her guidance, support and mentorship. I share the credit of my work with Dr. May

Alarab. Without her guidance, persistent help and moral support, this thesis would not have been possible.

iii

I consider it an honor to work with each member of the Lye and Kingdom Lab. Thank you

Anna Dorogin and Ela Matysiak-Zablocki for being wonderful office-space mates, and for your continuous help with troubleshooting my experiments when all else fails. Thank you

Dr. Lubna Nadeem, Dr. Caroline Dunk, Dr. Jianhong Zhang, Dora Baczyk and Dr. Mark

Kibschull for your advice and support. I am indebted to my many colleagues who supported me: Melissa Kwan, Yaryna Rybak, Richard Maganga, Tina Nguygen, Tam Lye, Farshad

Ghasemi, Christina Lee, Khrystyna Levytsky, Dr. Jan Heng, Dr. Kristin Connor and Dr.

Sascha Drewlo. Thank you Bev Bessey, I am so grateful for all your work and support in organizing my committee meetings, following up on recommendation letters and pulling application documents together. It gives me great pleasure in acknowledging the support and help of my supervisory committee members: Dr. Theodore Brown and Dr. Boris Hinz. Thank you for your continuous support, insightful discussions and help in directing my project and making it possible. Lastly, thank you Dr. Harold Drutz for your support and for securing funding for my position as a research assistant and Master student.

TABLE OF CONTENTS

ABSTRACT ...... ii ACKNOWLEDGMENTS ...... iii TABLE OF CONTENTS ...... v LIST OF TABLES ...... vii CHAPTER 1 ...... vii CHAPTER 2 ...... vii CHAPTER 3 ...... vii APPENDIX ...... vii LIST OF FIGURES ...... viii CHAPTER 1 ...... viii CHAPTER 2 ...... viii CHAPTER 3 ...... viii LIST OF ABBREVIATIONS ...... ix CHAPTER 1: LITERATURE REVIEW ...... 1 1.1 Overview: The Problem of Pelvic Floor Disorders ...... 2 1.1.1 Clinical Presentation of Pelvic Organ Prolapse ...... 2 1.1.2 Health, Social and Economic Aspects of Pelvic Organ Prolapse ...... 2 1.1.3 Risk Factors for Pelvic Organ Prolapse ...... 3 1.1.4 Anatomy and Histology of Normal Pelvic Floor ...... 5 1.2 The Extracellular Matrix of the Pelvic Floor ...... 6 1.2.1 Structural Components of ECM ...... 7 1.2.2 ECM turnover and breakdown ...... 17 1.2.3 Ground Substance ...... 21 1.2.4 Cell-Cell and Cell-Matrix Adhesion Molecules...... 24 1.3 Pelvic ECM and POP development...... 24 1.4 The Effects of Stretch on Pelvic Floor Tissue ...... 29 1.4.1 Vaginal Human Tissue and Mechanical Stretch ...... 29 1.4.2 Animal Models of Vaginal Stretch ...... 30 1.4.3 Cellular Response to Mechanical Stretch and POP ...... 31 1.4.4 Cell-Based Tissue Engineering and POP ...... 33 1.5 Rationale and Hypothesis ...... 34 1.5.1 Rationale ...... 34 1.5.2 Hypothesis and Objectives ...... 36 CHAPTER 2: MATERIALS and METHODS ...... 38 2.1 Patient Selection ...... 39 2.2 Tissue Collection ...... 39 2.3 Derivation and Maintenance of Primary Human Vaginal Fibroblasts ...... 39 2.4 Comparing the Biological Characteristics of Human Vaginal Fibroblasts ...... 42 2.4.1 Fibroblast cell verification by indirect immunofluorescence ...... 42 2.4.2 Cell Attachment on Different Extracellular Matrices ...... 42 v

2.4.3 Cell Proliferation on Different Extracellular Matrices ...... 44 2.5 Application of Static Mechanical Stretch ...... 45 2.6 Viability Studies ...... 45 2.6.1 Fluorescein Diacetate- Propidium Iodide Assay ...... 45 2.6.2 Lactate Dehydrogenase Cytotoxicity Assay ...... 46 2.7 Gene Expression Analysis ...... 46 2.7.1 ECM and Adhesion Molecules Quantitative Profiler PCR arrays ...... 47 2.7.2 Real Time Reverse Transcription Polymerase Chain Reaction (qRT-PCR) ...... 48 2.8 Quantitative Detection of Protein Expression in Conditioned Medium (CM) ...... 51 2.8.1 Quantibody Protein Array ...... 51 2.8.2 Western Immunoblot Analysis ...... 53 2.8.3 Zymography ...... 54 2.9 Statistical Analysis ...... 55 CHAPTER 3: RESULTS ...... 57 3.1 Patient Demographics ...... 58 3.2 Comparing the Biological Characteristics of the Vaginal Fibroblasts ...... 60 3.2.1 Fibroblast Cell Identification by Indirect Immunofluorescence ...... 60 3.2.2 Cell Attachment on Different Extracellular Matrices ...... 63 3.2.3 Cell Proliferation on Different Extracellular Matrices ...... 65 3.3 Viability Studies: Mechanical Stretch Does Not Induce Cell Injury ...... 67 3.4 Gene Expression Profile ...... 69 3.4.1 ECM and Adhesion Molecule Quantitative PCR Arrays ...... 69 3.4.2 Real Time Reverse Transcription Polymerase Chain Reaction (qRT PCR) Analysis ...... 73 3.5 Quantitative Detection of Protein Expression in Conditioned Media ...... 76 3.5.1 Quanti-body Protein Array ...... 76 3.5.2 Western Immunoblot Analysis ...... 78 3.6 Zymography ...... 85 CHAPTER 4: DISCUSSION ...... 87 4.1 Overall Summary ...... 88 4.2 Biological characteristics of VFs and their Ability to Produce ECM Proteins ...... 90 4.3 Mechano-responses of Primary Human Fibroblasts Derived from ...... 97 Non-Prolapsed and Prolapsed Vaginal Tissue ...... 97 CHAPTER 5: FUTURE DIRECTIONS ...... 103 REFERENCES ...... 113

LIST OF TABLES

CHAPTER 1

TABLE 1.1: THE COLLAGEN FAMILY OF PROTEINS...... 10 TABLE 1. 2: CLASSIFICATION OF MATRIX METALLOPROTEINASE ENZYMES...... 19 CHAPTER 2

TABLE 2. 1: REAL-TIME PCR PRIMER SEQUENCES OF A PANEL OF GENES STUDIED AND ...... 50 TABLE 2.2: SUMMARY OF ANTIBODIES USED IN IMMUNOBLOT ANALYSIS ...... 55 CHAPTER 3

TABLE 3.1: SUMMARY OF PATIENTS DEMOGRAPHICS ...... 59

APPENDIX

APPENDIX A: CONSENT FORM ...... 106 APPENDIX B: DATA TO BE COLLECTED FROM EACH PATIENT INVOLVED IN THE STUDY ...... 109 APPENDIX C : VAGINAL WALL BIOPSY SITE...... 110 APPENDIX D : LIST OF 84 ECM AND CELL ADHESION GENES PER FUNCTIONAL GROUP...... 111

vii

LIST OF FIGURES

CHAPTER 1

FIGURE 1.1: COLLAGEN AND ELASTIN METABOLISM...... 13

CHAPTER 2

FIGURE 2. 1 DERIVATION OF PRIMARY VAGINAL FIBROBLASTS...... 41 FIGURE 2.2 QUANTIBODY ARRAY-BASED MULTIPLEX SANDWICH ELISA SYSTEM...... 52 CHAPTER 3

FIGURE 3.1: IMMUNOFLUORESCENCE OF PRIMARY VAGINAL FIBROBLASTS (VFS) ...... 62 FIGURE 3.2: ATTACHMENT OF VAGINAL FIBROBLASTS ...... 64 FIGURE 3.3: PROLIFERATION OF VAGINAL FIBROBLASTS...... 66 FIGURE 3. 4: VIABILITY OF VAGINAL FIBROBLASTS ...... 68 FIGURE 3.5: GENE EXPRESSION HEAT MAP...... 70 FIGURE 3.6: RELATIVE EXPRESSION OF ADAMTS, MMPS AND TIMPS TRANSCRIPTS FROM POOLED RNA SAMPLES ...... 71 FIGURE 3.7: THE EXPRESSION LEVEL OF SELECTED GENES DETERMINED BY QRT-PCR ...... 74 FIGURE 3.8: QUANTI-BODY PROTEIN ARRAY ANALYSIS ...... 76 FIGURE 3.9: WESTERN IMMUNOBLOT ANALYSIS OF LOXL3 AND LOXL4 IN VAGINAL TISSUE AND CONDITONED MEDIUM ...... 79 FIGURE 3.10: COMMASSIE BLUE STAINING...... 80 FIGURE 3.11: WESTERN IMMUNOBLOT ANALYSIS OF LOX, LOXL1-2 IN CONDITIONED MEDIUM ...... 82 FIGURE 3.12: WESTERN IMMUNOBLOT ANALYSIS OF ADAMTS2 IN CONDITIONED MEDIUM ...... 83 FIGURE 3.13: WESTERN IMMUNOBLOT ANALYSIS OF BMP-1 IN CONDITIONED MEDIUM ...... 84 FIGURE 3.14: REPRESENTATIVE GELATIN ZYMOGRAPHY...... 86

CHAPTER 4

FIGURE 4.1:ECM SYNTHESIS AND DEGRADATION IN NON-POP AND POP VF’S UNDER NON-STRETCH AND STRETCH CONDITIONS……………………………………………………………………………………………………………………….101

viii

LIST OF ABBREVIATIONS

ADAM A Disintegrin and Metalloprotease ADAMTS A Disintegrin and Metalloproteinase with Thrombospondin Motifs ATT α-1-anti-trypsin BMI Body Mass Index BMP Bone Morphogenic Protein BSA Bovine Serum Albumin cDNA Complementary DNA CM Conditioned Media COL Collagen COPD Chronic Obstructive Pulmonary Disease CUB C-terminal complement –uegf-BMP1 EBP Elastin Binding Protein ECM Extracellular Matrix EDS Ehler Danlos Syndrome EGF Epidermal Growth Factor ELISA Enzyme-Linked Immunosorbent Assay FACIT Fibril Associated Collagens with Interrupted Triple Helices FBS Fetal Bovine Serum FDA-PI Fluorescein DiAcetate-Propidium Iodide GAG Glycosyaminoglycans HBSS Hank’s buffered salt solution without Ca2+ and Mg2+ VF Vaginal Fibroblast ICTP collagen Type I carboxyterminal telopeptide IPSC induced pluripotent stem cells ITS-A Insulin-Transferrin-Selenium-Sodium Pyruvate Solution K/O Knockout LDH Lactate Dehydrogenase LOX Lysyl Oxidase LOXL LOX-like LTRI Lunenfeld-Tanenbaum Research Institute MMP Matrix Metalloproteinase MT Membrane Type mTLD Mammalian Tolloid mTLL1 TLD-like 1 mTLL2 TLD-like 2 MTT Thiazolyl Blue Tetrazolium Bromide NCBI National Centre for Biotechnology Information NE neutrophil Elastase NS Non-stretched NVD Normal Vaginal Delivery

OD Optical Density PCP Pro-collagen-C-Proteinase PFD Pelvic Floor Disorders PG Proteoglycans PICP Procollagen Type 1 Carboxyterminal Propeptide PIIINP Procollagen type III Aminoterminal Propeptide PNP Pro-collagen-N-Proteinase POP Pelvic Organ Prolapse PVDF Polyvinylidene Difluoride Membrane QAH-MMP1 Quantibody Human MMP Array 1 qRT-PCR Real Time Reverse Transcription Polymerase Chain Reaction RT Room Temperature RT Reverse Transcription S Stretched SEM Standard Error of Mean SF-DMEM Serum Free DMEM SLRP Small Leucine-Rich Proteoglycans SMC Smooth Muscle Cell SUI Stress Urinary Incontinence TGF- β Transforming Growth Factor Beta TBST-T Tris-buffered saline (TBS), with 0.05% tween TIMP Tissue Inhibitors of Metalloproteinase TLD Tolloid TTP Thrombotic Thrombocytopenic Purpura VF Vaginal Fibroblast vWF Von Willebrand Factor vWFCP Von Willebrand factor-Cleaving Protease WT Wild Type ADAM A Disintegrin and Metalloprotease ADAMTS A Disintegrin and Metalloproteinase with Thrombospondin Motifs ATT α-1-anti-trypsin BMI Body Mass Index BMP Bone Morphogenic Protein BSA Bovine Serum Albumin cDNA Complementary DNA CM Conditioned Media COL Collagen COPD Chronic Obstructive Pulmonary Disease CUB C-terminal complement –uegf-BMP1 EBP Elastin Binding Protein ECM Extracellular Matrix EDS Ehler Danlos Syndrome EGF Epidermal Growth Factor ELISA Enzyme-Linked Immunosorbent Assay x

FACIT Fibril Associated Collagens with Interrupted Triple Helices FBS Fetal Bovine Serum FDA-PI Fluorescein DiAcetate-Propidium Iodide GAG Glycosyaminoglycans HBSS Hank’s buffered salt solution without Ca2+ and Mg2+ ICTP collagen Type I carboxyterminal telopeptide IPSC induced pluripotent stem cells ITS-A Insulin-Transferrin-Selenium-Sodium Pyruvate Solution K/O Knockout LDH Lactate Dehydrogenase LOX Lysyl Oxidase LOXL LOX-like LTRI Lunenfeld-Tanenbaum Research Institute MMP Matrix Metalloproteinase MT Membrane Type mTLD Mammalian Tolloid mTLL1 TLD-like 1 mTLL2 TLD-like 2 MTT Thiazolyl Blue Tetrazolium Bromide NCBI National Centre for Biotechnology Information NE neutrophil Elastase NS Non-stretched NVD Normal Vaginal Delivery OD Optical Density PCP Pro-collagen-C-Proteinase PFD Pelvic Floor Disorders PG Proteoglycans PICP Procollagen Type 1 Carboxyterminal Propeptide PIIINP Procollagen type III Aminoterminal Propeptide PNP Pro-collagen-N-Proteinase POP Pelvic Organ Prolapse PVDF Polyvinylidene Difluoride Membrane QAH-MMP1 Quantibody Human MMP Array 1 qRT-PCR Real Time Reverse Transcription Polymerase Chain Reaction RT Room Temperature RT Reverse Transcription S Stretched SEM Standard Error of Mean SF-DMEM Serum Free DMEM SLRP Small Leucine-Rich Proteoglycans SMC Smooth Muscle Cell SUI Stress Urinary Incontinence TGF- β Transforming Growth Factor Beta TBST-T Tris-buffered saline (TBS), with 0.05% tween

TIMP Tissue Inhibitors of Metalloproteinase TLD Tolloid TTP Thrombotic Thrombocytopenic Purpura VF Vaginal Fibroblast vWF Von Willebrand Factor vWFCP Von Willebrand factor-Cleaving Protease WT Wild Type

xii

CHAPTER 1: LITERATURE REVIEW

1.1 Overview: The Problem of Pelvic Floor Disorders

Pelvic floor disorders (PFDs) have a significant impact on the quality of life of women, and encompass many syndromes such as stress urinary incontinence (SUI), anal incontinence, chronic pain and pelvic organ prolapse (POP). These share a common pathophysiological process based on pelvic floor loss of support due to tissue and muscle laxity. A recent cross- sectional analysis in the United States concluded that 24% of women above 20 years of age suffer from at least one PFD; of these women, 16% are affected by SUI and 3% have been diagnosed with POP. With increasing age and/or parity, the proportion of women suffering from more than one PFD increases [1].

1.1.1 Clinical Presentation of Pelvic Organ Prolapse

Pelvic organ prolapse (POP) is characterized by the descent of the uterus, bladder or rectum into the vaginal canal. Patients can present with varying degrees of prolapse, the most severe with the pelvic organs protruding completely through the genital hiatus. POP covers a wide spectrum of clinical conditions, and it is closely related to other pelvic floor disorders including urinary incontinence and fecal incontinence. Symptoms include a sensation of vaginal fullness or pressure, sacral or lower back pain, vaginal spotting due to ulceration of the cervix or vagina, abdominal pain and sexual, voiding and defecatory dysfunction [2].

1.1.2 Health, Social and Economic Aspects of Pelvic Organ Prolapse

Alongside the physical symptoms that accompany these disorders, there is also substantial emotional impact on the women affected, which commonly results in social isolation, psychological distress, anxiety and depressive symptoms [2-4]. Hence, this condition significantly reduces the quality of life of those women. POP affects 1 of 3 premenopausal

2 women and nearly half of postmenopausal women, with a lifetime prevalence of 50% [5, 6].

The cumulative incidence for undergoing surgery for POP is 11%, with a re-operation rate of nearly 29% [5] . In other words, approximately 500,000 women undergo surgery for POP and pelvic floor dysfunction each year in the United States [7], which amounts to a collective cost of over $1 billion dollars [8] .The need for POP surgery increases with age, and it is estimated that demand for POP surgeries will increase by 46% over the next four decades [5]. These statistics demonstrate that POP is a major and growing burden on the health care system, especially since demographics show that women older than 80 years of age are the fastest growing population segment in developed countries [8].

1.1.3 Risk Factors for Pelvic Organ Prolapse

Risk factors for the development of POP can be categorized as inciting, predisposing, decompensating and promoting [9]. Epidemiological data supports the notion that vaginal delivery is the greatest independent inciting risk factor for the development of POP and other pelvic floor disorders [10-12]. This is due to the fact that the majority of women that undergo vaginal delivery have some anatomical evidence of damage or disruption to their pelvic floor

[13-15]. In fact, the relative risk for developing prolapse in women who have undergone one normal vaginal delivery (NVD) is 8.4, and rises to 10.4 after four or more NVDs [16] . There are three mechanisms whereby labor and vaginal delivery affect the pelvic floor. Firstly, increased mechanical distension leads to direct tearing of the connective tissue and the fibro- muscular components of the pelvic floor. In addition, vascular compression due to the increased pressures results in a hypoxic environment to the surrounding tissues and structures. Thirdly, the neurological bundles are compromised by the combined effect of the direct shearing forces in the hypoxic environment, leading to both motor and sensory nerve loss [17]. However, the majority of parous women do not progress to symptomatic prolapse, and those that do often

3 develop POP years to decades following transient or long-term labor-related injury [14].

Furthermore, POP has been observed in nulliparous women [18]. This has led to the conclusion that there are additional factors that contribute to the progression of POP by affecting the vagina and the surrounding connective tissue.

Factors that predispose to the development of POP include race, family history and genetics, which present as abnormalities in their connective tissue histology and morphology.

Women with connective tissue disorders, such as Ehler Danlos syndrome (EDS) and Marfan’s syndrome, have hyper-extensible skin and increased joint mobility. Post vaginal-delivery, the pelvic floor of these women does not undergo normal repair mechanisms, and thus there is an increased incidence of POP [5, 19]. It is now established that these women have intrinsic histological changes in the expression of collagen, elastin and their modulators, however, it is still not known whether those changes predispose to prolapse or are results of prolapse. Genetic predisposition to POP can be better understood through family studies. There is a high concordance of prolapse between nulliparous and parous sisters, and a 2 to 3 fold increase in the relative risk of developing POP among first-degree relatives (mother or sister).

Furthermore, at least 30% of women undergoing POP surgery under 45 years of age reported one first degree relative with POP [20]. Racial differences have also been reported in the prevalence of POP. Hispanic and Caucasian women have a five times higher risk of developing symptomatic prolapse in comparison to African-American women. Furthermore, Caucasian women have a 1.4-fold increase in the relative risk of developing severe POP (stage 3 and 4)

[21].

Virtually all studies on pelvic floor disorders agree that there is an increased incidence of POP with advanced age. Increasing age contributes to the development of POP through the combination of physiological aging, hypo-estrogenism precipitated by menopause, and age-

4 related organic and degenerative disease, all factors that decompensate the pelvic floor structures [22].

It has been identified that some repetitive activities that weaken the pelvic floor muscles may promote the development of POP. For instance, women working in factories or as laborers, exposed repeatedly to heavy lifting, are more likely to develop POP [22]. Chronic obstructive pulmonary disease (COPD) and asthma also predispose women to POP due to the increased intra-abdominal pressure that is transmitted to the pelvic connective tissue [9]. Similarly, chronic constipation promotes the development of POP due to the pelvic pressure induced by chronic straining [9]. Furthermore, a body mass index (BMI) greater than 30 has been associated with a 40-75% increased risk of POP [23]. Previous gynecological surgery is also a significant risk factor that promotes the development of POP, with an eight-fold risk of developing recurrent prolapse in patients in comparison to non-POP patients following hysterectomy for genital prolapse [24] .

1.1.4 Anatomy and Histology of Normal Pelvic Floor

The pelvic floor primarily supports pelvic organs, including the urethra, vagina, uterus, bladder and rectum. It comprises a highly interconnected system of striated muscle, smooth muscle and connective tissue [25, 26]. The striated muscle component is composed of three muscles (pubococcygeous, ileococcygeous and coccygeous muscles) referred together as the levator ani muscle complex. Tonic contraction of the levator ani supports and maintains the pelvic organs in place. The connective tissue support to the vagina is composed of a ligamentous component, the uterosacral, cardinal ligaments and lateral attachments to the arcus tendinous fasciae pelvis, as a well as the perineal body and membrane that support the distal vagina [27, 28]. Due to the load sharing relationship between the two, current literature agrees

5 that both the levator ani and the connective tissue complex are necessary for normal vaginal support [29].

The walls of the vagina are comprised of 4 layers: the epithelium, lamina propria, muscularis and adventitia. The outermost (luminal) epithelium is composed primarily of squamous cells, and undergoes constant turnover under hormonal control. The lamina propria consists primarily of fibroblasts and dense collagen fibers, produced by fibroblasts, which provide mechanical and structural integrity to the vagina. Below the lamina propria lies the muscularis, a fibro-muscular layer of smooth muscle cells (SMCs) that provides longitudinal and central support to the vagina [25]. The innermost adventitia is composed of a layer of loose areolar connective tissue, and is connected to the bladder anteriorly and to the rectum posteriorly [30].

1.2 The Extracellular Matrix of the Pelvic Floor

The connective tissue that supports the pelvic floor is mainly composed of the fibrous elements of extracellular matrix (ECM) - collagen and elastin, as well as ground substance, and cellular components [31]. Recently, there has been an increase in studying how tissue composition and content differs between patients presenting with severe POP and non-POP age-matched patients. Researchers have mainly focused on sampling the uterosacral ligaments

[32, 33], the pubocervical fascia [34], and the vaginal wall [35-40]. While conclusions are different between studies, the overall consensus is that connective tissue of the vaginal wall is representative of the pelvic floor tissue and therefore could be used to study POP in women

[41]. The vaginal connective tissue undergoes constant remodeling during a women’s premenopausal and postmenopausal lifetime, a process that is well balanced and closely regulated via several factors. It is important to understand how this remodeling can affect tissue integrity and strength [42]. It was suggested that an imbalance in the process of tissue 6 remodeling could result in weakening of this connective tissue, predisposing to pelvic floor dysfunction [35, 38, 39].

The cellular components of the connective tissue of the vagina consist mainly of fibroblasts and SMCs; a small number of fat cells and mast cells may also be present.

Fibroblasts and SMCs are mechano-sensitive cells, and are the primary modulators of ECM remodeling in the vaginal connective tissue [43]. By responding to hormonal, biochemical and physical stimuli, they act to produce compounds involved in ECM homeostasis, including collagens (collagen type I and III), elastin, matrix metalloproteinases (MMPs), tissue inhibitors of metalloproteinases (TIMPs) [32], bone morphogenic protein (BMP) [35] and a disintegrin and metalloproteinase with thrombospondin motifs (ADAMTS) family members [44].

Fibroblasts also produce ground substance, composed mainly of glycoproteins, proteoglycans

(PGs) and substrate adhesion molecules [29].

1.2.1 Structural Components of ECM 1.2.1.1 Collagens

Fibrillar Collagens. Collagen is the major insoluble fibrillar protein in connective tissue.

There are 29 members of the collagen family known so far, but 80 – 90 % of the collagen in the body consists of the fibril forming types I, II, III, V and XI. Collagen type I is ubiquitous, with large amounts present in the skin, fascia, organ capsules, fibro-cartilage and tendons. The content of collagen I within a connective tissue determines the tensile strength of the tissue[45,

46]. Mutations in the gene encoding collagen I cause osteogensis imperfecta and some forms of EDS [47]. Another form of EDS is due to mutations in collagen type III. Collagen III is present in large amounts in loose connective tissue of organs subjected to repetitive mechanical stretching, such as the skin, uterus, aorta, lungs and ligaments, and contributes to tissue elasticity and extensibility. It is also found to be the initial collagen type that is deposited at the

7 site of wound healing, and is replaced by the stronger type I collagen over several months.

Collagen I usually associates with collagen III and V to form collagen fibrils, while collagen XI co-distributes with collagen II. Collagen I is reported to be five times more abundant than collagen III in pelvic organ connective tissue. Furthermore, studies have shown that an increase in the ratio of collagen III or V to collagen I is associated with a decrease in the mechanical strength and integrity of the connective tissue [30, 48].

Microfibrillar Collagens. Collagens VI and XXVIII are the microfibrillar collagens, they appear on the structural level as fine filaments or microfibrils with faint cross-banding [49].

Type VI collagen fibrils first assemble inside the cell as overlapping dimers, and then align to form beaded tetramers. Once secreted into the ECM, the tetramers aggregate into filaments to form a microfibrillar network present in all connective tissue, except bone, that provide structural links to cells [50].

Fibril Associated Collagens with Interrupted Triple Helices (FACIT). The collagen types

IX, XII, XIV, XVI, XIX, and XX are known as the FACIT collagens. Their structure consists of short non-helical domains that interrupt their collagenous helical structures, and are associated with the surface of various fibrils. Collagen IX associates with collagen II in cartilage and vitreous body. Type XII and XIV collagens co-distribute with collagen I in skin, tendons, lung, liver, placenta, blood vessels, and have similar structures to collagen IX.

Collagen XIX is rare, found in muscle tissue, and is localized to basement membrane zones, while collagen XX is more widely distributed and mainly found in the corneal epithelium.

Although their structures and tissue distributions have been characterized, very little is known about the function of these collagens [46, 51].

Basement Membrane Collagen. Collagen IV is the most important structural protein in basement membranes. It forms a sheet like stable structure necessary for it to integrate

8 proteoglycans and other components of the basement membrane. Mutations in collagen IV genes have been implicated in Goodpasture’s disease and Alport’s disease, two conditions that require functional basement membrane interactions for correct renal/alveolar and sensori-neural function, respectively [52].

Transmembrane Collagens. The transmembrane collagens include collagens XIII and XXV.

These collagens are involved in cell adhesion, and have been implicated in malignancies, neural functions, eye development, and growth modulation. Of note, the extracellular domain is cleaved by ADAM (a disintegrin and metalloprotease) family proteinases [46].

Table 1.1: The Collagen Family of Proteins [46].

Structural Group Collagen Type

Fibril-forming I, II, III, V, XI

Microfibrillar VI, XXVIII

Anchoring Fibrils VII

Hexagonal network-forming VIII, X

FACIT IX, XII, XIV, XIX, XX, XXI

Basement Membrane IV

Transmembrane XIII, XVII, XXIII, XXV

Multiplexins XV, XVI, XVIII

Ungrouped XXII, XXIV, XXVI, XXVII, XXIX

1.2.1.2 Elastin

Elastin is produced by multiple cell types, including fibroblasts, chondroblasts, endothelial and mesothelial cells [53]. It is a major protein responsible for the stretch and recoil properties of connective tissue. Compared to collagen, which can only elongate to 4% of its length before rupturing, elastin elongates up to 70% and can return to its original shape [54].

The production of elastin is mainly confined to the late fetal and early neonatal period. Unlike collagen, elastin undergoes very little turnover during adult life, except in the female uterus, where elastic fibers degrade during labor and are re-synthesized after vaginal delivery [55].

This process is under the control of reproductive hormones, and decreases with age.

1.2.1.3 Biogenesis of Collagen and Elastin

Fibril collagen molecules are synthesized as a pro-molecule that contains three polypeptide chains. Two pro-collagen endopeptidases, pro-collagen-N-proteinase (PNP, also known as ADAMTS-2) and pro-collagen-C-proteinase (PCP, also known as bone morphogenetic protein-1) cleave the N terminal and C terminal pro-peptides respectfully, to yield a mature tropo-collagen monomer. Both enzymes belong to a family of Zn2+-dependent metalloproteinases [47]. Each molecule of collagen undergoes hydroxylation on lysine and proline residues to form the triple helical structure telo-peptide. In the telo-peptides, copper- dependent enzyme lysyl oxidase (LOX) catalyzes the conversion of lysine and hydroxylase residues to aldehydes. After processing and assembly, the pro-collagen I molecule is secreted into the extracellular space (see Figure 1.1).

Elastin is synthesized by fibroblasts as immature, soluble tropo-elastin monomers. It is secreted to the ECM by secretory vesicles derived from the Golgi apparatus, and then delivered to the microfibrillar site consisting of auxiliary proteins fibrillins and microfibril-associated 11 glycoproteins [53]. Other proteins associated with tropo-elastin are elastin binding protein

(EBP) [56] and fibulin-5 [57, 58] , the latter of which has been shown to be required for elastic fiber development. Crosslinking of immature elastin monomers to mature insoluble tropo- elastin polymers is also performed by members of the LOX family of enzymes (see Figure 1.1).

Specifically, LOX catalyzes the oxidative deamination of lysine to aldehyde residues. The aldehyde residues then spontaneously react with adjacent aldehydes or ε-amino groups of peptidyl lysine to form covalent cross-linkages. LOX-like-1 (LOXL-1) enzyme is also involved in elastogenesis by recruiting fibulin-5 to the tropoelastin monomers [59].

Figure 1.1: Collagen and Elastin Metabolism. The fibroblast is the most common cell that produces collagen and elastin. Collagen is synthesized by the fibroblast and secreted as a pro-collagen molecule to the extracellular space, where it is cleaved by procollagen-N-proteinase (PNP) and procollagen-C-proteinase (PCP) at its N-terminus and C-terminus respectively to yield mature tropo-collagen monomers. Elastin is secreted to the extracellular space as soluble tropo-elastin monomers. Once in the extracellular space, tropo-collagen and tropo-elastin monomers are cross-linked by LOX enzymes, also secreted by fibroblasts, to yield stable mature polymers. MMP enzymes are the major class of enzymes that degrade mature collagen and elastin polymers into their respective monomers. MMP enzyme activity is regulated by the Tissue Inhibitors of Matrix Metalloproteinases (TIMPs) family. A balance between the synthesis and degradation of mature, stable collagen and elastin is necessary to yield a mature, stable ECM. It is believed that this balance is tipped to result in increased degradation in women with POP.

1.2.1.4 Lysyl Oxidases Family of Enzymes

To date, there are five known members of the LOX family of enzymes, which are the copper-dependent monoamine oxidases secreted by fibroblasts and SMCs. The prototypic LOX is the most studied member of this family, while the individual roles of the LOXL-1-4 remain unclear [59]. All five members of the LOX family of enzymes share a common conserved C- terminus catalytic domain. The LOX gene encodes for a 417 amino acid protein, including a signal peptide of 21 amino acids [60]. N-glycosylation of the protein, incorporation of copper and cleavage of the signal peptide produces a 50 kDa pro-enzyme [61]. The pro-enzyme (pro-

LOX) is then secreted into the extracellular space, where it is cleaved by a PCP [62] and the related tolloid-like-1 and tolloid-like-2 enzymes [63] to yield the mature, non-glycosylated 32 kDa protein. LOXL-1 is also synthesized as a pro-peptide, and is converted to the active enzyme by the cleavage of the N-terminal pro-peptide. The variable domains of LOXL-2, -3, -4 contain highly conserved scavenger receptor cysteine-rich domains, present in numerous cell surfaces and secreted proteins, and thought to mediate cell adhesion and host defense [64].

LOX protein is highly expressed in connective tissue containing collagen and elastin fibers, such as the skin, lung, arteries, and vagina due to its main role in catalyzing the polymerization of collagen and elastin monomers to their respective mature polymers.

Importantly, LOX has also been localized to the nucleus of various tissues and has been reported to interact with histones to yield tightly packed chromatin [65-67]. Its interaction with histones has been suggested to affect condensation of chromatin, and hence transcription of genes. For instance, it has been shown to regulate the promoter activity of collagen III

(COL3A1) and elastin [68, 69]. Furthermore, LOX has been described to localize to the cytoplasm of certain cells and tissues in its pro-enzyme and mature enzyme form. Specifically,

14 pro-LOX has been localized to the cytoplasm of differentiated osteoblasts, and to the Golgi apparatus of proliferating osteoblasts [70]. Interestingly, the ECM linker molecule, fibronectin, has been shown to regulate the activity of LOX [71].

1.2.1.5 Pro-collagen-C-Proteinase / Bone Morphogenetic Protein-1

Pro-collagen-C-Proteinase, also known as bone morphogenetic protein-1 (BMP-1), is a calcium dependent, highly specific, multidomain, zinc endopeptidase. It has a key role in ECM formation by the proteolytic processing of the C-pro-peptides of collagen I-III to yield mature fibrillar collagens. The cleavage of the C-pro-peptide is a rate-limiting step in the synthesis of collagen. BMP-1 is the prototype of a small group of proteases that play key roles in the formation of functional, mature fibrillar collagen [72].

The protein domains of BMP-1 consists of an N-terminal pro-domain, a conserved metalloproteinase domain, C-terminal complement –uegf-BMP1 (CUB) domain and epidermal growth factor (EGF)-like domain [73]. The pro-domains are proteolytically cleaved in some cell types in the trans-Golgi apparatus, before secreted to the ECM. The CUB domain mediates protein-protein interactions [74], and the EGF domain is responsible for the binding of BMP-1 to Ca2+, conferring a structural and functional rigidity to the protein [75]. The drosophila protein Tolloid (TLD), involved in dorsa-ventral patterning in gastrulation, was found to have very similar protein domain structure to that of BMP-1 [76]. There are four known mammalian

BMP-1/TLD-like proteinases: BMP-1, mammalian TLD (mTLD), produced by alternative splicing of the BMP-1 gene, and mammalian TLD-like 1 and 2 (mTLL1 and mTLL2); each of the proteinases has some level of PCP activity [77]. Processing of pro-collagen I-III propeptides is cell/tissue specific, and may occur intra-cellularly or extra-cellularly. BMP-1/TLD- like proteinases are molecules secreted in the extra-cellular space; PCP activity and

15 unprocessed pro-collagen was also found in the ECM [78]. Other studies, however, have suggested that PCP activity occurs intra-cellularly [79].

In addition to their role in processing fibrillar pro-collagens, BMP-1/TLD like proteinases also activate LOX [80] and LOXL-1 [81] by cleaving the pro-domains to yield mature, functional enzymes.

1.2.1.6 Pro-collagen-N-Proteinase (PNP)/ ADAMTS-2, -3, -14

The ADAMTSs are a group of secreted proteases that include 19 members in humans.

They are metzincins, or zinc dependent proteases. The structure of the ADAMTS proteins comprise of few conserved domains. Synthesized initially as pre-pro-enzymes, ADAMTSs undergo processing to yield the mature enzymes. When they transit through the endoplasmic reticulum, the signal peptide and pro-domain are cleaved before being released to the ECM

[82].

ADAMTS-2, -3, and -14, also known as the procollagen-N-proteinases (PNP), cleave the smaller N-propeptides from pro-collagen molecules to release the mature triple helical tropo-collagen, which is then ready to be assembled into fibrils. Although the PNPs are structurally similar, their distribution in tissue differs. ADAMTS-2 is the main PNP in skin, and acts on procollagen I, II and III. ADAMTS-3 is a type II procollagen-N-propeptidase, whose expression is much lower than ADAMTS-2 in skin but is 5 times higher that ADAMTS-2 in cartilage [83]. ADAMTS-14 is the major type I-procollagen-N-pro-peptidase in tendon [84].

Defective ADAMTS-2 results in partially processed type-I procollagen consisting of monomers were the C-propeptides are removed, but the N-propeptides remain uncleaved. Persistence of the N-propeptides on the monomers gave rise to highly irregular collagen fibrils with decreased cross-linking and tensile strength due to failure of these fibrils to increase in diameter [85-87].

These results suggest that ADAMTS-2 is essential for maturation of type I collagen fibrils in

16 skin, and that neither ADAMTS-3 nor ADAMTS-14 compensate adequately for ADAMTS-2 deficiency in this tissue. Furthermore, as collagen is a major constituent of the functional ECM, a properly processed collagen is crucial for ECM-cell interactions. Therefore, the presence of defective collagen molecules will affect mechanical stability, as well as physiological events like embryogenesis, cell differentiation, migration and wound repair [88].

Another subset of the ADAMTS family is the aggrecanases, namely ADAMTS-1, -4, -

5, -8, -9 and -15. Aggrecan is a major constituent proteoglycan in cartilage, responsible for the ability of tissue to hydrate and thus resist compression [89]. Aggrecanases are proteolytic enzymes that cleave aggrecan at specific cleavage sites as it exits the matrix, resulting in aggrecan depletion in cartilage. The most efficient aggrecanase in this family is ADAMTS-4; however, ADAMTS-1 and ADAMTS-8 also contribute to the aggrecanase activity in cartilage.

ADAMTS-1 and ADAMTS-8 have also been shown to have anti-angiogenic activity [90], thought to be mediated through their thrombospondin motifs. ADAMTS-13, also known as Von

Willebrand factor-cleaving protease (vWFCP), is another significant member of the ADAMTS family. Its substrate, von Willebrand Factor (vWF), is an essential glycoprotein in plasma, platelets and vascular endothelial cells, and mediates platelet aggregation and adhesion to areas of vascular damage. Deficiency of ADAMTS-13 results in a condition known as thrombotic thrombocytopenic purpura (TTP), characterized by microthrombi, anemia, renal failure and neurological deficiency [82, 91].

1.2.2 ECM turnover and breakdown

The ECM and its components, comprising collagens, gelatin (irreversibly hydrolysed form of collagen), elastin, glycoproteins and proteoglycans (PGs), are under a state of constant remodeling and turnover. This process is mediated by a family of proteolytic enzymes known as the matrix metalloproteinases (MMPs). A group of proteins named Tissue Inhibitors of

Matrix Metalloproteinases (TIMPs) strictly control these enzymes, to offer a tight balance between ECM biogenesis and breakdown [92]. Both are also regulated by other factors, including hormones, cytokines, and growth factors. The balance between MMPs and TIMPs activities is involved in both physiological and pathological events, including wound healing, connective tissue remodeling, angiogenesis, metastasis and inflammation.

1.2.2.1 Matrix Metalloproteinases

MMPs are a 26-member family of calcium dependent, zinc endopeptidases. They have been subdivided into 6 main groups according to their primary substrates (see Table 1.2). The collagenases (MMP-1, -8, -13 and -18) degrade the majority of collagens, most importantly the fibrillar collagens I, II and III. The gelatinases (MMP-2 and -9) degrade gelatin, as well as smaller collagen fragments. The stromelysins (proteoglycanase, collagenase activating proteins

MMP-3, -10 and -11) can degrade smaller collagen fragments, but are also involved in several regulatory functions, including the activation of other MMPs (matrilysin MMP-7 and -26). The

“membrane-type” (MT) MMPs are cell membrane bound, and have diverse functions. For example, MMP-14 is involved in the activation of proMMPs, degradation of ECM, shedding of cell surface molecules, and cell signaling via binding to protein kinases [92, 93].

Table 1. 2: Classification of Matrix Metalloproteinase Enzymes [92] .

Group MMP Substrates

Collagenases Collagenase 1 MMP-1 collagens I, II, III, VII, VIII, X, XI, gelatins Neutrophil Elastase MMP-8 collagens I, II, III, V, VII, VIII, X Collagenase 3 MMP-13 collagens I, II, III, IV, V, VII, IX, X, gelatins Collagenase 4 MMP-18 Gelatinases Gelatinase A MMP-2 gelatins, collagens I, II, III, IV, VII, X, elastin, fibronectin , activates pro-MMP-13 Gelatinase B MMP-9 gelatins, collagens IV, V, VII, X, XI, elastin Stromelysins Stromelysin 1 MMP-3 collagens III, IV, V, VII, IX, X, XI, gelatins, proteoglycans, laminins, fibronectin Stromelysin 2 MMP-10 collagens I, III, IV, V, IX, X, gelatins Stromelysin 3 MMP-11 Matrilysins Matrilysin 1 MMP-7 gelatins, collagens I and IV Matrilysin 2 MMP-26 gelatins, collagens I and IV Membrane-type (MT) MT1-MMP MMP-14 gelatins, collagens I, II, III, activates pro-MMP-2 and pro-MMP-13 MT2-MMP MMP-15 gelatins, collagen III MT3-MMP MMP-16 MT4-MMP MMP-17 MT5-MMP MMP-24 gelatins MT6-MMP MMP-25 Other MMPs Macrophage MMP-12 collagens, gelatins Metalloelastase RASI MMP-19 Enamelysin MMP-20 MMP chromosome 1 MMP-21 gelatins MMP chromosome 1 MMP-22 Human ovary cDNA MMP-23 - MMP-27 Epilysin MMP-28 Unnamed MMP-29 19

The majority of the MMPs structure consists of 4 distinct domains: a N-terminal pro- domain, followed by a catalytic domain, a hinge region, and a hemopexin-like domain at the C- terminus. The latter is responsible for substrate specificity as well as interaction with TIMPs. In addition to these domains, the MT-MMPs contain a transmembrane domain that allows their interaction with the cell surface. MMPs are secreted as zymogens (pro-molecules) by connective tissue cells such as fibroblasts, osteoblasts, endothelial cells, and pro-inflammatory cells, neutrophils, macrophages and lymphocytes. They then undergo proteolytic cleavage to produce their active forms. Normally, the activity of MMPs is controlled either at the transcription level, by activation of the zymogen, or by inhibition of the active form through the activity of TIMPs. Under normal physiological conditions, there is little to no expression of

MMPs. In pathological conditions, this intricate balance is shifted towards an increase in MMP activity, or a decrease in TIMP efficacy, leading to cumulative tissue breakdown [94, 95].

1.2.2.2 Tissue Inhibitors of Metalloproteinases

There are 4 known members of the TIMP family, TIMP1-4, all of which reversibly inhibit MMPs by direct binding. TIMPs consist of two domains, one at the N-terminus and the

C-terminus [96]. The N-terminal domain has sufficient activity to inhibit MMPs, by binding with the Zn-binding site of MMPs in a 1:1 interaction [97] and is highly conserved between

TIMP members. The C-terminal domain is responsible for protein-protein interaction as well as binding to pro-MMPs [98]. The specificity and binding of TIMPs to MMPs appears to be overlapping; the only exception is the inability of TIMP-1 to inhibit MT-MMPs [94].

Besides inhibiting MMPs, TIMPs also regulate angiogenesis and cellular proliferation

[99]. While TIMP-2 appears to be ubiquitously expressed in tissue, the expression of TIMP-1, -

3 and -4 is inducible and is tissue specific. Precisely, TIMP-1 is enhanced in reproductive organ systems, TIMP-3 is enriched in the kidney, thymus and heart and TIMP-4 is highly expressed in

20 the heart, ovary, brain and skeletal muscle [100]. TIMP-1 and TIMP-2 are known to bind to pro-MMP-9 and pro-MMP-2 respectively through the C-terminus domain [101]. TIMP-2 also forms a tri-molecular complex with pro-MMP-2 and MT1-MMP, which allows MT1-MMP to cleave and release active pro-MMP-2 [102]. I recently reported a significant decrease in TIMP-

1 protein expression in POP patients in comparison to non-POP patients. Furthermore, I found that the expression of TIMP-2 protein was 10 times higher than TIMP-1 and TIMP-3 proteins in human vaginal tissues [103]. These results raise the possibility that a decrease in TIMP-1 expression contributes to the development of pelvic floor disorders, and that TIMP-2 may play a pivotal role in facilitating the activation of MMP-2.

1.2.3 Ground Substance

In addition to the structural proteins, the ECM contains a variety of multi-adhesive glycoproteins, and glycosyaminoglycans (GAGs), together known as ground substance. The ground substance occupies the space between the collagen and elastin fibers and the cells, and consists of a viscous, clear substance with high water content. Together with the fibers, the ground substance forms a dynamic and interactive system that anchors the cells within the tissue via cell-ECM adhesion molecules, as well as provides pathways for cell migration, differentiation and cell-cell signaling of biochemical and mechanical changes in the extracellular environment [104].

1.2.3.1 Glycoproteins

The adhesive glycoproteins, fibronectin, laminins, vitronectin, thrombospondin, fibrinogen and others, allow cells to adhere to the ECM. This is mediated via the binding of cells through cell-surface integrin receptors. Interactions between cells and ECM are crucial to many cellular responses, such as cell migration, growth, differentiation and survival. The process is dynamic; cells receive input from their surrounding extracellular environment, and in 21 turn modulate the ECM by secreting growth factors, as well as proteases and their inhibitors

[105].

Fibronectin is a high molecular weight, multifunctional dimeric or multimeric glycoprotein that is ubiquitously expressed in embryonic and adult tissue [106]. The dimeric, soluble plasma fibronectin is synthesized by the liver hepatocytes whereas the multimetric, insoluble, tissue fibronectin is secreted as a soluble dimer by fibroblastic cells and undergoes polymerization in the surrounding ECM. Plasma fibronectin is important for wound healing and thrombosis [107] and it is deposited at the site of injury along with fibrin. Tissue fibronectin is organized as a fibrillar network, and is crucial for cell adhesion, growth, migration, and differentiation. It also contributes to the ECM material stability, and allows interaction of various substrates to cell surface receptors [106]. Fibronectin contains specific functional domains that allow binding to cells via transmembrane receptors (integrins), and interaction with other proteins such as collagen, fibrin and heparin/heparan sulfate [106]. Furthermore, similar to other adhesive glycoproteins such as vitronectin and vWF, fibronectin also contains an RGD (Arg-Gly-Asp) motif that mediates cell adhesion via interaction with cell surface integrin receptors [108]. This interaction signals to cells that adhesion to ECM has occurred, which influences cell survival, metabolism and cell fate. ProNectin, a synthetic analogue of fibronectin, contains the tripeptide RGD cell attachment epitope, essentially enabling binding to adhesion receptors on the cell surface [109].

Integrins are large, heterodimeric transmembrane proteins, consisting of α and ß subunits, each with a large extracellular domain, a transmembrane domain, and a cytoplasmic domain. In humans, 18 α and 8 β subunits have been characterized. Through different combinations of α and β subunits, 24 unique integrins have been identified, although the number varies according to different studies. Integrins mainly act as receptors that allow the

22 interaction of ECM glycoproteins and other extracellular ligands with the cell to regulate intracellular signal transduction. Intriguingly, many of the integrin-triggered signaling pathways are very similar to the pathways triggered by growth factors, both of which require cells to be adherent. The majority of integrins are present in an inactive state. Activation occurs by either binding of protein talin or kindlin to the cytoplasmic domain of the ß tail, changing it from a

“bent” to an “unbent” conformation, and essentially activating the integrin from the inside-out

[110]. This allows for the integrin then to exercise high affinity binding to its ligand [111]. The interaction of fibronectin with integrins results in “outside-in” activation of integrins, and allows for cytoskeleton reorganization, actin microfilament assembly, focal adhesion formation, and fibronectin matrix assembly [106].

Laminins are major cell adhesive proteins that self-assemble into polymers on the cell surface[112]. They are large, heterotrimetric glycoproteins composed of three polypeptide chains: α, ß and ϒ, which can assemble into different combinations to create laminin variants.

Through self-polymerization, laminins form filaments and layered sheets that initiate basement membrane assembly. The network of collagen IV defines the basement membrane scaffold that integrates the laminins, PGs and other components, to form the highly organized tissue specific architecture necessary for cellular interactions [113]. Therefore, if this process is inhibited, basement membrane assembly is disrupted. Additionally, laminins have many roles in development and disease, and mediate cell adhesion, proliferation, migration, differentiation metastasis and angiogenesis [114].

Vitronectin is an adhesive glycoprotein present in blood plasma, amniotic fluid and urine, and in ECM of many tissues [115]. In humans, vitronectin is synthesized by the liver hepatocytes. It interacts with the ECM via its collagen- and heparin-binding domains, and with cells through its RGD integrin-binding domain. The main roles of vitronectin are in wound

23 healing, tumor growth and metastasis, and viral infection [116, 117].

1.2.3.2 PGs and Glycosyaminoglycans (GAG)

Tissue adhesiveness of the ECM is provided by PGs, which are heavily glycosylated proteins consisting of a “core protein” covalently linked to one or more GAG chains. PGs also serve as the major contributors to the viscoelastic properties of tissue. The GAG chains differ between different classes of PGs, and can be composed of a variety of proteins such as chondroitin, keratin, and heparan. Smaller PGs, such as decorin, fibromodulin, biglycan and chondroadherin, interact directly with the structural proteins collagen and elastin. This forms a network of fibers that work to prevent the compression of the ECM by trapping water molecules [118]. The smaller PGs also interact with growth factors that influence cell adhesion, migration and proliferation, and contribute to the turgor and viscoelastic pressures [119]. The larger PGs, such as versican and aggrecan, stabilize the ECM and contribute to the structural framework and spatial arrangement of proteins of the ECM [120].

1.2.4 Cell-Cell and Cell-Matrix Adhesion Molecules.

The selectin family of proteins consists of three closely related cell-surface molecules that are differentially expressed by leukocytes (L-selectin), vascular endothelium (E and P-selectin) and platelets (P-selectin). What distinguishes selectins from other adhesion molecules is that selectin function is restricted to leukocyte interaction with the vascular endothelium. Selectins play a significant role in regulating inflammatory processes via their interaction with other adhesion molecules and inflammatory mediators. Current selectin-directed therapeutics have been shown to be effective in blocking many pathological effects that result from leukocyte attachment and rolling to the sites of inflammation [121].

1.3 Pelvic ECM and POP Development.

Collagens I, III and V fibrils have been described in the vaginal wall. Using immunohistochemical analysis of the anterior vaginal wall, Lin et al [122] reported a statistically significant decrease in the density of collagen type III in women with POP in comparison to non-POP women after controlling for age, weight, parity, SUI and menopause status. Diminished collagen levels have also been found in multiple pelvic tissues of women with POP and SUI, including the round ligaments [123], uterosacral ligaments [124], anterior vaginal wall [125] and endopelvic fascia [126]. Salman et al [127] examined samples obtained from cardinal ligaments of women with and without POP, and concluded that women with prolapse have less dense ECM with loosely arranged connective tissue fibers compared to non-

POP women. Further investigation with electron microscopy showed that women with POP had less orderly, and loosely packed larger collagen fibrils than their non-POP counterparts.

Weakened pelvic floor tissue may also result from altered distribution of collagen fibers within the ECM; a disorderly arrangement of collagen fibers was observed in connective tissue samples from peri-urethral specimens of women with SUI in comparison to asymptomatic women [128].

Studies have reported conflicting results with respect to collagen turnover in connective tissue of women with PFDs. Studying markers of collagen synthesis and breakdown from sub- urethral tissue of women with and without POP, Edwall et al [129] reported increased procollagen type I carboxy-terminal pro-peptide (PICP) and pro-collagen type III amino-terminal pro-peptide (PIIINP) in women with POP in comparison to non-POP women, indicating an increase in collagen synthesis following collagen breakdown. Another study reported similar levels of PICP and PIIINP, but a lower content of hydroxyl-proline in uterosacral ligaments of women with POP [130]. However, Chen et al [126] examined the collagen content of fibroblast cultures from skin of women with and without POP and failed to find a difference in levels of

25 collagen synthesis. Collagen protein content in tissue has traditionally been assessed by immunohistochemistry and Western immunoblot techniques. Kannan et al [131] compared the histological changes between prolapsed and non-prolapsed vaginal skin, and found increased myofibroblast differentiation in prolapsed tissue, with increased fibrosis and condensed appearance of collagen fibers. More recently, collagen synthesis has been assessed using fibroblast cultures and specific markers, which allows the examination of the dynamic process of collagen synthesis and degradation. Despite controversial findings, the general consensus is that a decrease in the overall total collagen in pelvic floor tissue exists in women with POP, along with an increase in the collagen III: collagen I ratio and in immature collagen, suggesting weaker connective tissue in women with PFDs.

The expression and content of elastin in pelvic floor tissue has also been studied to further understand the pathophysiology of POP and other PFD’s. Using immunofluorescent staining, elastic fibers were found to be fragmented or even undetectable in uterosacral ligaments of women with POP [132]. Karam et al [133] confirmed that elastic fibers expression were smaller, fragmented and decreased in vaginal tissue of women with POP. Zong et al [134]

, however, reported an increase in the amount of tropo-elastin and mature elastin in women with

POP in comparison to non-POP women. Elastin content and metabolism were also found to be important in SUI etiology. For instance, a three-fold increase in systemic elastase activity has been reported in women with SUI in comparison to non-POP women, possibly promoting collagenolysis [134-136]. Furthermore, an increase in neutrophil elastase (NE) activity and a lower expression of an elastase inhibitor α-1-anti-trypsin (ATT) has been found in women with

SUI. Additionally, lower levels of fibrillin-1, an essential elastin scaffold protein, have been reported in peri-urethral tissue of women with SUI, which may result in a decrease in elastin deposition [137]. In conclusion, a decrease or disordered elastin content, coupled with an

26 increase in active elastin remodeling, may result in weakened elastin content in pelvic tissue of women with POP and SUI.

Man et al [138] studied the expression of the modulators of elastin in vaginal tissue of women with POP, namely ATT, NE, and LOXL-1. They concluded that the expression of these proteins varied between individuals and depended on the site of sample collection (the anterior or posterior vaginal wall). This observation highlights the importance of consistency in sample selection to ensure data reproducibility. Moalli et al [38] studied the activity of MMP-2 and MMP-9 in POP patients, and found an increase in MMP-9 activity, but a decrease MMP-2 activity, which could indicate the higher rate of remodeling in vaginal tissue of women with

POP.

Differential expression of the LOX family of enzymes in vaginal tissue of women with

POP was reported by our group [139]; LOX, LOXL1-4 gene expression as well as LOX, LOXL-

1 and LOXL-3 protein expression was reduced in vaginal samples of women with POP, which may result in defective ECM protein synthesis and assembly. Klutke et al [140] also observed a similar reduction the LOX family gene and protein expression in uterosacral ligament biopsies from women with POP, as well as an increase in FIB-5 mRNA [141] .They also identified 66 methylated CpG sites on the LOX gene promoter in the POP group, in comparison to only one methylated CpG site in the non-POP group [140]. However, several studies have reported a decrease in FIB-5 in biopsies from several pelvic floor tissue sites, including the anterior vaginal wall [142], uterosacral ligaments [143] and paraurethral tissue [144].

Our groups has previously studied the role of pro-collagen C and N proteinases in POP pathophysiology and observed that the mRNA expression of BMP-1 gene was decreased in women with POP in comparison to non-POP women. In particular, the expression of 130 kDa,

92.5 kDa, and 82.5 kDa isoforms of BMP-1 protein were down-regulated in postmenopausal

27 patients, whereas the 130 kDa isoform expression was up-regulated in premenopausal patients when compared with aged-matched non-POP women [35]. Furthermore, we reported that the expression of the 58 kDa isoform of ADAMTS-2 was up-regulated in patients with POP when compared to non-POP patients [103]. ADAMTS-\- knockout (K/O) mice develop fragile skin similar to that seen in dermatosparaxis in cattle and in patients with EDS type VIIC, which reflects weakened ECM and aberrant collagen fibril formation [44]. These findings, together with the well-known association between EDS and POP, suggest that deregulation of BMP-1 and ADAMTS-2 may contribute to deficient vaginal connective tissue, resulting in POP.

Other animal studies from targeted gene disruption models have also provided insight into the role of collagen and elastin metabolism in the pathophysiology of POP. Deficiency of

LOXL-1 led to severe POP in mice shortly after vaginal delivery, accompanied by marked weakness in the vaginal wall as well as SUI symptoms and paraurethral pathology [144, 145].

Furthermore, FIB-5 K/O mice develop POP shortly after pregnancy and vaginal delivery [146].

The vaginas of FIB-5 K/O mice exhibited increased distensibility and decreased maximal stress and stiffness [147]. Not only does FIB-5 enable assembly of elastic fibers, it also inhibits

MMP-9 mediated elastogenolysis in an integrin-dependent manner [148]. From the results of these studies, it was concluded that the intact synthesis and deposition of elastic fibers is necessary for recovery of the pelvic floor following vaginal delivery.

Several laboratories explored MMP expression and activity in the vaginal epithelial tissue of women with POP in comparison to non-POP women. Connell et al [124] reported a two-fold increase in MMP-2 gene expression in women with POP. Two other groups found that there was an 80% increase in the expression of MMP-1, coupled with a decrease in the expression of TIMP-1 [127, 149]. Jackson et al [150] and Alarab et al [103] described an increase in MMP-2 and MMP-9 activity in anterior vaginal tissue of women with POP in

28 comparison to non-POP women, concluding these reflected an increase in ECM turnover. Two other studies confirmed these results with immunohistochemical staining, reporting increased activity of pro-form as well as active form of MMP-2 in vaginal tissue of women with POP

[151]. Our group further showed an increase in both pro and active MMP-12 protein expression, coupled with a decrease in TIMP1-4 gene and TIMP-1 protein expression in anterior vaginal tissue of women with POP in comparison to non-POP women [103].

Using different methodologies, it has been established that women with SUI and POP have an increase in MMP expression and activity that may result in accelerated collagen and

ECM breakdown. With respect to markers of collagen synthesis and breakdown, Kushner et al

[152] described an increase in the concentration of helical peptide α1, a collagen breakdown product, in the urine of women with SUI, thus concluding increased degradation in urogenital tissue. Edwall et al [129] supported this theory by showing that women with SUI have a lower level of serum PICP and tissue collagen Type I carboxyterminal telopeptide (ICTP) in comparison to their non-POP counterparts.

PGs are involved in fibrillogensis of collagen fibers. An imbalance in PGs content can interfere with the formation, maintenance and destruction of collagen and possibly other ECM components [128]. Specifically, increased amounts of small leucine-rich proteoglycans (SLRP) such as decorin and fibromodulin have been reported in periurethral tissue of women with SUI

[128-130]. Another study reported the opposite results with respect to the transcript levels of decorin and lumican in pelvic floor tissue, which indicates altered PGs content in ECM of women with PFDs [128].

1.4 The Effects of Stretch on Pelvic Floor Tissue 1.4.1 Vaginal Human Tissue and Mechanical Stretch

Several studies have looked at the response of human tissue to stretch. Abramowitch et al [30] first defined the properties of vaginal wall tissue essential for testing stretch-induced changes. These studies established that vaginal tissue had to be anisotropic, or directionally dependent, in nature and exhibiting viscoelastic (both viscous and elastic) properties. Jean-

Charles [36] and Rubod [153] et al used post-mortem vaginal tissues from women with severe

POP and from women that had normal vaginal support, and subjected those tissues to cyclical loading and uniaxial testing. They found that tissues obtained from POP patients showed increased elasticity but only under large deformation, and that tissue obtained from the posterior wall of the vagina of POP was more rigid in nature when compared to the anterior wall of the vagina. Using suction-based devices to measure the biomechanical properties of dermal tissue, Epstein et al [154] confirmed that women with POP had significantly more extensible vaginal tissue than women with normal pelvic support. Another group [155] compared the properties of vaginal wall tissue obtained from pre and post-menopausal women undergoing vaginal hysterectomies, and reported that when subjected to uniaxial stretch, tissue from post-menopausal women was significantly more elastic in comparison to tissue obtained from premenopausal women. These results were explained by the higher collagen III content in postmenopausal tissue.

1.4.2 Animal Models of Vaginal Stretch

Many research groups have attempted to understand the behavior of vaginal tissue in response to stretch via the use of animal models, including rats and non-human primates [156-

158]. Alperin et al [144] compared the distensibility of vaginal tissue samples from wild type

(WT) and LOXL-1 K/O mice. They found that tissue obtained from LOXL-1 K/O mice failed at 69% of the uniaxial load that tissue from WT mice could withstand. Rahn et al [146] compared the characteristics of tissue collected from FIB-5 K/O mice, non-pregnant and

30 pregnant WT mice. Vaginal tissues were formed into ring-like structures, and subjected to uniaxial stretching until breakage or steady-state distension. They reported that tissue obtained from FIB-5 K/O mice behaved similarly to tissue from pregnant WT mice, with decreased stiffness and maximal load, however with increased distensibility and vaginal diameter when compared to non-pregnant WT mice [146].

The tangent modulus describes the behavior of materials stressed beyond their elastic properties by quantifying the “softening” of a material before it breaks. Feola et al [158] obtained tissue from nulliparous and parous rhesus macaques, and compared how these tissues reacted to uniaxial loading. They established that tissue from parous animals had a significantly decreased tangent modulus and tensile strength. Feola et al also quantified collagen ratios and alignments, and found that although there was no difference in the ratio of collagen I to collagen III/V in these tissues, collagen fibers were more disorganized with increased parity.

Since worsening pelvic organ support correlates negatively with decreased collagen alignment and mechanical properties, this likely predisposes such tissue to the development of POP [158].

1.4.3 Cellular Response to Mechanical Stretch and POP

Fibroblasts derived from women with POP seem to be preconditioned to the production of abnormal prolapsed matrix proteins. They do not respond to changes in their environment in the same way as non-POP fibroblasts, and thus are not able to restore ECM homeostasis. This in turn may result in an accelerated deterioration of the ECM, resulting in loss of tissue strength and tissue damage.

Several studies have looked at how pelvic floor fibroblasts respond to mechanical loading. It has been established that these cells respond to mechanical stimuli by remodeling their actin cytoskeleton, and that their alignment is perpendicular to the force after 48 hrs of cyclical mechanical stretch. This is especially true in the presence of a collagen I matrix [43,

159, 160]. However, differences between cell populations were seen after 24 hrs of stretch on collagen I substrates, where alignment of F-actin fibers from fibroblasts derived from vaginal tissue of women with POP was delayed in comparison to fibroblasts derived from non-POP women. Furthermore, mechanical loading induced activation of MMP-2 by fibroblasts in a time dependent manner up to 24 hrs. At a later time point of 48 hrs, levels of pro-MMP2 were similar between fibroblasts derived from POP patients and non-POP patients, and no active

MMP-2 was found in the conditioned media [160]. Zong et al [43] subjected vaginal fibroblasts plated on collagen-I-coated plates to cyclical stretch at a magnitude of 8 and 16% for 72 hrs.

They found that collective collagenase activity of MMP-1, -8 and -13 increased in the presence of mechanical stretch, and this activity correlated linearly with the magnitude of stretch.

Blaauboer et al [161] studied how primary lung fibroblasts from women with POP respond to cyclical mechanical stretch. They showed that stretch reduced the mRNA levels of the myofibroblast marker α-smooth muscle actin, as well as Collagen-I, -III, -V and Tenascin

C. Changes in the vaginal wall at the cellular level could indicate differential tissue behavior between women with and without POP, and should be considered when polymeric meshes or other artificial substrates are used to treat POP or SUI patients.

POP fibroblasts also respond differently to surface substrates in comparison to non-POP fibroblasts. Fibroblasts from POP patients showed decreased capacity to remodel the ECM; total levels of MMP-2 released by POP fibroblasts were lower than those released by non-POP fibroblasts, irrespective of surface substrate or exposure to stretch. Furthermore, POP fibroblasts displayed significantly lower gene expression of COL1A1 and COL3A1 than non-

POP fibroblasts, and gene expression of COL1A1 was down regulated by mechanical stretch only in non-POP fibroblasts. When the fibroblasts were plated on collagen-I coated plates and subjected to mechanical stretch, POP fibroblasts showed a decrease in COL1A1, COL3A1 and

MMP-2 gene expression. In addition, POP fibroblasts showed delayed cell alignment and rearrangement of F-actin fibers on collagen-I coated plates when compared to healthy fibroblasts [160].

Cell-matrix interaction seems to be impaired in fibroblasts derived from POP patients, and this may provide important cues for designing artificial polymers that appropriately mimic the vaginal ECM environment, and incite the restoration of the ECM metabolic balance.

1.4.4 Cell-Based Tissue Engineering and POP

Over the last decade, urogynecologists have resorted to the use of surgical implants to reinforce the weakened pelvic floor tissue and improve the outcome of pelvic floor surgery.

This approach was based on the successful use of synthetic mesh in the treatment of abdominal hernias, and synthetic vaginal tapes in the treatment of SUI. The majority of the implants used in the treatment of POP are composed of synthetic material, but biodegradable synthetic and biological implant options are also available. Even though these implants have been widely used, there still remains controversy on their efficacy [162], especially with the reported rates of up to 10% for complications such as erosion, pain, vaginal stenosis and infection [163].

Recently, attention has turned to the use of cell-based tissue engineering to compliment or replace the use of implants. Preclinical studies using autologous muscle-derived stem or progenitor cells cultured in vitro and injected to assist in the regeneration of the urethral sphincter to treat SUI have shown promising results [164]. Outcomes from clinical studies on

SUI patients have also been favorable, with 20-50% of patients becoming continent, however controversy remains since this method is still in its infancy and long-term results are lacking

[165].

The use of cell-based tissue engineering in the treatment of POP is a much more complicated process. Vaginal tissue consists of many different cell types, and the growth of

33 cells requires a scaffold for cell adhesion, which would also function as a support to the weakened pelvic floor. Thus the injection of one cell type without a scaffold will most likely fail to regenerate the vaginal tissue [166, 167]. Because of their accessibility, muscle-derived stem cells were the first to be explored in cell-based therapy for the treatment of POP in an animal model. When implanted on scaffolds in the rat vagina, skeletal muscle derived stem cells differentiated into SMCs, which is more inherent to the vaginal wall [166]. Boennelycke et al [168] introduced the use of fresh muscle fiber fragments as an alternative to the high cost stem cell method. After 8 weeks, they found that new striated muscle was formed and the synthetic scaffolds that were implanted in the rat abdomen disappeared.

The use of fibroblasts in conjunction with synthetic scaffolds has also been explored as a treatment for POP. This next step was understandable, since the connective tissue of the vaginal wall consisted primarily of fibroblastic cells. Tissue engineered fascia was successfully created by implanting human vaginal fibroblasts seeded onto synthetic scaffolds subcutaneously on the back of mice. Human fibroblasts derived from the vagina have also been seeded on various synthetic and biological meshes [169]. Vaginal fibroblasts isolated from women with POP have been successfully re-differentiated into induced pluripotent stem cells

(IPSC) and back to fibroblasts [170]. Whether those approaches can be used for the treatment of POP in humans remains to be clarified. However, since prolapse is the herniation of the involved organs through the vaginal wall, one can deduce the effectiveness of tissue-based engineering on the treatment of POP by looking at the studies that explored this approach to treat hernias [171, 172]. Implants have been used to reinforce hernias during surgery, and similar implants have now been tested in the treatment of POP in animal models [173].

1.5 Rationale and Hypothesis 1.5.1 Rationale

Numerous factors have been attributed to the development of POP. Childbirth and parity are considered to be the leading risk factors due to the injury sustained to the pelvic floor at the time of vaginal delivery. However, only some parous women develop POP of varying degrees, and usually years to decades following NVD. Currently, treatment of POP has been dominated by surgical repair with vaginal meshes, which generally results in good anatomic and subjective outcomes. However, the use of meshes is associated with considerable risk of erosion, pain, infection and vaginal stenosis. The formation of tissue-engineered fascia results from the interaction of vaginal tissue cells with synthetic scaffolds. The potential goal of regenerative medicine is to develop a bio-resorbable scaffold which, in combination with progenitor cells, will be able to contribute to tissue regeneration by proliferation and differentiation into fibroblasts (or myofibroblasts), thereby forming functional connective tissue. It is currently unknown whether autologous cells (myoblasts and fibroblasts) will be used. It is suggested that the successful outcome of vaginal reconstructive surgery will depend on utilizing an optimized

ECM substrate, such as collagen, to promote the cellular growth and distribution of human vaginal fibroblasts (VFs) with a high proliferation potential that would also exhibit rates of protein synthesis similar to non-POP VFs [174]. Therefore, research characterizing cellular component of vaginal tissue to improve surgical outcome is urgently needed. The question remains as to whether there are significant differences between the cellular and biological characteristics and the ability to attach to different synthetic substrates, and produce/modulate the ECM of vaginal fibroblasts derived from patients with severe prolapsed uteri and non-POP

VFs. I propose that the biological characteristics of primary VFs derived from pre-menopausal women with severe POP are distinct from that of VFs derived from non-POP patients. It is likely that fibroblasts from women with POP will not be able to produce an appropriate high- quality ECM due to intrinsic defects. Therefore, by studying the attachment, proliferative

35 ability and production of ECM proteins of VFs derived from POP patients in comparison to non-POP patients, I can optimize conditions that are used during surgery to generate sufficiently strong new tissue by replacing defective vaginal cells with healthy autologous

(from a different body part) or allogeneic fibroblast cells.

Other risk factors that predispose women to the development of POP, are associated with an increase loading of the pelvic floor, specifically chronic conditions i.e. obesity, asthma, constipation and heavy lifting. I propose therefore that chronic mechanical stretch of the vaginal connective tissue affects the expression of ECM proteins, collagen and elastin, their modulators and receptors, culminating in the failure of structures supporting the pelvic organs and the development of POP. I aim to compare the reaction of cells derived from POP and non-

POP patients to artificial mechanical loading. By exploring the cause-and-effect relationship between chronic stretch and POP, I hope to provide strategies to identify women at risk of developing POP and to determine whether risk factor modification will prevent the development of pelvic floor disorders.

1.5.2 Hypothesis and Objectives

I hypothesize that 1) primary VFs derived from pre-menopausal patients with severe

POP display different phenotypic characteristics in comparison to VFs derived from non-POP patients. 2) Continuous static mechanical stretch of VFs will change the expressions of ECM proteins and enzymes regulating ECM biogenesis and biodegradation.

The two main objectives of this thesis work are: 1) To compare the biological characteristics of VFs derived from patients with severe POP and non-POP VFs: cell morphology, attachment, proliferative ability, production of ECM proteins and their cell adhesion receptors; 2) To use the computer-controlled Flexcell in vitro stretch system to test the

36 effect of static mechanical stretch (25% elongation) on the expression of ECM proteins, and cell adhesion proteins, their receptors and modulators.

CHAPTER 2: MATERIALS and METHODS

2.1 Patient Selection

The Institutional Review Board of Mount Sinai Hospital at the University of Toronto approved the study. All clinical samples were collected at Mount Sinai Hospital, Toronto.

Written consent (Appendix A) and clinical data (Appendix B) were obtained from each patient at admission to the hospital for surgery. Strict criteria were determined to select a homogenous group of pre-menopausal women with severe POP and a non-POP group. Inclusion criteria: premenopausal adult women undergoing pelvic surgery for POP equal or greater than stage 3 by POP-Q were identified as patients. Non-POP patients were identified as premenopausal adult women with POP-Q at stage 0 undergoing abdominal hysterectomy for indications other than prolapse. Exclusion criteria: women with a history of gynecological malignancy, connective tissue disorders, emphysema, endometriosis, steroid therapy in the past six months and prior prolapse or incontinence surgery.

2.2 Tissue Collection

In all POP and non-POP patients, 1 cm2 full thickness vaginal tissue was collected during pelvic floor surgery from the anterior middle portion of the vaginal vault (Appendix C) to account for variations in stretch conditions and variations of muscularis thickness throughout the vaginal length. The tissue was placed in ice-cold basic salt solution HBSS-/- (Hank’s buffered salt solution without Ca2+ and Mg2+) which was prepared and autoclaved in-house by the media preparation department at the Lunenfeld-Tanenbaum Research Institute (LTRI). The tissue was then immediately transferred from the operating theatre to the tissue culture facility at LTRI for primary cell derivation.

2.3 Derivation and Maintenance of Primary Human Vaginal Fibroblasts

The process of primary VF derivation from vaginal biopsy samples from POP and non-

POP women was started within one hour of tissue collection during the surgery. Vaginal tissue was washed in ice-cold HBSS-/- three times, cut into small pieces (1 mm3) and placed in “an enzymatic digestion buffer” [1mg/ml Bovine Serum Albumin Fraction V (Fisher Scientific,

ON, Canada) supplemented with 2 mg/ml Collagenase Type IA (Sigma Aldrich, ON, Canada) and 0.15 mg/ml DNase 1 (Roche Applied Science, QC, Canada)] for 90 min at 37°C on shaker.

Every 30 min, tissue pieces were mechanically disrupted by vigorous pipetting. The suspension of isolated primary VFs was pelleted at 250 g for 8 min at 4°C, washed with HBSS-/- once, re- suspended in cell culture media [phenol red-free DMEM (Sigma, MO, USA) supplemented with 20% fetal bovine serum (FBS, Gibco, ON, Canada), 25 mM HEPES, 50mg/L Normocin

(Invivogen, CA, USA), 4g/L D-Glucose (Sigma, ON, Canada), 20 mM L-Glutamine (Life

Technologies, CA, USA)] and then plated on 10 cm tissue culture plates (Sarstedt, Germany) in a humidified incubator with 5% CO2 at 37°C to promote fibroblast attachment (see Figure 2.1).

After 18-24 hrs, culture plates were washed five times with HBSS-/- to remove un-attached cells. Usually after 2-6 weeks, primary VFs reach 90% confluence and were passaged. Primary

VF cell lines were cultured in DMEM supplemented with 20% FBS (see above). Cell culture media was changed every 2 to 3 days. All experiments conducted in this study were performed using cells between passage 2 and 4.

Figure 2. 1 Derivation of Primary Human Vaginal Fibroblasts. (1) During hysterectomy, 1 cm2 full thickness vaginal tissue was obtained from the

anterior vaginal wall. (2) Vaginal tissue was cut into small pieces, (3) enzymatically

digested in buffer at 37°C and mechanically disrupted by pipetting. (4) The suspension of

isolated primary cells was then pelleted and re-suspended in cell culture media. (5)

Finally, fibroblasts were plated on tissue culture plates and cultured at 37°C in 5% CO2.

2.4 Comparing the Biological Characteristics of Human Vaginal Fibroblasts

2.4.1 Fibroblast cell verification by indirect immunofluorescence

I employed specific biomarkers of fibroblastic versus myogenic origin via immunofluorescent staining. Primary VFs derived from premenopausal POP and non-POP patients on passages 0, 1 and 2 were cultured in 8-well chamber slides until 50% confluence.

The cells were then fixed with 4% paraformaldehyde for 15 min at room temperature (RT), permeabilized with 0.1% Triton X-100 (Sigma-Aldrich, MO, USA) for 30 sec, and blocked with Serum Free Protein Block (DAKO, ON, Canada) for 30 min at RT. The cells were then incubated with a specific antibody for fibroblasts biomarker, mouse anti-human Vimentin (Sc-

7558, Santa Cruz, Tx, USA), and with antibodies for myogenic biomarkers: mouse anti-human

Smooth Muscle Actin (M0851, DAKO), anti-Desmin (M076029, DAKO) and anti-H-

Caldesmon (Sc-58703, Santa Cruz Biotechnology) or epithelial biomarker anti-Cytokeratin

(M0821, DAKO), dilution 1:100 in PBS for 18 hrs at 4°C. VF cells were then washed in PBS, incubated with biotin-linked secondary antibody (GE Healthcare), or Streptavidin-Flourescein

(DAKO) plus DAPI (DAKO) in the dark for 40 min. After washing with PBS, the slides were mounted with Cytoseal XYL (Ricard-Allan Scientific, Kalamazoo, MI), viewed under a fluorescent microscope (Leica Microsystems, Richmond Hill, ON, Canada) and photographed with a Sony DXC-970 MD (Sony Ltd., Toronto, ON, Canada) 3CCD color video camera.

To quantitate the number of cells in culture that expressed the aforementioned biomarkers, I manually counted the number of VFs from passage 2 that expressed Vimentin,

Desmin and SM-actin per 100 VFs.

2.4.2 Cell Attachment on Different Extracellular Matrices

The Human ECM Cell Adhesion Array Kit from Millipore (MA, USA) was used to assess the attachment of primary VFs. Each 96-well plate contains 12 x 8 well strips coated with 7 different human ECM proteins (Collagen I, Collagen II, Collagen IV, Fibronectin,

Laminin, Tenascin, Vitronectin) and one Bovine Serum Albumin (BSA) coated well as a negative control.

13 primary VF cell cultures derived from 7 POP and 6 non-POP patients at passage 2 were used for this study. Confluent cultured cells were incubated with Trypsin-EDTA (Gibco,

ON, Canada) for 5 min at 37°C followed by gentle tapping. Next, trypsin activity reaction was stopped by adding DMEM/10% FBS. Cells were further detached mechanically by repeated pipetting, centrifuged and washed three times in HBSS-/- at 400xg for 5 min at RT. The cell pellets were re-suspended in 5 mls of provided Assay Buffer. Each primary VF cell line suspension was counted on CASY®Cell Counter (Roche Applied Science, QC, Canada) and then diluted to a final concentration of 1x106 cells per ml. Before use, each well of the ECM

Array plate was rehydrated with 200 µl of PBS at RT for 10 min. The PBS was then removed and 100 µl of each cell suspension were added in triplicates to 7 ECM-coated wells and the control well. The plate was then covered and incubated for 1 hr at 37°C in a CO2 chamber.

Following incubation, the medium was gently aspirated from the wells. Without allowing them to dry, the wells were washed three times with 200 µl of Assay Buffer per well. Next, 100 µl of provided Cell Stain Solution were added to each well, and incubated for 5 min at RT. The stain was then gently removed from the wells by aspiration, and the wells were washed five times with 200 µl of deionized water. The final wash was discarded and the wells were allowed to air dry. Next, 100 µl of Extraction Buffer was added to each well. The plates were incubated in the dark under gentle rotation on an orbital shaker at RT for 10 min, until the cell-bound stain was

43 completely solubilized. Finally, the absorbance was determined at 560 nm on a microplate reader (uQuant Biotek, VT, USA).

2.4.3 Cell Proliferation on Different Extracellular Matrices

I employed the Thiazolyl Blue Tetrazolium Bromide (MTT) assay to investigate the effect of different ECM on the proliferation ability of 10 primary VFs cell lines (5 were derived from POP patients and 5 from non-POP patients). All VF cell lines used for the proliferation study were at passage 2. I used 24-well black HT Bioflex culture plates coated with Collagen I

(HTPB-3001C) or ProNectin (HTPB-3001P) from Flexcell (Flexcell International Corp., NC,

USA). Cells were trypsinzed, centrifuged and washed in HBSS-/- as described above. The cell pellets were re-suspended in DMEM/20% FBS to a final concentration of 5x103 cells/ml. One ml from each primary VF cell line (5000 cells) was plated in quadruplicates on the HT Bioflex culture plate. Four wells from each plate contained only medium to be used as a background control. Identical plates for both ECM substrates were prepared to study proliferation of cell lines derived from POP and non-POP patients over the span of five days.

MTT solution (Sigma-Aldrich, MO, USA) was prepared to a final concentration of 5 mg/ml MTT in DMEM/20% FBS. The solution was filter sterilized, aliquoted and kept frozen in the dark. After 1, 2, 3, 4 and 5 days in culture, frozen MTT solution was thawed to 37°C, and

0.5 mls of MTT was added to each well including the control (media only) wells. The plate was then incubated at 37°C for 3.5 hrs in a CO2 chamber. Next, the medium was carefully aspirated, and 1 ml of DMSO (Sigma, MO, USA) was added to each well. The plate was incubated in the dark under gentle rotation on an orbital shaker at RT for 10 min. Finally, the absorbance was determined at 590 nm on a microplate reader (µQuant). The procedure was repeated at the same time every 24 hrs for five consecutive days.

2.5 Application of Static Mechanical Stretch

The impact of mechanical stretch on the expression of ECM proteins and their modulators was investigated in vitro using primary VF cell lines and a computer-driven

Flexcell vacuum system (FX-5000, Flexcell International Corp, NC, USA). VFs cells from passages 2 or 3 were seeded at a density of 350,000 cells/well into the flexible-bottom

Collagen-I-coated 6-well Flexcell culture plates with 2.5 ml of medium containing

DMEM/20% FBS. Once the cells reached approximately 80% confluence, they were washed twice with HBSS-/- and then serum starved O/N with 5 ml of serum-free DMEM (SF-DMEM) supplemented with 1X Insulin-Transferrin-Selenium-Sodium Pyruvate Solution (ITS-A, Gibco,

ON, Canada). The next morning, static stretch of 25% elongation was applied for 6, 24 or 48

o hrs inside a humidified incubator with 5% CO2 at 37 C. An identical non-stretched plate was kept in the same incubator for the same time interval. Once the protocol was completed, conditioned medium (CM) from POP and non-POP patients, non-stretched (NS) and stretched plates (S), were collected, centrifuged at 1000xg for 10 min at 4oC and filtered through 0.22-µm

Milliplex® syringe filter units (EMD Millipore Corp, MA, USA) to remove cellular debris. The conditioned medium was then aliquoted into 5 ml portions and kept at -20oC until protein content analysis. In addition, VFs cells were washed, collected and total RNA extracted.

2.6 Viability Studies

2.6.1 Fluorescein Diacetate- Propidium Iodide Assay

To assess the viability of VF cells following the stretch protocol, I employed the fluorescein diacetate-propidium iodide (FDA-PI) staining (Sigma-Aldrich, MO, USA). The non-fluorescent FDA molecules are taken up by live cells and converted to fluorescin, a green fluorescent dye, by cytoplasmic esterases. PI is membrane impermeable and generally excluded from viable cells while in cells with compromised (damaged) plasma membranes this 45 intercalating agent will easily penetrate into the cell and bind to DNA which will fluoresce red

(a sign of cell death). Individual VF cell lines derived from POP (n=3) and non-POP (n=3) patients were mechanically stretched for a maximum of 48 hrs at 25% elongation. After conditioned medium was collected, the stretched cells were washed twice in HBSS-/-, and then stained with FDA and PI (both at 20 µg/ml) for 3 min at RT. The flexible membranes of the

Flexcell plates were then carefully cut out with a blade and immediately mounted on to a glass slide (Fisher Scientific, ON, Canada), viewed under a fluorescent microscope using a 520 nm filter (Leica Microsystems) and photographed with Sony DXC-970 MD 3CCD color video camera (Sony).

2.6.2 Lactate Dehydrogenase Cytotoxicity Assay

The CytoTox 96 non-radioactive Cytotoxicity Assay (Promega, WI, USA) was used to quantitatively measure the concentration of Lactate Dehydrogenase (LDH) enzyme, which is released from the cells upon cell lysis/damage. The concentration of LDH in the culture supernatant is measured with the enzymatic assay that results in the conversion of a tetrazolium salt (INT) into a red formazan product. The amount of color is then quantified, which is proportional to the number of damaged cells. In a 96 well plate, 50 µl of substrate mix were added in triplicates to 50 µl of CM from non-stretched (NS-CM) and stretched (S-CM) VF cells

(n=3). The plate was covered and incubated in the dark at RT for 30 min. Next, 50 µl of stop solution was added to each well. The plate was gently rotated in the dark on an orbital shaker for 5 min. Absorbance was then recorded at 492 nm (uQuant).

2.7 Gene Expression Analysis

I examined the basal expression of multiple genes in non-stretched VF cell lines derived from premenopausal patients with severe POP and non-POP patients, and the effect of static mechanical stretch at 25% elongation for 24 hrs on mRNA levels. 46

For extraction of total RNA, VFs cells were washed twice with ice-cold HBSS-/-, scraped and lysed by adding 250 µl of TRIZOL (Gibco, Burlington, ON) to each well. Total

RNA was extracted according to the manufacturer`s protocol, RNA samples were column purified using RNeasy MiniElute Cleanup Kit (Qiagen, Mississauga, ON, Canada), and treated with 2.5µl DNase I (2.73 Kunitz unit/µl, Qiagen) for 15 min at RT to remove genomic DNA contamination. The RNA concentration was then measured by a NanoDrop 1000 spectrophotometer (Thermo Fisher Scientific Inc., DE, USA).

50 ng/ µl stock cDNA solutions were generated with iScript Reverse Transcription (RT) supermix (Bio-Rad Laboratories Inc., CA, USA) from 1 µg of RNA following the manufacturer`s recommended protocol. All PCR reactions were carried out on the CFX96 or

CFX384 Touch™ Real-Time PCR Detection Systems (Bio-Rad Laboratories Inc., CA, USA).

2.7.1 ECM and Adhesion Molecules Quantitative Profiler PCR arrays

I screened for the expression of 84 genes in VFs using the Human ECM and Adhesion

Molecules RT2 Profiler PCR array (SaBiosciences Corp., Frederick, MD, USA) according to the manufacturer’s instructions (Appendix, Table 2.1). cDNA was prepared from pooled samples of RNA extracted from 8 VFs cell cultures derived from POP (P) and 7 from non-POP

(C) patients exposed to static mechanical stretch at 25% elongation for 24 hrs (P-S and C-S respectively), and from their non-stretched counterparts (P-NS and C-NS respectively). cDNA was prepared from 1 µg total pooled RNA using the iScript Reverse Transcription (RT) supermix (Bio-Rad Laboratories Inc., CA, USA). PCR amplification was conducted with an initial 10-min step at 95°C followed by 40 cycles of 95°C for 15 s and 60°C for 1 min. The experiment was repeated three times. Data were imported into the integrated Web-based software package (http://pcrdataanalysis.sabiosciences.com/pcr/arrayanalysis.php; Qiagen

GmbH). Quantitative analysis was based on the ΔΔCt method, with normalization of the raw 47 data the geometric mean of 9 reference genes determined by the software as being the most stable within the samples (CD44, COL6A1, COL6A2, COL7A1, ITGA5, ITGB4, ITGB5,

THBS1, GAPDH). The Web portal automatically performs all ΔΔCt-based calculations from the uploaded raw threshold cycle (Ct) data.

2.7.2 Real Time Reverse Transcription Polymerase Chain Reaction (qRT-PCR)

To confirm the results of the RT2 Profiler PCR array, I conducted qRT-PCR on individual cDNA samples produced from VFs from 8 POP and 7 non-POP patients (stretch and non- stretch) using CFX384 Touch™ Real-Time PCR Detection Systems (BioRad). Specific sequences of oligonucleotide primers for LOX, LOXL-1-4, MMP-1, -2, -3, -8, -10, -12, -13, 14,

TIMP-1-4, ADAMTS-2 and BMP-1 were obtained from Gene Bank Database of the National

Centre for Biotechnology Information (NCBI) of the National Institutes of Health using Primer-

BLAST (see Table 2.2). Using pooled cDNA samples, I first tested primer efficiency and specificity, and created standard curves with 4-fold serial dilutions (from 50 ng/ µl to 0.195 ng/

µl). The efficiencies of the primers of interest were similar to those of the reference genes.

Using the geNorm algorithm, we selected the 3 most stable from the commonly used reference genes: YWHAZ, B2M, ACTΒ, TBP, HPRT1 and SDHA. Gene expression levels in all studies were normalized against the geometric mean of YWHAZ, TBP and ACTB. RT-PCR reactions were carried out in triplicate, where each reaction contained 10 ng of cDNA, LuminoCt®

SYBR® Green qPCR ReadyMix™ (Sigma), forward and reverse primers at a final concentration of 300 nM in a total of 10 µl per well reaction mix. The cycling protocol started with an initial denaturation 95oC for 30 sec, then 40 cycles of denaturation 95 oC for 5 sec and annealing/extension 60oC for 20 sec. We followed each qRT-PCR run by a melting curve analysis to confirm the specificity of the primers used; reaction was heated slowly from 65oC to

o o 95 C (0.1 C/sec). Gene expression values were analyzed using the ΔΔCq mode on the CFX

TM Manager software 2.0 (Bio-Rad Laboratories Inc., CA, USA). Sample replicates with a Cq

>35 were excluded from data analysis, and the cut-off quantification cycle standard deviation

(ΔCq) was set at ≤ 0.2. A no template control of each primer mastermix was also loaded on every plate to ensure the absence of contaminations.

Table 2. 1: Real-time PCR Primer Sequences of a Panel of Genes Studied and Reference Genes.

Target Genes Primer Sequences GenBank Accession # LOX Forward 5’-AGGCCACAAAGCAAGTTTCTG-3’ NM_002317 Reverse 5’-AACAGCCAGGACTCAATCCCT-3’ LOXL1 Forward 5’-CTGTGACTTCGGCAACCTCAA-3’ NM_005576 Reverse 5’-TGCACGTCGGTTATGTCGAT-3’ LOXL2 Forward 5’-TCGAGGTTGCAGAATCCGATT-3’ NM_002318 Reverse 5’-TTCCGTCTCTTCGCTGAAGGA-3’ LOXL3 Forward 5’-CGGATGTGAAGCCAGGAAACT-3’ NM_032603 Reverse 5’-AGGCATCACCAATGTGGCA-3’ LOXL4 Forward 5’-ACCGGCATGACATTGATTGC-3’ NM_032211 Reverse 5’-CATCATACTTGCAGCGGCACT-3’ MMP1 Forward 5’-TACGAATTTGCCGACAGAGATG-3’ NM_002421 Reverse 5’-GCCAAAGGAGCTGTAGATGTCC-3’ MMP2 Forward 5’-GAATACCATCGAGACCATGCG-3’ NM_004530 Reverse 5’-CGAGCAAAGGCATCATCCA-3’ MMP3 Forward 5’TGCTGTTTTTGAAGAATTTGGGTT-3’ NM_002422.3 Reverse 5’ACAATTAAGCCAGCTGTTACTCT-3’ MMP8 Forward 5’-GCCATCCCTTCCAACTGGTAT-3’ NM_002424.2 Reverse 5’-ATCATAGCCACTCAGAGCCCA-3’ MMP10 Forward 5’-TCATGCCTACCCACCTGGAC-3’ NM_002425.2 Reverse 5’-AATTGGTGCCTGATGCATCTTC-3’ MMP12 Forward 5’-AAGGCCGTAATGTTCCCCA-3’ NM_002426 Reverse 5’-CAGGATTTGGCAAGCGTTG-3’ MMP14 Forward 5’-TGCCATGCAGAAGTTTTACGG-3’ NM_004995 Reverse 5’-CCTTCGAACATTGGCCTTGAT-3’ TIMP1 Forward 5’-TTCTGGCATCCTGTTGTTGCT-3’ NM_003254 Reverse 5’-CCTGATGACGAGGTCGGAATT-3’ TIMP2 Forward 5’-GCGTTTTGCAATGCAGATGTAG-3’ NM_003255 Reverse 5’-TCTCAGGCCCTTTGAACATCTT-3’ TIMP3 Forward 5’-CTGCTGACAGGTCGCGTCTA-3’ NM_000362 Reverse 5’-GCTGGTCCCACCTCTCCAC-3’ TIMP4 Forward 5’-TCTGAACTGTGGCTGCCAAAT-3’ NM_003256 Reverse 5’-GCTTTCGTTCCAACAGCCAG-3’ ADAMTS2 Forward 5’-GTGTGCACCTGGCAAGCATTGTTT3’ NM_014244.4 Reverse 5’-GCCAAACGGACTCCAAGCGC-3’ BMP1 Forward 5’-GCCACATTCAATCGCCCAA-3’ NM_006132 Reverse 5’-TGGCGCTCAATCTCAAAGGAC-3’ ACTB Forward 5’ACCTTCAACACCCCAGCCATGTACG-3’ NM_001101 Reverse 5’CTGATCCACATCTGCTGGAAGGTGG-3’ TBP Forward 5’-TGCACAGGAGCCAAGAGTGAA-3’ NM_003194 Reverse 5’-CACATCACAGCTCCCCACCA-3’ YWHAZ Forward 5’-ACTTTTGGTACATTGTGGCTTCAA-3’ NM_003406 Reverse 5’-CCGCCAGGACAAACCAGTAT-3’

2.8 Quantitative Detection of Protein Expression in Conditioned Medium (CM)

CM was concentrated from 5 ml to a final volume of 500 µl per sample using the

Amicon® Ultra-4 Centrifugal Filter Units (UFC800308, Millipore, MA, USA) by spinning at

4000g for 15 min at 4oC in a swinging bucket rotor. The filter units contain a cellulose membrane with a molecular weight cut off of 3 kDa. Total protein concentration was determined using the Bovine Serum Albumin (BSA) Protein assay (Thermo Scientific Pierce,

DE, USA) using the manufacturer’s instructions. Concentrated CM was then diluted to a final concentration of 1µg/µl of total protein by SF-DMEM and stored in -80oC until use.

2.8.1 Quantibody Protein Array

The protein expression of 7 different Matrix Metalloproteinases (MMP-1, MMP-2,

MMP-3, MMP-8, MMP-9, MMP-10, MMP-13) and 3 Tissue Inhibitors of Metalloproteinases

(TIMP-1, TIMP-2, TIMP-4) were quantitatively measured using the Quantibody Human MMP

Array 1 (QAH-MMP1, Raybiotech, GA, USA) following manufacturer’s instructions (see

Figure 2.2). The array uses the multiplexed sandwich enzyme-linked immunosorbent assay

(ELISA)-based technology to accurately measure the concentration of multiple proteins simultaneously. The kit is supplied as a glass slide spotted with 16 identical antibody arrays; each array included positive controls in quadruplicates. After incubation with the sample, the target proteins are trapped onto the solid surface. Next, a second biotin-labeled detection antibody is added that recognizes a different isotope of the target protein. The protein-antibody- biotin complex is visualized by adding the streptavidin-labeled Cy3 dye and then reading it on a laser scanner. Quantification of concentrations of unknown samples is conducted by comparing their signals to array-specific standards whose concentrations have been predetermined.

Figure 2.2 Quantibody Array-Based Multiplex Sandwich ELISA System to simultaneously detect quantitatively the measurement of 7 MMPs and 3 TIMPs secreted in one sample of medium conditioned by stretched (S) and non-stretched (NS) vaginal fibroblasts derived from non-POP (C) and POP (P) patients. Shown here is location of the quadruplicate spot for each capture antibody.

The glass slide was first equilibrated to RT for 30 min, followed by air-drying at RT for

1-2 hrs. Seven standard serial dilutions were prepared as per manufacturer’s instructions from a supplied lyophilized standard protein mix. Slides were blocked for 30 min with 100 µl of sample diluent at RT, then the diluent was decanted and 100 µl of each standard protein mix or samples (CM samples collected from stretched and non-stretched VFs derived from POP (n=6) and from non-POP (n=6) patients at a concentration of 0.5 µg/µl) were added to the antibody array. The array was covered with an adhesive film and incubated overnight (ON) at 4oC.

The next day, the samples were decanted from each well, washed 5 times for 5 min with

150 µl of wash buffer at RT with gentle shaking, 80 µl of detection antibody cocktail were pipetted into each well and incubated at RT for 1-2 hrs. The samples were then decanted and washed 5 times. Another wash with 30 µl of wash buffer was carried out after the slides were dissembled at RT for 15 min with gentle shaking. A final wash with 30 µl of wash buffer II at

RT for 5 min concluded the washing steps. The slides were dried by centrifugation at 1000 rpm for 3 min without cap. The signals were visualized with a laser scanner equipped with a Cy3 wavelength (Tecan, Männedorf, Switzerland). Data extraction was done using ArrayVision microarray analysis software (Imaging Research Inc, St. Catharines, ON, Canada) and quantitative data analysis was carried out using the Quantibody Q-Analyzer Software purchased from the manufacturer.

2.8.2 Western Immunoblot Analysis

As there were no commercially available ELISA assay to detect the expression of the secreted LOX, LOXL-1-4, ADAMTS-2 and BMP-1 proteins by stretched (S) and non-stretched

(NS) VFs from POP (n=6) and non-POP (n=6) patients, we measured the expression of these proteins by Western Immunoblot .50 μg of CM were loaded on 10-12% SDS-polyacrylamide gel and protein samples were resolved by electrophoresis under reducing conditions at 100V for 53

2 hrs. The separated proteins were then transferred ON at 30V at 4oC onto polyvinylidene difluoride (PVDF) membrane (Millipore, Bedford, MA) in 25 mM Tris-HCl, 250 mM glycine,

0.1 % (wt/vol) SDS, pH 8.3 using an electrophoretic transfer cell (Bio-Rad). Membranes were then blocked with filtered 5% non-fat milk for 1 hour at RT, and incubated with the primary antibody (see Table 2.3) ON at 4oC. All antibodies were previously validated in our lab [35,

103, 139]. After incubation, the membrane was washed in TBS-T (Tris-buffered saline (TBS), with 0.05% tween), and incubated for one hour at RT with the required secondary antibody (see table 2.2). Proteins were detected using ECL detection system (Amersham Pharmacia Biotech) exposed to X-ray film (HyBlot CL, Denville, Metuchen, NJ) and analyzed by densitometry, using Image J software analysis (National Institutes of Health, USA). As no suitable reference protein was determined for conditioned medium, blots where stained with Coomassie Blue for

2 hrs at RT to determine total protein content. To verify equal protein loading, total protein content per sample and average protein content per patient group was calculated for each blot.

The values were expressed as a relative optical density (OD) of the protein of interest on the

Western blot to the total protein content as shown by Coomassie blue staining.

2.8.3 Zymography

Gelatinase zymography was used to detect the enzymatic activities of MMP-2 and

MMP-9. Total protein (6 µg /lane) was mixed with an equal volume of 2X non-reducing sample buffer to a volume of 9 µl/well, and CM samples from VFs were loaded on a gel (Novex 10%

Zymogram Gelatin Gel, Invitrogen, Carlspad, CA, USA). In addition to the CM samples, 10 µl of full range Kaleidoscope Precision Plus ProteinTM Prestained protein molecular weight marker, (BioRad, Hercules, CA, USA) were loaded. The gel was electrophoresed in Novex®

1X Tris-Glycine SDS Running Buffer (LC675, Invitrogen, CA, USA) in non-denaturing conditions at 90V for 3 hrs. After electrophoresis, the gels were washed twice for 30 min each

54 in 1X Novex® Renaturing buffer (LC2670, Invitrogen, CA, USA) at RT to remove SDS.

Zymograms were subsequently developed by incubation in Novex® Developing Buffer

(LC2671, Invitrogen, CA) for 18 h at 37ºC with gentle shaking. Incubated gels were then stained with SimplyBlue™ SafeStain (LC6060, Invitrogen, CA, USA). Proteolytic activities were detected as clear bands indicating the lysis of the substrate. The gels were scanned and quantification of band density was carried out using Image J software analysis (NIH, USA).

2.9 Statistical Analysis

Statistical analysis for continuous data was performed using ANOVA for gene expression, protein array and zymography and independent Student’s t-test for western blotting.

Independent Student’s –t-test was used for western blots since each blot contained only two groups (non-POP NS vs. POP NS, non-POP NS vs non-POP S, POP-NS vs. POP S). Fisher’s exact test was used for categorical data. Experimental error was reported as standard error of the mean (SEM). The statistical program Prism (version 4.0, Graph Pad Software Inc., San

Diego, CA) was used with the level of significance for comparison set at P<0.05. The number

(n) of mRNA samples from VF cell lines derived from POP and non-POP patients used in gene expression analysis were 8 and 7 respectively; for protein array, immunoblot and zymography analysis the number used was 6 protein samples from POP patients and 6 protein samples from non-POP patients. For the proliferation assay, I used 5 protein samples from POP patients and 5 samples from non-POP patients.

Table 2.2: Summary of Antibodies Used in Immunoblot Analysis

Antibody Primary Monoclonal Dilution Source Company or or Secondary Polyclonal LOX Primary Polyclonal 1:200 Rabbit Abcam, Cambridge, MA, USA LOXL1 Primary Polyclonal 1:200 Goat Santa Cruz Biotech, USA. LOXL2 Primary Polyclonal 1:200 Rabbit Abcam, Cambridge, MA, USA LOXL3 Primary Polyclonal 1:1000 Mouse Abnova, Taipei City, Taiwan LOXL4 Primary Polyclonal 1:1000 Mouse Abnova, Taipei City, Taiwan BMP1 Primary Polyclonal 1:1000 Rabbit Abcam, Cambridge, MA, USA ADAMTS-2 Primary Monoclonal 1:200 Mouse Abcam, Cambridge, MA, USA MMP-14 Primary Monoclonal 1:200 Mouse Abcam, Cambridge, MA, USA HRP-conjugated Secondary Polyclonal 1:3000 Goat GE anti-rabbit IgG Healthcare, Buckingham shire, UK HRP-conjugated Secondary Polyclonal 1:3000 Donkey Serotec, anti-sheep/ 3200, goat IgG Raleigh, NC, USA HRP-conjugated Secondary Polyclonal 1:4000 Rabbit GE anti-mouse IgG Healthcare, Buckingham shire, UK

CHAPTER 3: RESULTS

3.1 Patient Demographics

Vaginal vault biopsy samples were obtained from 15 Caucasian premenopausal women (8 in the POP group, and 7 in the non-POP group) matched for age and body mass index (BMI).

The mean age for the POP and non-POP patients was 43 and 44.85, respectively. The mean

BMI for POP and non-POP patients was 30.1 and 26.9, respectively. Furthermore, 88% of POP patients and 50% of non-POP patients reported symptoms of stress urinary incontinence. There was no statistical difference on those parameters. However, the mean parity was 3.1 and 1.3

(p=0.03) and the mean vaginal delivery was 3 and 1 (p=0.02) for POP and non-POP patient groups respectively. We also noted a statistical difference between POP and non-POP patients with respect to a family history of POP (p=0.019) (Table 3.1).

Table 3.1: Summary of Patients Demographics

POP Non-POP P value

n 8 7

Mean age 43 (37-50) 44.85 (42-49) NS

Mean BMI 30.1 (20.6-41.6) 26.88 (21.6-32.8) NS

Mean Parity 3.1 (1,6) 1.3 (0,3) 0.03

Mean Vaginal Deliveries 3 (1,3) 1 (0,1) 0.02

SUI 88% 57% NS

Family History of POP 63% 0% 0.019

3.2 Comparing the Biological Characteristics of the Vaginal Fibroblasts

3.2.1 Fibroblast Cell Identification by Indirect Immunofluorescence

I employed specific biomarkers of fibroblastic vs. myogenic origin via immunoflourescent staining to identify the phenotype of the VFs derived from 8 POP and 7 non-POP patients (Figure 3.1.A). My results show that at derivation (passage 0), the VFs were positive for anti-Vimentin (fibroblastic marker), as well as the smooth muscle markers anti-H-

Caldesmon and anti-alpha Smooth Muscle Actin. Thus, the cells in tissue were a mixture of fibroblasts and SMCs. However, by passage 2, 100% of cells were stained positive for anti-

Vimentin, although 5-6% stained still stained positive for the smooth muscle markers in both the POP and non-POP groups. To further confirm our results, I employed another smooth muscle biomarker in passage 2, anti-Desmin, as a specific marker of differentiation of cells into myofibroblasts in comparison to anti-H-caldesmon. Furthermore, I noted that 0% of cells stained for the anti-epithelial cell marker, cytokeratin, in passages 0-2 (Figure 3.1.B). I concluded that the majority of VF cells in culture were fibroblastic in origin, with some myofibroblastic presence and no epithelial contamination.

Figure 3.1: Immunofluorescence of Primary Vaginal Fibroblasts derived from premenopausal patients with POP. A) Representative images of VFs incubated with antibody against Vimentin (red), Smooth Muscle (SM) Actin, Desmin, H-Caldesmon and Cytokeratin (CK) (all green), and nuclear DAPI staining (blue) at passage 0, 1 and 2. Original magnifications: x400 B) Quantitative analysis of VFs from passage 2 derived from non-POP (black bars) and POP patients (grey bars) that expressed Vimentin, Desmin and SM-Actin respectively (percentage of positive cells versus all cells).

3.2.2 Cell Attachment on Different Extracellular Matrices

To determine whether POP (n=8) and non-POP (n=7) VFs derived from premenopausal patients preferentially attach to different ECM proteins, we used the commercially available

Human Cell Adhesion Array Kit. Our results indicate no differential attachment of VFs on

Collagen I, Collagen II, Fibronectin, Laminin, Tenascin, Vitronectin proteins except for a significant (p<0.05) decrease in attachment ability of VFs derive from POP patients on basement membrane protein Collagen IV in comparison to non-POP patients. However, these results were not expected, and we have concerns that the adhesion assay matrix protein’s concentration was too high to detect any differential attachment. Since we did not detect differential attachment on Collagen I, the most abundant collagen in the ECM, we selected this protein as an attachment substrate for the mechanical stretch studies (Figure 3.2).

Figure 3.2: Attachment of Vaginal Fibroblasts derived from premenopausal non-POP (black bars) and POP patients (white bars) on 7 different human ECM proteins: Collagen I (Col I), Collagen II (Col II), Collagen IV (Col IV), Fibronectin (FN), Laminin (LN), Tenascin (TN), Vitronectin (VN) and the control ECM protein, Bovine Serum Albumin (BSA). The results shown are the mean ± SEM of absorbance. A significant difference is indicated by ** (P<0.01).

3.2.3 Cell Proliferation on Different Extracellular Matrices

To examine the effect of different ECM substrates on the proliferative ability of VFs derived from non-POP and POP patients, I plated five primary cell lines from each group on

24-well Flexcell plates coated with collagen type I or proNectin, a synthetic analogue of fibronectin. MTT assay was carried out to assess a daily proliferation rate over a period of five days. My results indicate no difference in the proliferation rate of POP VFs plated on collagen I and proNectin in comparison to non-POP VFs. These results, in addition to the attachment assay data, confirm that collagen I could be used as an attachment substrate for the mechanical stretch studies (Figure 3.3).

Figure 3.3: Proliferation of Vaginal Fibroblasts derived from premenopausal non-POP (black line) and POP patients (grey dashed line) on (A) proNectin and (B) Collagen I coated flexcell plates respectively. The results shown are the mean ± S.E.M of wells per patient group.

3.3 Viability Studies: Mechanical Stretch Does Not Induce Cell Injury

To elucidate the viability of the VFs after application of static mechanical stretch for 6,

24 or 48 hrs, the viability of the mechanically stretched and non-stretched cells was determined by the PI/FDA vital staining and the LDH cytotoxicity assay. The majority of VFs (~ 95%) were viable, confirmed by green fluorescein intracellular staining and absence of the red staining, reflecting the ability of live cells to take up FDA and exclude PI. (Figure 3.4.A). Next, necrotic cell death was further quantified by the release of LDH from the VFs into the surrounding medium when exposed to static mechanical stretch at 25% elongation for a different time interval from 6 to 48 hrs (Figure 3.4.B). In non-stretched VFs, 9% of total LDH was released after 24 hrs in culture, compared to 15% in stretched VFs (p=0.06, NS). After 48 hrs, the relative cytotoxicity of non-stretch VFs versus stretched VFs was 22% and 23%, respectively (p=0.5). Thus, these results demonstrate that the application of static mechanical stretch to VFs does not induce significant cell damage. Based on these data the time interval of

24 hrs was chosen for further stretch experiments.

Control 24 25% Stretch 24 25% Stretch 48 hours hours hours B

Figure 3.4: Viability of Vaginal Fibroblasts derived from POP patients (n=3) exposed to static mechanical stretch at 25% elongation for different time intervals. A) Fluorescein Diacetate - Propidium Iodide Staining and B) LDH Cytotoxicity Assay. Black bars and white patterned bars represent mean ± S.E.M % cytotoxicity (damaged/dead cells) of non-stretched and stretched VF cells for the indicated time periods (0, 6, 24 and 48 hrs). The red dotted line indicates the maximized cell cytotoxicity after 48 hrs.

3.4 Gene Expression Profile

3.4.1 ECM and Adhesion Molecule Quantitative PCR Arrays

The PCR array detects the expression of 84 encoding genes involved in cell adhesion and communication and ECM remodeling, including ECM ligands and their trans-membrane receptors. In particular, I compared the gene expression of multiple collagen molecules (Figure

3.5A), ECM proteases and protease inhibitors (Figure 3.5.B), cell-cell adhesion molecules and basement membrane constituents (Figure 3.5.C), trans-membrane integrin receptors and cell- matrix adhesion molecules (Figure 3.5.D) in stretched (S) and non-stretched (NS) primary VFs derived from POP and non-POP patients.

My results demonstrate that in static culture, non-stretched POP VFs expressed significantly lower levels of COL14A1 and COL15A1 mRNA, and a significantly higher

COL7A1 levels as compared to non-stretched VFs derived from non-POP patients. Importantly, application of static stretch significantly down-regulated the expression of multiple collagens including COL1A1, COL5A1, COL6A1, COL4A2, COL5A2, COL11A1 and COL12A1 genes in

VFs derived from POP patients (Figure 3.5A). There was no significant effect of stretch on the expression of collagens in VFs derived from non-POP patients. Examining the differential expression of genes involved in cell-cell adhesion molecules and basement membrane constituents by non-stretched POP and non-POP VFs, I noted that SELE (Selectin-E), CDH1

(E-cadherin) and SELP (P-selectin) were significantly up-regulated, while CLEC3B

(tetranectin) and THBS2 (thrombospondin-2) were significantly down-regulated in POP VFs

(Figure 3.5.C). Similar to collagen molecules, stretch of non-POP VFs did not affect the expression of basement membrane or cell adhesion molecules. On the other hand, stretch significantly up-regulated the expression of TNC (tenascin-C), CTGF (connective tissue growth

69 factor), ECM1 (Extracellular matrix protein 1), SGCE (epsilon-sarcoglycan), SPP1

(Osteopontin), LAMC1 (laminin gamma-1), LAMA3 (laminin gamma-3), and significantly down-regulated the expression of CLEC3B (C-type lectin domain family 3, member B) and

THBS2 (Thrombospondin 2) in VFs derived from POP patients (Figure 3.5.C).

Furthermore, in non-stretched VFs derived from POP patients, I observed a significant up-regulation of ITGA1 (integrin alpha1), ITGA4, ITGA8, ITGB1, ITGB2, ITGB3, NCAM1

(Neural cell adhesion molecule 1) and CTNNA1 (catenin alpha-1) mRNA, and a significant down-regulation in ICAM1 (Intercellular adhesion molecule 1) and VCAM1 (Vascular cell adhesion molecule 1) gene expression as compared to non-stretched non-POP VFs (Figure

3.5D). Static mechanical stretch further up-regulated the expression of ITGA4, ITGB1 and

CTNNA1 genes in POP in addition to ITGA6, ITGA2, and CTNNB1 genes, and further down- regulated the expression of VCAM1 as well as ITGA8 and CNTN1 (contactin 1) genes (p<0.05).

In VFs derived from non-POP patients, mechanical stretch down-regulated the expression of

ITGAV, ITGA1, ITGB1, and significantly up-regulated the expression of ITGA4 (Figure 3.5.D).

Next, I analyzed the basal expression of ECM remodeling genes in non-stretched VFs isolated from POP and non-POP patients. I found that under control static conditions, in POP patients when compared to non-POP patients, MMP-2 and TIMP-3 were significantly (p<0.05) down-regulated, whereas MMP-10, MMP-13 and ADAMTS13 were significantly (p<0.05) up- regulated (Figure 3.5B and 3.6). Importantly, mechanical stretch significantly down-regulated the transcript levels of ADAMTS8, ADAMTS13 and TIMP-2 (p<0.05 for all), but up-regulated the expression of MMP-1, MMP-3 and MMP-10 genes (p<0.05) in VFs from POP patients.

Furthermore, mechanical stretch significantly down-regulated MMP-2, MMP-8 and MMP-13 genes in non-POP VFs (p<0.05) (Figure 3.5.B and Figure 3.6).

Figure 3.5: Gene Expression Heat Map from PCR-array analysis of genes of which expression levels differ among the indicated cell populations involved in A) Collagens & ECM Structural Constituents; B) ECM proteases and protease inhibitors; C) Cell-Cell Adhesion Molecules and Basement Membrane Constituents and D) Trans-membrane and Cell-Matrix Adhesion Molecules. Pooled RNA samples of stretched (S) and non- stretched (NS) non-POP (n=7) and POP VFs (n=8) were used. Genes highlighted in red are statistically significant (P<0.05).

Figure 3.6: Relative Expression of ADAMTS, MMPs and TIMPs transcripts from pooled RNA samples of stretched (S) and non-stretched (NS) VFs derived from non-POP (n=7) and POP VFs (n=8), which expression levels differ among the indicated cell populations as determined using an RT-PCR-based mRNA expression array. The results shown are the mean ± S.E.M relative to non-stretched non-POP patients. A significant difference is indicated by * (P<0.05). ** (P<0.01).

3.4.2 Real Time Reverse Transcription Polymerase Chain Reaction (qRT PCR) Analysis

The results of the PCR array experiment for ECM proteases and protease inhibitors

(Figure 3.5.B) were chosen for verification by quantitative RT-PCR using individual RNA samples isolated from VFs derived POP (n=8) and non-POP patients (n=7). qRT-PCR results confirmed that basal expression of MMP-2, MMP-14, TIMP-2 and TIMP-3 genes were significantly lower, whereas MMP-3 expression was significantly higher in cells isolated from

POP patients in comparison to non-POP VFs (p<0.05). We further confirmed that mechanical stretch significantly (p<0.05) up-regulated MMP-1 and TIMP-3 mRNA levels in POP patients

(Figure 3.7).

A previous in vivo study indicated differential mRNA expression of proteins involved in the processing and maturation of collagen and elastin polymers, specifically LOX, LOXL1-4

[139], BMP [35] and ADAMTS2 [103], in vaginal biopsy samples from POP and non-POP patients. Unfortunately, these genes were not included in the PCR array. Hence, to understand whether these in vivo changes mirrored the differences in cellular expressions, I examined the mRNA levels of these genes in vitro in the static VFs cultures derived from POP and non-POP patients. Similar to earlier in vivo data, a significant decrease in LOX, LOXL1-3, ADAMTS2 and

BMP1 mRNA levels were noted in vitro in cultured POP VFs in comparison to non-POP VFs.

Importantly, mechanical stretch up-regulated the expression of LOXL4 in non-POP VFs

(p<0.05), and further decreased LOXL3 and LOXL4 expression in POP VFs (p<0.05) (Figure

3.7).

Figure 3.7: The Expression Level of A) LOXs B) MMPs C) ADAMTS2 and BMP-1 and D) TIMPs Genes Determined by qRT-PCR in VFs from non-POP (n=7, black bars - non- stretched, white dotted bars - stretched) and POP patients (n=8, grey bars - non-stretched, grey dotted bars - stretched) normalized to three reference genes. The results shown are the mean ± S.E.M relative to non-stretched non-POP patients. A significant difference is indicated by * (P<0.05), ** (p<0.01), *** (p<0.001).

3.5 Quantitative Detection of Protein Expression in Conditioned Media

3.5.1 Quanti-body Protein Array

To confirm that the transcript levels of ECM proteases and protease inhibitors correspond to their protein expression levels, I applied antibody array technology to determine the profile of 7 proteases and 3 protease inhibitors (namely MMP-1, -2, -3, -8, -9, -10, -13 and

TIMP-1, -2, -4) in culture media conditioned by VFs derived from non-POP (n=6) and POP patients (n=6). I did not detect significant changes in the secretion of these proteins between the two patient groups under static conditions. However, significant up-regulation in secreted

MMP-1 (p=0.0074), MMP-8 (p=0.02) and MMP-9 (p=0.02) by stretched POP VFs was noted.

Furthermore, TIMP-4 secretion was significantly up-regulated by stretch in from both POP and non-POP VFs (p<0.01 for both); whereas, TIMP-2 protein expression was down- regulated by stretch in POP patients (p=0.02) (Figure 3.8).

Figure 3.8: Quanti-body Protein Array Analysis of A) MMPs and B) TIMPs secreted by VFs derived from non-POP (n=6, black bars: non-stretched (NS), white dotted bars – stretched (S) VFs) and POP patients (n=6, grey bars- non-stretched, grey dotted bars - stretched VFs). The bars represent the mean ± SEM relative to non-POP NS patients. A significant difference is indicated by *(p<0.05),** (p<0.01).

3.5.2 Western Immunoblot Analysis

Western Blot analysis of media conditioned by VFs derived from POP and non-POP patients (n=6/group) was performed using antibodies that specifically recognize proteins that showed a significant difference at the transcript level (LOX, LOXL1-4, ADAMTS2 and BMP-

1). LOX, LOXL1-2, ADAMTS2, BMP-1 proteins were detected in non-stretched and stretched

CM samples, however, LOXL3 and LOXL4 proteins were not detected. To confirm that VFs do not secrete detectable levels of LOXL3 or LOXL4, I used tissue lysates from vaginal tissue biopsies as positive controls (Figure 3.9). To quantitate the expression of selected proteins in the CM, I calculated a relative optical density (OD) of the protein of interest to the total protein content detected by Coomassie blue staining (Figure 3.10).

LOX was detected as a 37 kDa (active form) protein, whereas LOXL1 was noted as two bands, 91 kDa (pro-form) and 35 kDa (active form). LOXL2 was detected as 85 kDa (pro-form) and 63 kDa (active form). There was no difference in the expression of LOX, LOXL1 and

LOXL2 between VFs derived from POP and non-POP patients under non-stretched conditions.

When exposed to static mechanical loading, however, VFs reacted differently. Stretch did not change the secretion of LOX in non-POP VFs but induced significantly smaller amounts of both secreted forms of LOXL1 and pro-LOXL2 in the conditioned medium (p<0.05 for all).

POP VFs exposed to stretch showed a decrease in the secretion of active form of LOX (37 kDa, p=0.0013) in conditioned medium (Figure 3.11). The 63 kDa active form was barely detectable in the CM from non-stretched cells.

ADAMTS2 protein was detected as a double band: an intermediate 75 kDa form and 58 kDa, however there was no difference in expression of both isoforms of ADAMTS2 between all study groups. (Figure 3.12).

BMP1 secreted by VFs derived from non-POP and POP patients showed a pattern of 4

78 bands with molecular weights that correspond to different isoforms derived from the same gene

(130 kDa, 92.5 kDa, 82.5 kDa, and 70 kDa). The expression of the 92 kDa, 82.5 kDa and 70 kDa bands was significantly down-regulated in the CM of stretched non-POP VFs (p<0.05)

(Figure 3.13).

Figure 3.9: Western Immunoblot Analysis of LOXL3 and LOXL4 in Vaginal Tissue (Tissue) and in Conditioned Medium (CM) of VFs derived from POP (PT5) and non-POP patients (CT7). The bands represent proteins with the following molecular weights: LOXL3 (37 kDa), LOXL4 (90kDa), showing the secretion of these proteins in tissue, but not in conditioned media samples of VFs derived from POP and non-POP patients.

B Average of Total Protein Ratio of Averages SEM of Average Values

POP NS 14198.1 0.94 2268.6 Vs Non-POP NS 14949.5 2668.7 Non POP S 3044.5 1.05 1684.5 Vs. Non-POP NS 2897.4 1741.7 POP S 110146.4 1.16 22898.69 Vs. POP NS 94340.1 19776.9

Figure 3.10: Coomassie Blue Staining of A) Duplicate PVDF membranes representing total protein content in CM from stretched (S) and non-stretched (NS) VFs derived from non-POP and POP patients B) Total protein values were used in Western Immunoblot analysis (Figures 3.9, 3.11, 3.12, 3.13) to normalize for slight variances in loading. Shown here are the average of the total protein values, the ratio and the SEM.

Figure 3.11 Western Immunoblot Analysis of LOX, LOXL1-2 in Conditioned Medium (CM) of VF derived from non-POP (n=6, black bars - non-stretched, white dotted bars - stretched) and POP patients (n=6, grey bars - non-stretched, grey dotted bars - stretched). Representative (A) Western Blot and (B) densitometric analysis. The bands represent proteins with the following molecular weights: LOXL1 (91-35kDa), LOXL2 (85-63kDa). The bars represent the mean ± S.E.M. A significant difference is indicated by *(p<0.05).

Figure 3.12: Western Immunoblot Analysis of ADAMTS2 in Conditioned Medium of VF derived from non-POP (black bars - non-stretched, white dotted bars - stretched) and POP patients (grey bars - non-stretched, grey dotted bars - stretched). (A) Representative Western Blot and (B) densitometric analysis. The bands represent proteins with the following molecular weights: ADAMTS-2 intermediate (75kDa) and pro-form of isoform-2 (58kDa) .The bars represent the mean ± S.E.M (n=6/group).

** * *

Figure 3.13: Western Immunoblot Analysis of BMP-1 in Conditioned Medium of VFs derived from non-POP (black bars - non-stretched, white dotted bars - stretched) and POP patients (grey bars - non-stretched, grey dotted bars - stretched). (A) Representative Western Blot and (B) densitometric analysis. The bars represent the mean ± S.E.M. (n=6/group). A significant difference is indicated by * (p<0.05) and ***(p<0.01).

3.6 Zymography

Enzymatic activity of proteins secreted by VFs from POP and non-POP patients was analyzed in conditioned culture media (n=6/group). Zymography detected gelatinase activities corresponding to latent pro-form and active form for both MMP-2 and MMP-9. MMP-2 activity is observed as a prominent band at 68 kDa (latent pro-form) and 62 kDa (active form)

(Figure 3.14.A). Densitometric analysis indicated very low levels of active-MMP-2 in CM from non-POP VFs, irrespective of mechanical loading; and static mechanical stretch significantly induced active MMP-2 secreted by stretched POP-VFs (p<0.05, Figure 3.14.B). Importantly,

MMP-9 activity was detected at 92 kDa (latent pro-form) and 87 kDa (active form), however, this was much lower than MMP-2 activity and was not statistically different between patient groups.

Figure 3.14: Representative Gelatin Zymography (A) and Densitometric analysis (B) showing gelatinase activity (visualized as light bands) of total protein in conditioned medium of VFs derived from non-POP (black bars - non-stretched (NS), white dotted bars – stretched (S) and POP patients (grey bars - non-stretched, grey dotted bars - stretched). MMP-2 activity was observed predominantly at 68 kDa (latent pro-form) and 62 kDa (active form). MMP-9 activity was observed at 92 kDa (latent pro-form), and 87 KDa (active form). B) Densitometric analysis of the gelatin zymography; bars represent the relative density of MMP-2. Non-POP NS sample #5 was chosen as a calibrator for the two zymograms. Bars represent the mean ± S.E.M. (n=5- 6/group). A significant difference is indicated by * (p<0.05) and **(p<0.01).

CHAPTER 4: DISCUSSION

4.1 Overall Summary

The weakening of the pelvic floor and extrusion of the pelvic organs during POP is associated with serious inconveniences, significantly disrupting the quality of life of the affected women. While conservative therapies for POP are available, they are not always sufficient or effective. Furthermore, reconstructive surgery for POP is associated with a high failure rate due to the already weakened native tissue [5]. In the 1970’s, urogynecologists began using vaginal meshes to reinforce tissue during POP surgery. However, this approach was associated with multiple and severe complications. At the same time, with the increased life expectancy and activity levels of modern women, the need for effective long-lasting repair therapies for POP has grown. Hence, cell-based solutions could represent a novel alternative to minimize the negative effects of mesh application.

Recently, there has been a growing interest in characterizing the tissue composition of patients with POP. Samples were obtained from the vaginal wall [35-39, 103, 139-141, 151], pubocervical fascia [34] and uterosacral ligaments [32, 33]. Conflicting results have been reported, mainly due to lack of homogeneity in tissue sample sites and in patient populations with respect to age, parity and menopausal status, however, the overall consensus is that connective tissue of women with POP is weakened due to an imbalance in its remodeling [35,

38, 39, 41, 103, 139]. In normal tissue, remodeling is a rigorously balanced process, and the mechano-sensitive fibroblasts play a central role in maintaining ECM homeostasis. These cells produce ECM proteins, and their modulators, namely LOXs, ADAMTS, BMP1, MMPs and

TIMPs, to control the anabolic and catabolic processes that remodel collagen and elastin fibers and the surrounding matrix. It has been shown that the expression of MMP-1 [103, 125], MMP-

2 [30, 103, 151], MMP-9 [103] and MMP-12 [103] is increased in tissue from patients with

POP in comparison to non-POP patients. Concurrently, the expression of TIMP1-4 gene [103,

125] and TIMP1 protein [103] was decreased in prolapsed pelvic tissue of POP patients.

Furthermore, LOX [139] family gene and protein expression was significantly decreased, in addition to differential expression of BMP-1 [35] and ADAMTS-2 [103] genes and proteins in the anterior vaginal wall tissue of POP patients in comparison to non-POP patients.

Collectively, these in-vivo results demonstrate that the biomechanical environment of prolapsed tissue is altered, resulting in loss of tissue strength and affecting the quality of the pelvic floor support.

Little is known about the active interactions between the cell and the ECM, and how this affects the pathophysiology of POP. Consequently, researchers have now turned their attention towards studying characteristics of pelvic floor cells (mainly fibroblasts) and how they respond to mechanical loading. It was recently reported in a minimal number of samples (n=1) that fibroblasts derived from vaginal tissue of women with mild and severe POP displayed decreased expression of COL1A1 and COL3A1, delayed cell alignment and rearrangement of F- actin cytoskeleton in comparison to fibroblasts derived from a non-POP patients [160]. When

POP VFs were exposed to mechanical loading, expression of COL1A1, COL3A1 and MMP-2 was down-regulated, while the collective activity of MMP-1, -8 and -13 was up-regulated [43].

Still, the question remains as to whether there are inherited differences in the biological characteristics, attachment to different ECM substrates, proliferative capacity and ability to produce ECM ligand proteins and their receptors (integrins) between VFs derived from POP and non-POP patients. In addition, I questioned whether the changes in ECM gene and protein expression previously detected in prolapsed vaginal tissue in vivo, LOXs [139], BMP1 [35],

ADAMTS2, MMPs and TIMPs [103], were due to intrinsic defects of POP vaginal cells, or as a consequence of exposure to increased intra-abdominal pressure. To address this, I examined the

89 expression of these proteins in vaginal cells derived from severely prolapsed tissue biopsies and from the vaginal tissue collected from non-POP patients. The experiments were performed under normal cell culture conditions and when VFs were exposed to static mechanical loading for different intervals at 25% elongation. This study was designed to assess in vitro the dynamic cell-ECM interactions that are important for understanding the pathophysiology of POP.

Firstly, I examined the biological characteristic of cells derived from POP patients and compared them to cells derived from non-POP patients. To control for variability within patient populations, I collected samples from POP and non-POP patients that were matched for age,

BMI and hormonal status (all premenopausal). My demographical data supports the results of other studies, which showed that the risk of developing POP escalates with an increased mean parity [9-12, 16] as well as with a family history of POP [20, 21].

4.2 Biological characteristics of VFs and their Ability to Produce ECM Proteins

Recent clinical studies have focused on the regenerative capacity of vaginal connective tissue through the application of cell-based tissue engineering. Understanding of the cell-matrix interaction in tissue of women with POP is necessary to cultivate new therapeutic approaches, especially in the development of new scaffolds. Similar to other connective tissues in the human body, vaginal connective tissue consists of several constituents, both cellular and extracellular, but its major ECM structural protein is collagen I, and the predominant cell type is the fibroblast. The fibroblast serves in the maintenance and remodeling of the ECM; however, little information is known regarding the regulation of proliferation, differentiation and activity of these cells during normal vaginal function. Even less is known about the alterations of these cells in response to different pathological environmental stimuli, chemical and/or mechanical. There is evidence that the vaginal connective tissue from patients with POP is phenotypically different than the connective tissue derived from non-POP patients, showing

90 diminished collagen I and III levels [122, 123], and an ECM that is disordered, less dense and loosely arranged [127]. Kerkhof et al [175] compared vaginal wall tissue samples from non-

POP and POP sites from the same patient, and found that there was a tendency towards an increase in the collagen III and elastin content, and a significantly increased number of SMCs and collagen cross-links at the POP site.

My current data indicate that human vaginal cells derived from POP and non-POP patients expressed the fibroblastic marker vimentin and smooth muscle markers, alpha smooth muscle actin and desmin, in similar proportions. Thus, both cultures were essentially fibroblastic in origin, with equal myofibroblastic contributions. Several reasons for why both

POP-VF’s and non-POP VF’s cultures de-differentiate from a culture of myofibroblasts in passage 0 to fibroblasts by passage 2 include the increased proliferation rate fibroblasts in comparison to smooth muscle cells and possibly the high concentration of FBS in the cell culture media. Both cell populations showed a similarity in the attachment to different ECM components, except for collagen IV. Again these results are not typical, and possibly due to the saturated concentration of matrix proteins used to coat the wells in this commercial attachment assay. Type IV collagen is present ubiquitously, but only in basement membranes. It provides a scaffold for cellular assembly and mechanical stability, and plays an important role in cell adhesion, growth, migration, proliferation and differentiation. It is also the main ECM protein secreted from fibroblasts following trauma, and induces scar formation [176]. Attachment of cells to collagen IV is mediated via multiple binding sites, and involves several integrin and non-integrin receptors [177]. Therefore, decreased attachment of POP-VFs to collagen IV may contribute to the pathological wound healing environment of the weakened pelvic floor in patients with POP.

The RGD attachment domain, found in plasma fibronectin and its synthetic counterpart

91 proNectin, serves as the cell attachment site of many proteins including blood, cell surface and extracellular matrix proteins. Integrins function as protein receptors for these proteins, and together they form a major recognition system for the adhesion of cells to the ECM [108].

Cellular proliferation of several cell types, including bronchial epithelial cells [178] and embryonic mesenchymal cells [179] is promoted by adhesion to surfaces containing the RGD sequence. Because of the importance of this sequence in cellular interactions, I chose to examine if it affected the proliferative ability of VFs derived from non-POP and POP patients.

My results indicate no difference in the proliferation rate of POP VFs plated on collagen I and proNectin in comparison to non-POP VFs. Collagen I also contains an RGD sequence, which is only exposed once collagen is in its non-helical, denatured form. In normal wound healing, this allows fibronectin to bind denatured collagen with a high affinity, concentrating fibronectin at the sites of tissue injury, and promoting wound matrix strengthening [180]. Since Collagen I is the most abundant protein in connective tissue, and because the attachment and proliferation of

VFs derived from POP patients in comparison to non-POP VFs on Collagen I were not different, I choose this substrate for the experiments using static mechanical stretch. Collagen and its degradation products are involved in many other cellular processes that are essential to wound healing. Collagen fibrils signal to platelets to release clotting factors [181], and collagen fragments also recruit other cell types, including fibroblasts, to promote the formation of granulation tissue and epithelization.

In normal, intact tissue, the collagen matrix undertakes the majority of the tension, and thus the fibroblasts are under relatively low stress. Once wounded, the adult skin undergoes a complex process involving the cooperative effort of many different cell types. Two predominant phenomena are involved in the process of normal wound healing: re- epithelialization and the formation of granulation tissue. During re-epithelialization, epidermal

92 cells replicate and deposit along the wound. Granulation tissue consists of fibroblasts, myofibroblasts, inflammatory cells and small vessels, and is formed when a fibrin clot is deposited in a full-thickness dermal wound, releasing growth factors from local intact dermis that stimulate the fibroblasts to invade the wound ECM [182]. Migration of fibroblasts exerts tractional forces on the collagen surface, and organizes the cells along the stress lines. This stress stimulates the fibroblasts to differentiate into myofibroblasts, which in turn produce collagen, other ECM molecules and proteases. Myofibroblasts also generate an increased contractile force that shortens the collagen matrix and results in wound closure [183]. Once re- epithelialization is complete, apoptosis ensues to decrease the cellularity of the granulation tissue, resulting in the formation of a poorly cellularized scar [184]. In pathological wound healing, due to mechanisms not yet well understood, myofibroblasts persist and continue to remodel the ECM, and granulation tissue does not undergo apoptosis. This results in a fibrotic, or hypertrophied scar, containing excess myofibroblasts with an imbalance in the surrounding

ECM. Consequently, the surrounding tissue becomes deformed and malfunctioned [183]. In

POP, the pathological tissue environment may be a result of impaired myofibroblast physiology, or compromised interaction of myofibroblasts with growth factors and ECM receptors, including those for collagen and fibronectin. Thus, when pelvic floor tissue undergoes acute injury during vaginal delivery, or chronic injury due to an increase in intra- abdominal pressure, defective wound healing can further weaken the pelvic connective tissue, and cause the tissue prolapse.

Our previous studies demonstrated impairment in the expression and activity of ECM- homeostasis related factors (multiple enzymes regulating ECM assembly, synthesis and degradation) in vaginal tissue biopsies derived from POP patients in comparison to non-POP patients in vivo [35, 103, 139]. However, the correlation of these in vivo changes in collagen

93 and elastin metabolism in women with POP required further exploration using an in vitro approach. To date there are limited investigations on the metabolism of collagen and elastin in

VFs. Makinen et al [185] found that the rate of collagen synthesis and pro-collagen mRNA levels in fibroblasts derived from POP patients were similar or slightly higher than those from age-matched non-POP patients. We also reported key differences in collagen and integrin mRNA expression between POP VFs and non-POP VFs. Under normal culture conditions, POP

VFs expressed lower mRNA levels of collagens that co-distribute with collagen I (COL14A1 and COL15A1) in comparison to non-POP VFs. Research groups have traditionally used immunohistochemical and Western immunoblot approaches to assess collagen content in pelvic floor tissue of women with and without POP [126, 130, 131, 186]. While conflicting results have been reported, the conclusion is that overall collagen content is decreased in pelvic floor tissue of women with POP in comparison to non-POP patients.

I also noted differential expression of cell-cell adhesion molecules and basement membrane proteins between POP VFs and non-POP VFs. My results indicate that POP VFs expressed higher mRNA levels of several alpha and beta integrins, which may implicate altered cell-matrix interactions and potential differences in mechano-transduction mechanism in POP

VFs. The endothelial and platelet selectins (Selectin-E and Selectin-P, respectively) were also up-regulated in POP VFs. Selectins play a significant role in inflammatory processes, and their up-regulation may indicate an increase in cellular interaction with inflammatory mediators, contributing to the pathological wound-healing environment in POP pelvic floor tissue [121].

However, I detected lower transcript levels of intercellular and cell adhesion molecules, ICAM1 and VCAM1 respectively, in fibroblasts derived from POP patients in comparison to non-POP

VFs. Altogether these results confirm that fibroblasts derived from POP patients do not respond to changes in their environment in the same way as non-POP VFs. When exposed to pathologic

94 environmental stimuli, POP VFs might not be able to restore ECM homeostasis, resulting in loss of tissue strength and subsequent tissue damage.

In addition, I demonstrated that under non-stretch conditions, cultured VFs from POP patients expressed lower transcript levels of BMP1 and the LOX family members, which was similar to the results of our earlier in vivo studies [35, 139]. However, I did not detect significant changes in the levels of secreted BMP1, LOX, LOXL1 and LOXL2 by POP VFs.

Frequently, LOX shows a similar expression pattern with collagen type III, and the active LOX protein stimulates the collagen 3A1 promoter [68]. In pathological wound healing, LOX production is under the regulation of growth factors and cytokines, thus LOX may also interact with some cytokines and other ECM constituents. Furthermore, LOX expression in fibroblasts corresponds with their differentiation to active myofibroblasts, which plays a role in fibrotic disorders [187]. Although the key known function of LOX is to contribute to collagen and elastin fiber crosslinking, all of these features indicate that the LOX proteins may have roles over and above this known function. Hence, while I did not detect changes in the secreted LOX and LOXL1-2 protein levels between POP and non-POP VFs, I speculate that intracellular levels of the LOX proteins may be significantly different. In contrast to our in vivo results, I detected in vitro a decrease in the expression of PCP and PNP in POP VFs in comparison to non-POP VFs. It has been shown before that mechanical tension can decrease transcript levels of LOX, pro-collagen I and III, PNP and PCP [188]. Thus, it is possible that VFs derived from severely prolapsed stretched tissues would express lower levels of LOX, PNP and PCP.

I previously reported a reduction in TIMP-2 and TIMP-3 mRNA in POP vaginal biopsies in comparison to non-POP patients [103]. My current results showed that cultured VFs derived from POP patients expressed lower levels of MMP-2, TIMP-2 and TIMP-3 mRNA, and a decrease in pro-MMP-2 activity in comparison to VFs derived from non-POP patients. During

95 wound healing, fibroblasts differentiate from a quiescent to a migratory myofibroblastic state.

Recently, Howard et al [189] showed that the myofibroblast phenotype suppresses expression of MMP-2, which was inversely related to the expression of the contractile protein, α-smooth muscle actin. Furthermore, they reported that factors that increased myofibroblast contractility, such as TGF-β and serum, decreased MMP-2 expression. In pathological wound healing environments, such as that of the pelvic floor of POP patients, the myofibroblast phenotype persists. This could explain why MMP-2 expression was lower in POP-VFs in comparison to non-POP VFs when exposed to mechanical stretch.

MMP-2 has received special attention due to its ability to digest collagen I. Pro-MMP-2 is cleaved to active MMP-2 by MMP-14/MT1-MMP in a tri-molecular complex involving

TIMP-2 [190]. This MMP-14 mediated activation is up-regulated by certain ECM molecules, including collagen I, in fibroblasts in order to regulate collagen I synthesis and prevent fibrosis

[191]. MMP-2 co-localizes with ITGB3 on the surface of many cells, including angiogenic blood cells and melanoma [192] , and VFs derived from POP patients demonstrated an increased expression of ITGB3 transcripts. ITGB3 selectively suppresses the collagen I induced

MMP-2 activation [193]. Thus, overexpression of ITGB3 on the surface of VFs may serve as a protective mechanism to decrease the activity of MMP-2 in POP patients. The result of these studies confirm the differential expression of the ECM degrading proteins and their inhibitors in cells derived from severe POP patients vs. non-POP VFs, confirming genotypical and phenotypical differences in these fibroblasts, which potentially can predispose them to different modes of reaction with respect to remodeling of the ECM.

A difference in expression of MMP-1, -3, -8, -10 and -13, by POP and non-POP VFs was not found, which was in accordance with our recently published in vivo data [103]. Based

96 on these findings, we speculate that the differential expression of these MMPs is under the control of exogeneous factors such as ovarian hormones and/or mechanical stretch.

In conclusion, there is a distorted ECM biogenesis by VFs derived from POP patients compared to VFs derived from non-POP patients. POP-VF’s express lower levels of MMPs,

TIMPs, LOXs, BMP1 and ADAMTS2. Thus, there is a decreased turnover of ECM in POP

VF’s in comparison to NON-POP VF’s (Figure 4.1).

4.3 Mechano-responses of Primary Human Fibroblasts Derived from Non-Prolapsed

and Prolapsed Vaginal Tissue

Previous studies have shown that an increase in intra-abdominal pressure is transmitted directly to the pelvic floor and vagina [194] , and the outcome of this increased mechanical load is tissue stretch [195] and remodeling by inducing MMPs expression and activation [196, 197] via a process known as mechano-transduction. The vagina is clearly a load-bearing tissue, and although mechano-transduction has been confirmed in many other load-bearing tissues [183,

184], very little is known about the impact of increased mechanical load and tissue stretch on the behavior of VFs.

To investigate the cellular mechanisms through which stretch modulates ECM metabolism in the pelvic floor tissue, I compared the effect of static mechanical load on the expression of enzymes involved in collagen and elastin synthesis in VFs derived from POP and non-POP patients. My present data demonstrates a clear difference in mechano-responses of these VFs. In particular, I found that when static mechanical stretch was applied, VFs derived from POP patients massively down-regulated the mRNA expression of the fibril forming collagens, COL1A1, COL5A1, COL5A2, COL6A1 and COL11A1, as well as COL12A1 which co-distributes with collagen I and the basement membrane collagen, COL7A1. Ruiz-Zapata et al [160] similarly reported a decrease in COL1A1 mRNA expression in POP fibroblasts when 97 exposed to cyclic mechanical stretch. These changes could explain that the increased distensibility [146, 158] of vaginal tissue of women with POP in comparison non-POP patients is due to the decreased collagen I: collagen III ratio. Stretch also down-regulated several other

ECM proteins, including Laminin ɣ1 and ɣ3 in POP-VFs. Laminins contribute to cell adhesion, proliferation, migration and differentiation, and are crucial to proper basement membrane assembly [112]. Their down-regulation affects fundamental biological characteristics of tissue fibroblasts that are essential to proper function and production of ECM proteins. Importantly, static mechanical stretch did not affect transcript levels of collagens, basement membrane proteins or cell adhesion molecules in non-POP VFs. The expression of α and β integrins in stretched non-POP-VFs was also different from POP-VFs. This clearly demonstrates the altered mechano-responses of POP-VFs as compared to non-POP VFs derived from women with healthy pelvic floor.

I also examined the effect of mechanical stretch on the activity of collagen-degrading enzymes and their endogenous inhibitors. In VFs derived from POP patients, mechanical stretch decreased TIMP-2 expression, and upregulated the expression of collagenases MMP-1 and MMP-8, the gelatinase MMP-9, and the enzyme activity of active-MMP-2. On the other hand, in non-POP VFs, mechanical stretch decreased levels of pro-MMP-2 and increased the expression of TIMP-4. Zong et al [43] similarly reported that mechanical stretch increased the collective collagenase activity in VFs derived from POP patients. Mechanical stretch also increased the expression of MMP-2 in fibroblasts derived from fetal lung tissue [198], and anterior cruciate ligaments (ACL) and medial collateral ligaments (MCL) [199]. As mentioned above, collagenases degrade the major constituents of the ECM, including collagen I, II and III, gelatin, and aggrecan [200]. Hence, a net and augmented degradative activity due to mechanical stretch explains why the risk factors that result in increased intra-abdominal pressure, such as

98 heavy lifting, chronic cough and constipation, are associated with the progression of POP [194].

The expression of TIMP-2 (which directly binds to MMP-2) is ten times higher in the vaginal tissues biopsies than the expression of TIMP-1 and TIMP-3 proteins [103], and thus can be regarded as the major inhibitor of metalloprotienases in VFs. Collectively, these results clearly demonstrate that mechanical stretch on VFs derived from previously injured vaginal tissue induces collagenase activity that is not counteracted by their endogenous inhibitors. This results in weakening of their supportive ability as a direct consequence of the net loss of structural collagens and elastin.

Stretch decreased the expression of the active form of LOX in POP-VFs. We have previously shown that the expression of LOX family proteins in anterior vaginal wall biopsies of women with advanced POP was significantly reduced as compared with non-POP women

[139]. Since LOX regulates the promoter activity of COL3A1 [68], the down regulation of

LOX in response to mechanical stretch in POP VFs explains the previously reported significantly decreased expression of COL3A1 in POP VFs in comparison to non-POP VFs

[160].

Stretch also down-regulated the expression of the secreted pro and active forms of LOXL1, the pro-form of LOXL2 and BMP1 in non-POP VFs. We previously reported in vivo an increase BMP1 [35] expression in vaginal tissue biopsies derived from premenopausal POP patients in comparison to non-POP patients. Hence, the down regulation of BMP1 in non-POP

VF’s in response to mechanical stretch clearly demonstrates the differential reaction to the pre- stretch distorted collagen biogenesis environment in POP-VFs. These present in vitro results indicate that the decrease in LOX, LOXL1, LOXL2 and BMP1 enzyme expression by VFs maybe a consequence of exposure to mechanical stretch, resulting in weakened connective tissue due to decreased crosslinking of collagen and elastin polymers. Thus, mechanical stress

99 contributes to the remodeling of the ECM by controlling the expression of its fibrillar constituents and the enzymes that assemble and degrade the ECM [68]. We recognize, however, that these methods measure steady state levels. Hydroxyproline incorporation and degradation studies would have been more suitable to directly measure collagen synthesis and degradation respectively.

In conclusion, I was able to derive an enriched VF cell population, with similar attachment and proliferative abilities on collagen I, from POP and non-POP patients. However,

I detected key differences in the biological characteristics, ability to produce ECM and response to static mechanical stretch between these two cell populations. ECM turnover by VFs derived from prolapsed tissue is decreased as compared to VFs derived from vaginal tissues of non-

POP patients. In addition, POP-VF’s respond differently to stretch in comparison to non-POP

VFs. Specifically, stretched POP-VF’s express proteins that increase ECM degradation, while stretched non-POP VFs express proteins that protect tissue from acceleration of ECM turnover

(Figure 4.1).

100

Conclusions

Synthesis Degradation SynthesisSynthesis Degradation

Non-POP NS STRETCH POP-NS STRETCH STRETCH STRETCH

SynthesisSynthesis Degradation Synthesis Degradation

Figure 4.1. ECM Synthesis and Degradation in Non-POP and POP VF’s under Non-

Stretch and Stretch Conditions. There is a decreased ECM turnover primary VFs derived

from POP patients as compared to VFs derived from non-POP patients; POP-VF’s respond

differently to stretch in comparison to non-POP VFs: stretched POP-VF’s express proteins

that increase ECM degradation, while stretched non-POP VFs express proteins that protect

tissue from acceleration of ECM turnover.

101

I suggest that cultured fibroblasts may be used to improve tissue repair and augment strength in a variety of conditions, from acute to chronic wounds, and in aesthetic and reconstructive surgery. Furthermore, while fibroblasts from different anatomical sites may have similar morphology, gene expression profiling studies have shown that molecular differences, including the ability to produce ECM proteins and cytokines, are site-specific [201]. Increased

MMP expression in POP-VF’s in response to stretch may increase the risk of mesh autolysis and failure after POP surgeries and. Hence pre-stretched POP-VF’s may not be a suitable option for vaginal mucosa biological meshes. These parameters are crucial to take into account when considering the use of autologous or allogenic fibroblasts in mesh-augmented reconstructive surgery of the pelvic floor. In contrast to autologous fibroblasts, allogeneic fibroblasts may carry with them a risk of rejection or cross-infection [202]. Furthermore, the use of autologous fibroblasts has been shown to result in better restoration of the dermal skin and less scar formation in comparison with allogeneic fibroblasts [203]. However, allogenic fibroblasts can be harvested, cultured and cryopreserved prior to surgery, and are hence more readily available. Hence, allogeneic fibroblasts can be used to precondition the pelvic floor prior to the application of a permanent graft that contains autologous VFs.

102

CHAPTER 5: FUTURE DIRECTIONS

103

To date, there is a relative lack of basic scientific research focusing on the study of female pelvic floor disorders. This current study explored the fundamental differences in the biological characteristics between VFs derived from POP and non-POP patients, and the cause and effect relationship between mechanical stretch and POP development in premenopausal women. Our PCR array data demonstrated differences in expression profile of genes encoding cell-cell adhesion molecules and basement membrane constituents between POP-VFs and non-

POP VFs, under normal culture conditions and when a particular mechanical stretch environment was applied. Currently, studies are underway to confirm these data using quantitative RT-PCR, Western immunoblot and immunohistochemical analysis.

As mentioned above, the vast majority of women with POP are post-menopausal. To further understand the differences in POP development before and after menopause, the biological characteristics of cells derived from postmenopausal patients with prolapsed uteri could be examined and compared with cells from age-matched subjects with normal pelvic floor support. Using an in vitro computer-controlled stretch system, the direct effect of mechanical load on the ability of primary VFs derived from post-menopausal women to synthesize ECM proteins and enzymes responsible for ECM integrity could be tested. To understand the effect of ovarian hormones on the modulation of ECM metabolism in vaginal tissue after menopause, the potential protective effect of hormone replacement therapy or topical hormonal application against vaginal tissue deterioration would be examined. This would be done by treating POP and non-POP VFs with estrogen or a combination of estrogen and progesterone therapy before and after the application of static mechanical stretch, and studying how these ovarian hormones might modulate ECM metabolism in VFs.

I have shown that VFs derived from the prolapsed vaginal tissue of women with severe

POP are defective in their attachment, proliferative capacity and ability to produce ECM 104 proteins. Future studies should now be aimed at characterizing and comparing autologous fibroblasts derived from non-prolapsed and prolapsed vaginal tissue of POP patients, and autologous fibroblasts derived from different body sites of patients with POP. Such studies would provide more insight into if the changes in expression of the ECM proteins are inherited, or acquired due to mechanical loading in vivo and culture conditions in vitro. Consequently, by replacing defective vaginal cells with healthy autologous cells, or allogeneic non-POP VFs, the question of the potential suitability of these cells for the development of patient-specific tissue can be answered. This would enable the identification of the ideal combination of fibroblasts and mesh properties that would yield optimized epithelial-mesenchymal interactions. Potential development of a healthy tissue-engineered fascia equivalent that would be developed with autologous or allogeneic cells which could be used to reinforce the defective supportive tissue of the vagina may lead to a reduction in the wound healing time, recovery and cost of reconstructive pelvic floor surgery.

105

Appendix A: Consent Form

CONSENT TO PARTICIPATE IN A RESEARCH STUDY

INVESTIGATOR: Dr. May Alarab MBChB, MRCOG, MRCPI Fellow in Urogynecology and reconstructive pelvic surgery Department of Obstetrics & Gynecology University of Toronto, Mount Sinai Hospital 700 University Avenue, Toronto, Ontario M5G 1Z5 Tel: 416 586 4642 Fax: 416 586 3208 Email: [email protected]

TITLE: Patterns of gene expression in pelvic tissue of women, with or with out pelvic organ prolapse

You are being asked to take part in a research study. Before agreeing to participate in this study, it is important that you read and understand the following explanation of the proposed study procedures. The following information describes the purpose, procedures, benefits, discomforts, risks and precautions associated with this study. It also describes your right to refuse to participate or withdraw from the study at any time. In order to decide whether you wish to participate in this research study, you should understand enough about its risks and benefits to be able to make an informed decision. This is known as the informed consent process. Please ask the study doctor or study staff to explain any words you don’t understand before signing this consent form. Make sure all your questions have been answered to your satisfaction before signing this document.

Purpose You have been asked to participate in a study, which is designed to help understand the cause of pelvic organ prolapse (herniation or dropping of the uterus and / or the vaginal tissues past the vaginal opening). This is a very uncomfortable condition that effects women and interferes with their quality of life. It is managed either conservatively, or surgically. Both types of care have limitations and do not always offer permanent cure. Because patients affected by this condition have varying backgrounds and histories, it has been very hard to pinpoint the cause. We are asking patients who have this problem to participate in this study; we are also asking patients who do not have this condition, to participate to enable us to compare the tissue characteristics between both groups. We hope to identify the genes/proteins responsible for this

106 highly prevalent condition. This information will assist in the management and prevention of pelvic organ prolapse in the future. You will be told which group of patients you are in.

Procedures You are booked to have a hysterectomy, and / or repair of pelvic organ prolapse as agreed by you and your specialist. You could be either the effected group (have pelvic organ prolapse), or the control group (no pelvic organ prolapse). With your consent, we will take and use for testing a small amount of the upper part of the vaginal tissues (0.5 cm), removed as part of this procedure. The sample comes from the top of the vagina at its attachment to the cervix at the time of the hysterectomy, and or pelvic repair. The vaginal sample will be analyzed looking at the genetic sequencing and/or specific protein expression in the tissue sample. The aim is to identify any genes/proteins that are related to the causation of genital prolapse and the way to prevent this disorder. The vaginal tissue will be used for the purpose only. As part of the study, we may need to take 10 ml of your peripheral blood (from the arm). In addition, Dr. May Alarab will ask you to answer a brief history questionnaire (5 minutes time) after you consent to participate in the study. This questionnaire will include a review of previous medications you have been prescribed. If required, Dr. May Alarab may need to review your hospital record.

Risks There are no known risks applied to you being involved in this study.

Benefits You may not receive any medical benefit from your participation in this study. Information learned from this study may benefit other patients in the future.

Confidentiality All information obtained during the study will be held in strict confidence. No names or identifying information will be used in any publication or presentations. No information identifying you will be transferred outside the investigators in this study. During the regular monitoring of your study or in the event of an audit, your medical record may be reviewed by the Mount Sinai Hospital Research Ethics Board and/or the Investigator involved in the study.

Participation Your participation in this study is voluntary. You can choose not to participate or you may withdraw at any time without affecting your medical care.

Questions If you have any questions about the study, please call (Dr. Alarab) at (416 586 4642). If you have any questions about your rights as a research subject, please call Dr. R. Heslegrave, Chair of the Mount Sinai Hospital Research Ethics Board at (416) 586-4875. This person is not involved with the research project in any way and calling him will not affect your participation in the study.

107

Consent I have had the opportunity to discuss this study and my questions have been answered to my satisfaction. I consent to take part in the study with the understanding I may withdraw at any time without affecting my medical care. I have received a signed copy of this consent form. I voluntarily consent to participate in this study.

______Patient’s Name (Please Print) Patient’s Signature Date

I confirm that I have explained the nature and purpose of the study to the subject named above. I have answered all questions.

______Name of Person Signature Date Obtaining Consent

Do you agree to allow your tissue to be used for future research purposes? Yes ___ No __

108

Appendix B: Data To Be Collected From Each Patient Involved In The Study Age

Time of menstrual cycle.

Parity

Mode of delivery, and gestation

Forceps Vacuum

Weight of each child at delivery

Smoking

BMI Wt:……….. Ht: ………

Grade of prolapse, and duration

Family history of POP

History of stress urinary incontinence.

Bowel symptoms, (constipation).

Fecal incontinence: Yes No

Medical history

List of medication

Hormone Replacement Therapy? Yes No

Oral? Permarin? Vagifem?

Duration? When was it stopped?

Past surgical history; hernia repair varicose veins

pelvic prolapse surgery SUI surgery

Occupation

OR:

Indication

109

Appendix C: Vaginal wall biopsy site.

Full thickness specimen of anterior vaginal wall to be removed from the vaginal cuff after hysterectomy.

110

Appendix D: List of 84 ECM and Cell Adhesion Genes per Functional Group. (http://www.sabiosciences.com/rt_pcr_product/HTML/PAHS-013Z.html)

Refseq Symbol Description Collagens NM_000088 COL1A1 Collagen, type I, α 1 NM_001846 COL4A2 Collagen, type IV, α 2 NM_000093 COL5A1 Collagen, type V, α 1 NM_001848 COL6A1 Collagen, type VI, α 1 NM_001849 COL6A2 Collagen, type VI, α 2 NM_000094 COL7A1 Collagen, type VII, α 1 NM_001850 COL8A1 Collagen, type VIII, α 1 NM_080629 COL11A1 Collagen, type XI, α 1 NM_004370 COL12A1 Collagen, type XII, α 1 NM_021110 COL14A1 Collagen, type XIV, α 1 NM_001855 COL15A1 Collagen, type XV, α 1 NM_001856 COL16A1 Collagen, type XVI, α 1 ECM proteases and protease inhibitors NM_002421 MMP1 Matrix metallopeptidase 1 (interstitial collagenase) NM_004530 MMP2 Matrix metallopeptidase 2 (gelatinase A, 72kDa gelatinase, 72kDa type IV collagenase) NM_002422 MMP3 Matrix metallopeptidase 3 (stromelysin 1, progelatinase) NM_002423 MMP7 Matrix metallopeptidase 7 (matrilysin, uterine) NM_002424 MMP8 Matrix metallopeptidase 8 (neutrophil collagenase) NM_004994 MMP9 Matrix metallopeptidase 9 (gelatinase B, 92kDa gelatinase, 92kDa type IV collagenase) NM_002425 MMP10 Matrix metallopeptidase 10 (stromelysin 2) NM_005940 MMP11 Matrix metallopeptidase 11 (stromelysin 3) NM_002426 MMP12 Matrix metallopeptidase 12 (macrophage elastase) NM_002427 MMP13 Matrix metallopeptidase 13 (collagenase 3) NM_004995 MMP14 Matrix metallopeptidase 14 (membrane-inserted) NM_002428 MMP15 Matrix metallopeptidase 15 (membrane-inserted) NM_005941 MMP16 Matrix metallopeptidase 16 (membrane-inserted) NM_006988 ADAMTS1 ADAM metallopeptidase with thrombospondin type 1 motif, 1 NM_139025 ADAMTS13 ADAM metallopeptidase with thrombospondin type 1 motif, 13 NM_007037 ADAMTS8 ADAM metallopeptidase with thrombospondin type 1 motif, 8 NM_003254 TIMP1 TIMP metallopeptidase inhibitor 1 NM_003255 TIMP2 TIMP metallopeptidase inhibitor 2 NM_000362 TIMP3 TIMP metallopeptidase inhibitor 3 Trans-Membrane and Cell-Matrix Adhesion Molecules NM_001843 CNTN1 Contactin 1 NM_001903 CTNNA1 Catenin (cadherin-associated protein), α 1, 102kDa NM_001904 CTNNB1 Catenin (cadherin-associated protein), β 1, 88kDa NM_001331 CTNND1 Catenin (cadherin-associated protein), δ 1 NM_001332 CTNND2 Catenin (cadherin-associated protein), δ 2 (neural plakophilin-related arm-repeat protein) NM_181501 ITGA1 Integrin, α 1 NM_002203 ITGA2 Integrin, α 2 (CD49B, alpha 2 subunit of VLA-2 receptor) NM_002204 ITGA3 Integrin, α 3 (antigen CD49C, alpha 3 subunit of VLA-3 receptor) NM_000885 ITGA4 Integrin, α 4 (antigen CD49D, alpha 4 subunit of VLA-4 receptor) NM_002205 ITGA5 Integrin, α 5 (fibronectin receptor, alpha polypeptide) NM_000210 ITGA6 Integrin, α 6 NM_002206 ITGA7 Integrin, α 7 111

NM_003638 ITGA8 Integrin, α 8 NM_002209 ITGAL Integrin, α L (antigen CD11A (p180), lymphocyte function-associated antigen 1; α polypeptide) NM_000632 ITGAM Integrin, α M (complement component 3 receptor 3 subunit) NM_002210 ITGAV Integrin, α V (vitronectin receptor, α polypeptide, antigen CD51) NM_002211 ITGB1 Integrin, β 1 (fibronectin receptor, β polypeptide, antigen CD29 includes MDF2, MSK12) NM_000211 ITGB2 Integrin, β 2 (complement component 3 receptor 3 and 4 subunit) NM_000212 ITGB3 Integrin, β 3 (platelet glycoprotein IIIa, antigen CD61) NM_000213 ITGB4 Integrin, β 4 NM_002213 ITGB5 Integrin, β 5 NM_000201 ICAM1 Intercellular adhesion molecule 1 NM_000615 NCAM1 Neural cell adhesion molecule 1 NM_000442 PECAM1 Platelet/endothelial cell adhesion molecule NM_001078 VCAM1 Vascular cell adhesion molecule 1 Cell-Cell Adhesion Molecules and Basement Membrane Constituents NM_004425 ECM1 Extracellular matrix protein 1 NM_002026 FN1 Fibronectin 1 NM_001523 HAS1 Hyaluronan synthase 1 NM_000216 KAL1 Kallmann syndrome 1 sequence NM_005559 LAMA1 Laminin, α 1 NM_000426 LAMA2 Laminin, α 2 NM_000227 LAMA3 Laminin, α 3 NM_002291 LAMB1 Laminin, β 1 NM_000228 LAMB3 Laminin, β 3 NM_002293 LAMC1 Laminin, γ 1 (formerly LAMB2) NM_001901 CTGF Connective tissue growth factor NM_003919 SGCE Sarcoglycan, epsilon NM_000582 SPP1 Secreted phosphoprotein 1 NM_003118 SPARC Secreted protein, acidic, cysteine-rich (osteonectin) NM_000450 SELE Selectin E NM_000655 SELL Selectin L NM_003005 SELP Selectin P (granule membrane protein 140kDa, antigen CD62) NM_003119 SPG7 Spastic paraplegia 7 (pure and complicated autosomal recessive) NM_002160 TNC Tenascin C NM_003246 THBS1 Thrombospondin 1 NM_003247 THBS2 Thrombospondin 2 NM_007112 THBS3 Thrombospondin 3 NM_000358 TGFBI Transforming growth factor, beta-induced, 68kDa NM_004385 VCAN Versican NM_000638 VTN Vitronectin NM_003278 CLEC3B C-type lectin domain family 3, member B NM_004360 CDH1 Cadherin 1, type 1, E-cadherin (epithelial) NM_000610 CD44 CD44 molecule (Indian blood group)

112

REFERENCES

1. Nygaard, I., et al., Prevalence of symptomatic pelvic floor disorders in US women. JAMA, 2008. 300(11): p. 1311-6. 2. Bump, R.C., et al., The standardization of terminology of female pelvic organ prolapse and pelvic floor dysfunction. Am J Obstet Gynecol, 1996. 175(1): p. 10-7. 3. Heit, M., et al., Predicting treatment choice for patients with pelvic organ prolapse. Obstet Gynecol, 2003. 101(6): p. 1279-84. 4. Barber, M.D., et al., Sexual function in women with urinary incontinence and pelvic organ prolapse. Obstet Gynecol, 2002. 99(2): p. 281-9. 5. Olsen, A.L., et al., Epidemiology of surgically managed pelvic organ prolapse and urinary incontinence. Obstet Gynecol, 1997. 89(4): p. 501-6. 6. Rogers, G.R., et al., Sexual function in women with and without urinary incontinence and/or pelvic organ prolapse. Int Urogynecol J Pelvic Floor Dysfunct, 2001. 12(6): p. 361-5. 7. Subak, L.L., et al., Cost of pelvic organ prolapse surgery in the United States. Obstet Gynecol, 2001. 98(4): p. 646-51. 8. Brown, J.S., et al., Pelvic organ prolapse surgery in the United States, 1997. Am J Obstet Gynecol, 2002. 186(4): p. 712-6. 9. Bump, R.C. and P.A. Norton, Epidemiology and natural history of pelvic floor dysfunction. Obstet Gynecol Clin North Am, 1998. 25(4): p. 723-46. 10. Carley, M.E., et al., Obstetric history in women with surgically corrected adult urinary incontinence or pelvic organ prolapse. J Am Assoc Gynecol Laparosc, 1999. 6(1): p. 85-9. 11. Lukacz, E.S., et al., Parity, mode of delivery, and pelvic floor disorders. Obstet Gynecol, 2006. 107(6): p. 1253-60. 12. Moalli, P.A., et al., Risk factors associated with pelvic floor disorders in women undergoing surgical repair. Obstet Gynecol, 2003. 101(5 Pt 1): p. 869-74. 13. O'Boyle, A.L., et al., Pelvic organ support in pregnancy and postpartum. Int Urogynecol J Pelvic Floor Dysfunct, 2005. 16(1): p. 69-72; discussion 72. 14. DeLancey, J.O., et al., The appearance of levator ani muscle abnormalities in magnetic resonance images after vaginal delivery. Obstet Gynecol, 2003. 101(1): p. 46-53. 15. DeLancey, J.O., et al., Comparison of levator ani muscle defects and function in women with and without pelvic organ prolapse. Obstet Gynecol, 2007. 109(2 Pt 1): p. 295-302. 16. Mant, J., R. Painter, and M. Vessey, Epidemiology of genital prolapse: observations from the Oxford Family Planning Association Study. Br J Obstet Gynaecol, 1997. 104(5): p. 579-85. 17. Jeffery, S., J. P, and D.o.O.a. Gynecology, Textbook of Urogynecology 2010 University of Cape Town 18. Harris, R.L., et al., Urinary incontinence and pelvic organ prolapse in nulliparous women. Obstet Gynecol, 1998. 92(6): p. 951-4. 19. Sardeli, C., S.M. Axelsen, and K.M. Bek, Use of porcine small intestinal submucosa in the surgical treatment of recurrent rectocele in a patient with Ehlers-Danlos syndrome type III. Int Urogynecol J Pelvic Floor Dysfunct, 2005. 16(6): p. 504-5.

113

20. Buchsbaum, G.M., et al., Pelvic organ prolapse in nulliparous women and their parous sisters. Obstet Gynecol, 2006. 108(6): p. 1388-93. 21. Whitcomb, E.L., et al., Racial differences in pelvic organ prolapse. Obstet Gynecol, 2009. 114(6): p. 1271-7. 22. Swift, S., et al., Pelvic Organ Support Study (POSST): the distribution, clinical definition, and epidemiologic condition of pelvic organ support defects. Am J Obstet Gynecol, 2005. 192(3): p. 795-806. 23. Diez-Itza, I., I. Aizpitarte, and A. Becerro, Risk factors for the recurrence of pelvic organ prolapse after vaginal surgery: a review at 5 years after surgery. Int Urogynecol J Pelvic Floor Dysfunct, 2007. 18(11): p. 1317-24. 24. Dallenbach, P., et al., Risk factors for pelvic organ prolapse repair after hysterectomy. Obstet Gynecol, 2007. 110(3): p. 625-32. 25. DeLancey, J.O., The anatomy of the pelvic floor. Curr Opin Obstet Gynecol, 1994. 6(4): p. 313-6. 26. Hall, R.J., et al., Translabial ultrasound assessment of the anal sphincter complex: normal measurements of the internal and external anal sphincters at the proximal, mid-, and distal levels. Int Urogynecol J Pelvic Floor Dysfunct, 2007. 18(8): p. 881-8. 27. DeLancey, J.O., Anatomic aspects of vaginal eversion after hysterectomy. Am J Obstet Gynecol, 1992. 166(6 Pt 1): p. 1717-24; discussion 1724-8. 28. DeLancey, J.O., Anatomy and biomechanics of genital prolapse. Clin Obstet Gynecol, 1993. 36(4): p. 897-909. 29. Barber, M.D., Contemporary views on female pelvic anatomy. Cleve Clin J Med, 2005. 72 Suppl 4: p. S3-11. 30. Abramowitch, S.D., et al., Tissue mechanics, animal models, and pelvic organ prolapse: a review. Eur J Obstet Gynecol Reprod Biol, 2009. 144 Suppl 1: p. S146-58. 31. Wen, Y., et al., Differences in mRNA and protein expression of small proteoglycans in vaginal wall tissue from women with and without stress urinary incontinence. Hum Reprod, 2007. 22(6): p. 1718-24. 32. Liang, C.C., et al., Expression of matrix metalloproteinase-2 and tissue inhibitors of metalloproteinase-1 (TIMP-1, TIMP-2 and TIMP-3) in women with uterine prolapse but without urinary incontinence. Eur J Obstet Gynecol Reprod Biol, 2010. 153(1): p. 94-8. 33. Phillips, C.H., et al., Collagen metabolism in the uterosacral ligaments and vaginal skin of women with uterine prolapse. BJOG, 2006. 113(1): p. 39-46. 34. Berger, M.B., et al., Is cervical elongation associated with pelvic organ prolapse? Int Urogynecol J, 2012. 23(8): p. 1095-103. 35. Bortolini, M.A., et al., Expression of Bone Morphogenetic Protein-1 in vaginal tissue of women with severe pelvic organ prolapse. Am J Obstet Gynecol, 2011. 204(6): p. 544 e1-8. 36. Jean-Charles, C., et al., Biomechanical properties of prolapsed or non-prolapsed vaginal tissue: impact on genital prolapse surgery. Int Urogynecol J, 2010. 21(12): p. 1535-8. 37. Martins, P., et al., Biomechanical properties of vaginal tissue in women with pelvic organ prolapse. Gynecol Obstet Invest, 2013. 75(2): p. 85-92. 38. Moalli, P.A., et al., Remodeling of vaginal connective tissue in patients with prolapse. Obstet Gynecol, 2005. 106(5 Pt 1): p. 953-63. 39. Mosier, E., V.K. Lin, and P. Zimmern, Extracellular matrix expression of human prolapsed vaginal wall. Neurourol Urodyn, 2010. 29(4): p. 582-6.

114

40. Zhou, L., et al., Biomechanical properties and associated collagen composition in vaginal tissue of women with pelvic organ prolapse. J Urol, 2012. 188(3): p. 875-80. 41. Word, R.A., S. Pathi, and J.I. Schaffer, Pathophysiology of pelvic organ prolapse. Obstet Gynecol Clin North Am, 2009. 36(3): p. 521-39. 42. Goh, J.T., Biomechanical and biochemical assessments for pelvic organ prolapse. Curr Opin Obstet Gynecol, 2003. 15(5): p. 391-4. 43. Zong, W., et al., Repetitive mechanical stretch increases extracellular collagenase activity in vaginal fibroblasts. Female Pelvic Med Reconstr Surg, 2010. 16(5): p. 257- 262. 44. Le Goff, C. and V. Cormier-Daire, The ADAMTS(L) family and human genetic disorders. Hum Mol Genet, 2011. 20(R2): p. R163-7. 45. Doillon, C.J., et al., Collagen fiber formation in repair tissue: development of strength and toughness. Coll Relat Res, 1985. 5(6): p. 481-92. 46. Kadler, K.E., et al., Collagens at a glance. J Cell Sci, 2007. 120(Pt 12): p. 1955-8. 47. Prockop, D.J., A.L. Sieron, and S.W. Li, Procollagen N-proteinase and procollagen C- proteinase. Two unusual metalloproteinases that are essential for procollagen processing probably have important roles in development and cell signaling. Matrix Biol, 1998. 16(7): p. 399-408. 48. Niyibizi, C. and D.R. Eyre, Structural characteristics of cross-linking sites in type V collagen of bone. Chain specificities and heterotypic links to type I collagen. Eur J Biochem, 1994. 224(3): p. 943-50. 49. Keene, D.R., E. Engvall, and R.W. Glanville, Ultrastructure of type VI collagen in human skin and cartilage suggests an anchoring function for this filamentous network. J Cell Biol, 1988. 107(5): p. 1995-2006. 50. Aigner, T., et al., The C5 domain of Col6A3 is cleaved off from the Col6 fibrils immediately after secretion. Biochem Biophys Res Commun, 2002. 290(2): p. 743-8. 51. Gelse, K., E. Poschl, and T. Aigner, Collagens--structure, function, and biosynthesis. Adv Drug Deliv Rev, 2003. 55(12): p. 1531-46. 52. Khoshnoodi, J., V. Pedchenko, and B.G. Hudson, Mammalian collagen IV. Microsc Res Tech, 2008. 71(5): p. 357-70. 53. Mithieux, S.M. and A.S. Weiss, Elastin. Adv Protein Chem, 2005. 70: p. 437-61. 54. Wu, M.-P., Regulation of Extracellular Matrix Remodeling Associated With Pelvic Organ Prolapse. Journal of Experimental & Clinical Medicine, 2010. 2(1): p. 11-16. 55. Wieslander, C.K., et al., Regulation of elastolytic proteases in the mouse vagina during pregnancy, parturition, and puerperium. Biol Reprod, 2008. 78(3): p. 521-8. 56. Hinek, A. and M. Rabinovitch, 67-kD elastin-binding protein is a protective "companion" of extracellular insoluble elastin and intracellular tropoelastin. J Cell Biol, 1994. 126(2): p. 563-74. 57. Yanagisawa, H., et al., Fibulin-5 is an elastin-binding protein essential for elastic fibre development in vivo. Nature, 2002. 415(6868): p. 168-71. 58. Kagan, H.M., et al., Ultrastructural immunolocalization of lysyl oxidase in vascular connective tissue. J Cell Biol, 1986. 103(3): p. 1121-8. 59. Noblesse, E., et al., Lysyl oxidase-like and lysyl oxidase are present in the dermis and epidermis of a skin equivalent and in human skin and are associated to elastic fibers. J Invest Dermatol, 2004. 122(3): p. 621-30. 60. Hamalainen, E.R., et al., Molecular cloning of human lysyl oxidase and assignment of the gene to chromosome 5q23.3-31.2. Genomics, 1991. 11(3): p. 508-16.

115

61. Kosonen, T., et al., Incorporation of copper into lysyl oxidase. Biochem J, 1997. 327 ( Pt 1): p. 283-9. 62. Kessler, E., et al., Bone morphogenetic protein-1: the type I procollagen C-proteinase. Science, 1996. 271(5247): p. 360-2. 63. Uzel, M.I., et al., Molecular events that contribute to lysyl oxidase enzyme activity and insoluble collagen accumulation in osteosarcoma cell clones. J Bone Miner Res, 2000. 15(6): p. 1189-97. 64. Hohenester, E., T. Sasaki, and R. Timpl, Crystal structure of a scavenger receptor cysteine-rich domain sheds light on an ancient superfamily. Nat Struct Biol, 1999. 6(3): p. 228-32. 65. Hayashi, K., et al., Comparative immunocytochemical localization of lysyl oxidase (LOX) and the lysyl oxidase-like (LOXL) proteins: changes in the expression of LOXL during development and growth of mouse tissues. J Mol Histol, 2004. 35(8-9): p. 845- 55. 66. Li, P.A., et al., Up-regulation and altered distribution of lysyl oxidase in the central nervous system of mutant SOD1 transgenic mouse model of amyotrophic lateral sclerosis. Brain Res Mol Brain Res, 2004. 120(2): p. 115-22. 67. Li, W., et al., Localization and activity of lysyl oxidase within nuclei of fibrogenic cells. Proc Natl Acad Sci U S A, 1997. 94(24): p. 12817-22. 68. Giampuzzi, M., R. Oleggini, and A. Di Donato, Demonstration of in vitro interaction between tumor suppressor lysyl oxidase and histones H1 and H2: definition of the regions involved. Biochim Biophys Acta, 2003. 1647(1-2): p. 245-51. 69. Lelievre, E., et al., VE-statin/egfl7 regulates vascular elastogenesis by interacting with lysyl oxidases. EMBO J, 2008. 27(12): p. 1658-70. 70. Guo, Y., et al., Intracellular distribution of the lysyl oxidase propeptide in osteoblastic cells. Am J Physiol Cell Physiol, 2007. 292(6): p. C2095-102. 71. Fogelgren, B., et al., Cellular fibronectin binds to lysyl oxidase with high affinity and is critical for its proteolytic activation. J Biol Chem, 2005. 280(26): p. 24690-7. 72. Hopkins, D.R., S. Keles, and D.S. Greenspan, The bone morphogenetic protein 1/Tolloid-like metalloproteinases. Matrix Biol, 2007. 26(7): p. 508-23. 73. Bond, J.S. and R.J. Beynon, The astacin family of metalloendopeptidases. Protein Sci, 1995. 4(7): p. 1247-61. 74. Bork, P. and G. Beckmann, The CUB domain. A widespread module in developmentally regulated proteins. J Mol Biol, 1993. 231(2): p. 539-45. 75. Handford, P.A., et al., The first EGF-like domain from human factor IX contains a high- affinity calcium binding site. EMBO J, 1990. 9(2): p. 475-80. 76. Shimell, M.J., et al., The Drosophila dorsal-ventral patterning gene tolloid is related to human bone morphogenetic protein 1. Cell, 1991. 67(3): p. 469-81. 77. Takahara, K., et al., Characterization of a novel gene product (mammalian tolloid-like) with high sequence similarity to mammalian tolloid/bone morphogenetic protein-1. Genomics, 1996. 34(2): p. 157-65. 78. Kessler, E., et al., Partial purification and characterization of a procollagen C- proteinase from the culture medium of mouse fibroblasts. Coll Relat Res, 1986. 6(3): p. 249-66. 79. Canty, E.G., et al., Coalignment of plasma membrane channels and protrusions (fibripositors) specifies the parallelism of tendon. J Cell Biol, 2004. 165(4): p. 553-63. 80. Uzel, M.I., et al., Multiple bone morphogenetic protein 1-related mammalian metalloproteinases process pro-lysyl oxidase at the correct physiological site and 116

control lysyl oxidase activation in mouse embryo fibroblast cultures. J Biol Chem, 2001. 276(25): p. 22537-43. 81. Borel, A., et al., Lysyl oxidase-like protein from bovine aorta. Isolation and maturation to an active form by bone morphogenetic protein-1. J Biol Chem, 2001. 276(52): p. 48944-9. 82. Porter, S., et al., The ADAMTS metalloproteinases. Biochem J, 2005. 386(Pt 1): p. 15- 27. 83. Fernandes, R.J., et al., Procollagen II amino propeptide processing by ADAMTS-3. Insights on dermatosparaxis. J Biol Chem, 2001. 276(34): p. 31502-9. 84. Colige, A., et al., Cloning and characterization of ADAMTS-14, a novel ADAMTS displaying high homology with ADAMTS-2 and ADAMTS-3. J Biol Chem, 2002. 277(8): p. 5756-66. 85. Holmes, D.F., et al., Ehlers-Danlos syndrome type VIIB. Morphology of type I collagen fibrils formed in vivo and in vitro is determined by the conformation of the retained N- propeptide. J Biol Chem, 1993. 268(21): p. 15758-65. 86. Hulmes, D.J., et al., Pleomorphism in type I collagen fibrils produced by persistence of the procollagen N-propeptide. J Mol Biol, 1989. 210(2): p. 337-45. 87. Watson, R.B., et al., Surface located procollagen N-propeptides on dermatosparactic collagen fibrils are not cleaved by procollagen N-proteinase and do not inhibit binding of decorin to the fibril surface. J Mol Biol, 1998. 278(1): p. 195-204. 88. Tang, B.L., ADAMTS: a novel family of extracellular matrix proteases. Int J Biochem Cell Biol, 2001. 33(1): p. 33-44. 89. Roughley, P.J., Articular cartilage and changes in arthritis: noncollagenous proteins and proteoglycans in the extracellular matrix of cartilage. Arthritis Res, 2001. 3(6): p. 342-7. 90. Vazquez, F., et al., METH-1, a human ortholog of ADAMTS-1, and METH-2 are members of a new family of proteins with angio-inhibitory activity. J Biol Chem, 1999. 274(33): p. 23349-57. 91. Levy, G.G., et al., Mutations in a member of the ADAMTS gene family cause thrombotic thrombocytopenic purpura. Nature, 2001. 413(6855): p. 488-94. 92. Verma, R.P. and C. Hansch, Matrix metalloproteinases (MMPs): chemical-biological functions and (Q)SARs. Bioorg Med Chem, 2007. 15(6): p. 2223-68. 93. Ren, X.H., et al., Association between serum soluble membrane type matrix metalloproteinase-1 (MT1-MMP) levels and bone mineral density, and biochemical markers in postmenopausal women. Clin Chim Acta, 2008. 390(1-2): p. 44-8. 94. Baker, A.H., D.R. Edwards, and G. Murphy, Metalloproteinase inhibitors: biological actions and therapeutic opportunities. J Cell Sci, 2002. 115(Pt 19): p. 3719-27. 95. Cheng, M., et al., Design and Synthesis of Piperazine-Based Matrix Metalloproteinase Inhibitors. Journal of Medicinal Chemistry, 2000. 43(3): p. 369-380. 96. Murphy, G., et al., The N-terminal domain of tissue inhibitor of metalloproteinases retains metalloproteinase inhibitory activity. Biochemistry, 1991. 30(33): p. 8097-102. 97. Hidalgo, M. and S.G. Eckhardt, Development of matrix metalloproteinase inhibitors in cancer therapy. J Natl Cancer Inst, 2001. 93(3): p. 178-93. 98. Lambert, E., et al., TIMPs as multifacial proteins. Crit Rev Oncol Hematol, 2004. 49(3): p. 187-98. 99. Chirco, R., et al., Novel functions of TIMPs in cell signaling. Cancer Metastasis Rev, 2006. 25(1): p. 99-113.

117

100. Crocker, S.J., A. Pagenstecher, and I.L. Campbell, The TIMPs tango with MMPs and more in the central nervous system. J Neurosci Res, 2004. 75(1): p. 1-11. 101. Olson, M.W., et al., Kinetic analysis of the binding of human matrix metalloproteinase- 2 and -9 to tissue inhibitor of metalloproteinase (TIMP)-1 and TIMP-2. J Biol Chem, 1997. 272(47): p. 29975-83. 102. Hernandez-Barrantes, S., et al., Differential roles of TIMP-4 and TIMP-2 in pro-MMP-2 activation by MT1-MMP. Biochem Biophys Res Commun, 2001. 281(1): p. 126-30. 103. Alarab, M., et al., Expression of extracellular matrix-remodeling proteins is altered in vaginal tissue of premenopausal women with severe pelvic organ prolapse. Reprod Sci, 2014. 21(6): p. 704-15. 104. Wheatley, D.N., Diffusion, perfusion and the exclusion principles in the structural and functional organization of the living cell: reappraisal of the properties of the 'ground substance'. J Exp Biol, 2003. 206(Pt 12): p. 1955-61. 105. Funakoshi, Y. and T. Suzuki, Glycobiology in the cytosol: the bitter side of a sweet world. Biochim Biophys Acta, 2009. 1790(2): p. 81-94. 106. Pankov, R. and K.M. Yamada, Fibronectin at a glance. J Cell Sci, 2002. 115(Pt 20): p. 3861-3. 107. Ni, H., et al., Plasma fibronectin promotes thrombus growth and stability in injured arterioles. Proc Natl Acad Sci U S A, 2003. 100(5): p. 2415-9. 108. Pierschbacher, M.D. and E. Ruoslahti, Cell attachment activity of fibronectin can be duplicated by small synthetic fragments of the molecule. Nature, 1984. 309(5963): p. 30-3. 109. Butler, M., Animal cell culture and technology, in The basics. 2004, BIOS Scientific Publishers: London ; New York. p. xii, 244 p. 110. Hynes, R.O., Integrins: bidirectional, allosteric signaling machines. Cell, 2002. 110(6): p. 673-87. 111. Ginsberg, M.H., A. Partridge, and S.J. Shattil, Integrin regulation. Curr Opin Cell Biol, 2005. 17(5): p. 509-16. 112. Yurchenco, P.D. and W.G. Wadsworth, Assembly and tissue functions of early embryonic laminins and netrins. Curr Opin Cell Biol, 2004. 16(5): p. 572-9. 113. Poschl, E., et al., Collagen IV is essential for basement membrane stability but dispensable for initiation of its assembly during early development. Development, 2004. 131(7): p. 1619-28. 114. Scheele, S., et al., Laminin isoforms in development and disease. J Mol Med (Berl), 2007. 85(8): p. 825-36. 115. Preissner, K.T., Structure and biological role of vitronectin. Annu Rev Cell Biol, 1991. 7: p. 275-310. 116. Felding-Habermann, B. and D.A. Cheresh, Vitronectin and its receptors. Curr Opin Cell Biol, 1993. 5(5): p. 864-8. 117. Schvartz, I., D. Seger, and S. Shaltiel, Vitronectin. Int J Biochem Cell Biol, 1999. 31(5): p. 539-44. 118. Ulmsten, U. and C. Falconer, Connective tissue in female urinary incontinence. Curr Opin Obstet Gynecol, 1999. 11(5): p. 509-15. 119. Nakashima, Y., T.N. Wight, and K. Sueishi, Early atherosclerosis in humans: role of diffuse intimal thickening and extracellular matrix proteoglycans. Cardiovasc Res, 2008. 79(1): p. 14-23. 120. Kuettner, K.E. and J.H. Kimura, Proteoglycans: an overview. J Cell Biochem, 1985. 27(4): p. 327-36. 118

121. Tedder, T.F., et al., The selectins: vascular adhesion molecules. FASEB J, 1995. 9(10): p. 866-73. 122. Lin, S.Y., et al., Changes in the extracellular matrix in the anterior vagina of women with or without prolapse. Int Urogynecol J Pelvic Floor Dysfunct, 2007. 18(1): p. 43-8. 123. Ulmsten, U., et al., Different biochemical composition of connective tissue in continent and stress incontinent women. Acta Obstet Gynecol Scand, 1987. 66(5): p. 455-7. 124. Connell, K.A., et al., HOXA11 is critical for development and maintenance of uterosacral ligaments and deficient in pelvic prolapse. J Clin Invest, 2008. 118(3): p. 1050-5. 125. Chen, B.H., et al., Collagen metabolism and turnover in women with stress urinary incontinence and pelvic prolapse. Int Urogynecol J Pelvic Floor Dysfunct, 2002. 13(2): p. 80-7; discussion 87. 126. Chen, Y., et al., Collagen synthesis is not altered in women with stress urinary incontinence. Neurourology and Urodynamics, 2004. 23(4): p. 367-373. 127. Salman, M.C., et al., Light and electron microscopic evaluation of cardinal ligaments in women with or without uterine prolapse. Int Urogynecol J, 2010. 21(2): p. 235-9. 128. Trabucco, E., et al., Role of proteoglycans in the organization of periurethral connective tissue in women with stress urinary incontinence. Maturitas, 2007. 58(4): p. 395-405. 129. Edwall, L., K. Carlstrom, and A. Fianu Jonasson, Markers of collagen synthesis and degradation in urogenital tissue and serum from women with and without uterovaginal prolapse. Mol Hum Reprod, 2008. 14(3): p. 193-7. 130. Suzme, R., et al., Connective tissue alterations in women with pelvic organ prolapse and urinary incontinence. Acta Obstet Gynecol Scand, 2007. 86(7): p. 882-8. 131. Kannan, K., et al., Microscopic alterations of vaginal tissue in women with pelvic organ prolapse. J Obstet Gynaecol, 2011. 31(3): p. 250-3. 132. Goepel, C., Differential elastin and tenascin immunolabeling in the uterosacral ligaments in postmenopausal women with and without pelvic organ prolapse. Acta Histochem, 2008. 110(3): p. 204-9. 133. Karam, J.A., et al., Elastin expression and elastic fibre width in the anterior vaginal wall of postmenopausal women with and without prolapse. BJU Int, 2007. 100(2): p. 346-50. 134. Zong, W., et al., Alteration of vaginal elastin metabolism in women with pelvic organ prolapse. Obstet Gynecol, 2010. 115(5): p. 953-61. 135. Mathrubutham, M. and S.K. Rao, Single microassay for matrix degrading enzymes. Front Biosci, 2001. 6: p. A13-6. 136. Rao, S.K., et al., Reduced capacity to inhibit elastase in abdominal aortic aneurysm. J Surg Res, 1999. 82(1): p. 24-7. 137. Soderberg, M.W., et al., Decreased gene expression of fibrillin-1 in stress urinary incontinence. Neurourol Urodyn, 2010. 29(3): p. 476-81. 138. Man, W.C., et al., Is lysyl oxidase-like protein-1, alpha-1 antitrypsin, and neutrophil elastase site specific in pelvic organ prolapse? Int Urogynecol J Pelvic Floor Dysfunct, 2009. 20(12): p. 1423-9. 139. Alarab, M., et al., LOX family enzymes expression in vaginal tissue of premenopausal women with severe pelvic organ prolapse. Int Urogynecol J, 2010. 21(11): p. 1397-404. 140. Klutke, J., et al., Suppression of lysyl oxidase gene expression by methylation in pelvic organ prolapse. Int Urogynecol J, 2010. 21(7): p. 869-72. 141. Klutke, J., et al., Decreased endopelvic fascia elastin content in uterine prolapse. Acta Obstet Gynecol Scand, 2008. 87(1): p. 111-5. 119

142. Jung, H.J., et al., Changes in expression of fibulin-5 and lysyl oxidase-like 1 associated with pelvic organ prolapse. Eur J Obstet Gynecol Reprod Biol, 2009. 145(1): p. 117-22. 143. Liu, X., et al., Failure of elastic fiber homeostasis leads to pelvic floor disorders. Am J Pathol, 2006. 168(2): p. 519-28. 144. Alperin, M., et al., LOXL1 deficiency negatively impacts the biomechanical properties of the mouse vagina and supportive tissues. Int Urogynecol J Pelvic Floor Dysfunct, 2008. 19(7): p. 977-86. 145. Drewes, P.G., et al., Pelvic organ prolapse in fibulin-5 knockout mice: pregnancy- induced changes in elastic fiber homeostasis in mouse vagina. Am J Pathol, 2007. 170(2): p. 578-89. 146. Rahn, D.D., et al., Biomechanical properties of the vaginal wall: effect of pregnancy, elastic fiber deficiency, and pelvic organ prolapse. Am J Obstet Gynecol, 2008. 198(5): p. 590 e1-6. 147. Budatha, M., et al., Extracellular matrix proteases contribute to progression of pelvic organ prolapse in mice and humans. J Clin Invest, 2011. 121(5): p. 2048-59. 148. Yue, W., et al., Fibulin-5 suppresses lung cancer invasion by inhibiting matrix metalloproteinase-7 expression. Cancer Res, 2009. 69(15): p. 6339-46. 149. Chen, B., et al., Differences in estrogen modulation of tissue inhibitor of matrix metalloproteinase-1 and matrix metalloproteinase-1 expression in cultured fibroblasts from continent and incontinent women. Am J Obstet Gynecol, 2003. 189(1): p. 59-65. 150. Jackson, S.R., et al., Changes in metabolism of collagen in genitourinary prolapse. Lancet, 1996. 347(9016): p. 1658-61. 151. Gabriel, B., et al., Increased expression of matrix metalloproteinase 2 in uterosacral ligaments is associated with pelvic organ prolapse. Int Urogynecol J Pelvic Floor Dysfunct, 2006. 17(5): p. 478-82. 152. Kushner, L., et al., Excretion of collagen derived peptides is increased in women with stress urinary incontinence. Neurourol Urodyn, 2004. 23(3): p. 198-203. 153. Rubod, C., et al., Biomechanical properties of vaginal tissue: preliminary results. Int Urogynecol J Pelvic Floor Dysfunct, 2008. 19(6): p. 811-6. 154. Epstein, L.B., C.A. Graham, and M.H. Heit, Correlation between vaginal stiffness index and pelvic floor disorder quality-of-life scales. Int Urogynecol J Pelvic Floor Dysfunct, 2008. 19(7): p. 1013-8. 155. Lei, L., Y. Song, and R. Chen, Biomechanical properties of prolapsed vaginal tissue in pre- and postmenopausal women. Int Urogynecol J Pelvic Floor Dysfunct, 2007. 18(6): p. 603-7. 156. Couri, B.M., et al., Animal models of female pelvic organ prolapse: lessons learned. Expert Rev Obstet Gynecol, 2012. 7(3): p. 249-260. 157. Feola, A., et al., Impact of pregnancy and vaginal delivery on the passive and active mechanics of the rat vagina. Ann Biomed Eng, 2011. 39(1): p. 549-58. 158. Feola, A., et al., Parity negatively impacts vaginal mechanical properties and collagen structure in rhesus macaques. Am J Obstet Gynecol, 2010. 203(6): p. 595 e1-8. 159. Kong, D., B. Ji, and L. Dai, Stability of adhesion clusters and cell reorientation under lateral cyclic tension. Biophys J, 2008. 95(8): p. 4034-44. 160. Ruiz-Zapata, A.M., et al., Fibroblasts from women with pelvic organ prolapse show differential mechanoresponses depending on surface substrates. Int Urogynecol J, 2013. 24(9): p. 1567-75. 161. Blaauboer, M.E., et al., Cyclic mechanical stretch reduces myofibroblast differentiation of primary lung fibroblasts. Biochem Biophys Res Commun, 2011. 404(1): p. 23-7. 120

162. Maher, C.M., et al., Surgical management of pelvic organ prolapse in women: the updated summary version Cochrane review. Int Urogynecol J, 2011. 22(11): p. 1445-57. 163. Abed, H., et al., Incidence and management of graft erosion, wound granulation, and dyspareunia following vaginal prolapse repair with graft materials: a systematic review. Int Urogynecol J, 2011. 22(7): p. 789-98. 164. Gras, S. and G. Lose, The clinical relevance of cell-based therapy for the treatment of stress urinary incontinence. Acta Obstet Gynecol Scand, 2011. 90(8): p. 815-24. 165. Wang, H.J., Y.C. Chuang, and M.B. Chancellor, Development of cellular therapy for the treatment of stress urinary incontinence. Int Urogynecol J, 2011. 22(9): p. 1075-83. 166. Ho, M.H., et al., Stimulating vaginal repair in rats through skeletal muscle-derived stem cells seeded on small intestinal submucosal scaffolds. Obstet Gynecol, 2009. 114(2 Pt 1): p. 300-9. 167. Hung, M.J., et al., Tissue-engineered fascia from vaginal fibroblasts for patients needing reconstructive pelvic surgery. Int Urogynecol J, 2010. 21(9): p. 1085-93. 168. Boennelycke, M., et al., Fresh muscle fiber fragments on a scaffold in rats-a new concept in urogynecology? Am J Obstet Gynecol, 2011. 205(3): p. 235 e10-4. 169. Skala, C.E., et al., Isolation of fibroblasts for coating of meshes for reconstructive surgery: differences between mesh types. Regen Med, 2009. 4(2): p. 197-204. 170. Wen, Y., et al., Reprogramming of fibroblasts from older women with pelvic floor disorders alters cellular behavior associated with donor age. Stem Cells Transl Med, 2013. 2(2): p. 118-28. 171. Ayele, T., et al., Tissue engineering approach to repair abdominal wall defects using cell-seeded bovine tunica vaginalis in a rabbit model. J Mater Sci Mater Med, 2010. 21(5): p. 1721-30. 172. Zhang, L., et al., Musculature tissue engineering to repair abdominal wall hernia. Artif Organs, 2012. 36(4): p. 348-52. 173. Slack, M., et al., A standardized description of graft-containing meshes and recommended steps before the introduction of medical devices for prolapse surgery. Consensus of the 2nd IUGA Grafts Roundtable: optimizing safety and appropriateness of graft use in transvaginal pelvic reconstructive surgery. Int Urogynecol J, 2012. 23 Suppl 1: p. S15-26. 174. Chen, G., et al., Culturing of skin fibroblasts in a thin PLGA–collagen hybrid mesh. Biomaterials, 2005. 26(15): p. 2559-2566. 175. Kerkhof, M.H., et al., Changes in tissue composition of the vaginal wall of premenopausal women with prolapse. Am J Obstet Gynecol, 2014. 210(2): p. 168 e1-9. 176. Kawano, H., et al., Role of the lesion scar in the response to damage and repair of the central nervous system. Cell Tissue Res, 2012. 349(1): p. 169-80. 177. Chelberg, M.K., et al., Type IV collagen-mediated melanoma cell adhesion and migration: involvement of multiple, distinct domains of the collagen molecule. Cancer Res, 1989. 49(17): p. 4796-802. 178. Han, S.W. and J. Roman, Fibronectin induces cell proliferation and inhibits apoptosis in human bronchial epithelial cells: pro-oncogenic effects mediated by PI3-kinase and NF-kappa B. Oncogene, 2006. 25(31): p. 4341-9. 179. Armstrong, M.T. and P.B. Armstrong, Growth factor modulation of the extracellular matrix. Exp Cell Res, 2003. 288(2): p. 235-45. 180. DiCosmo, F., Edge Effect : The Role of Collagen in Wound Healing. Advances in Skin & Wound Care 2008. 22(1): p. 12-15.

121

181. Luzak, B., et al., Inhibition of collagen-induced platelet reactivity by DGEA peptide. Acta Biochim Pol, 2003. 50(4): p. 1119-28. 182. Martin, P., Wound healing--aiming for perfect skin regeneration. Science, 1997. 276(5309): p. 75-81. 183. Tomasek, J.J., et al., Myofibroblasts and mechano-regulation of connective tissue remodelling. Nat Rev Mol Cell Biol, 2002. 3(5): p. 349-63. 184. Desmouliere, A., et al., Apoptosis mediates the decrease in cellularity during the transition between granulation tissue and scar. Am J Pathol, 1995. 146(1): p. 56-66. 185. Makinen, J., et al., Collagen synthesis in the vaginal connective tissue of patients with and without uterine prolapse. Eur J Obstet Gynecol Reprod Biol, 1987. 24(4): p. 319- 25. 186. Klauschie, J., et al., Histologic characteristics of vaginal cuff tissue from patients with vaginal cuff dehiscence. J Minim Invasive Gynecol, 2014. 21(3): p. 442-6. 187. Szauter, K.M., et al., Lysyl oxidase in development, aging and pathologies of the skin. Pathol Biol (Paris), 2005. 53(7): p. 448-56. 188. Lambert, C.A., et al., Coordinated regulation of procollagens I and III and their post- translational enzymes by dissipation of mechanical tension in human dermal fibroblasts. Eur J Cell Biol, 2001. 80(7): p. 479-85. 189. Howard, E.W., et al., MMP-2 expression by fibroblasts is suppressed by the myofibroblast phenotype. Exp Cell Res, 2012. 318(13): p. 1542-53. 190. Lafleur, M.A., A.M. Tester, and E.W. Thompson, Selective involvement of TIMP-2 in the second activational cleavage of pro-MMP-2: refinement of the pro-MMP-2 activation mechanism. FEBS Lett, 2003. 553(3): p. 457-63. 191. Guo, C. and L. Piacentini, Type I collagen-induced MMP-2 activation coincides with up-regulation of membrane type 1-matrix metalloproteinase and TIMP-2 in cardiac fibroblasts. J Biol Chem, 2003. 278(47): p. 46699-708. 192. Brooks, P.C., et al., Localization of matrix metalloproteinase MMP-2 to the surface of invasive cells by interaction with integrin alpha v beta 3. Cell, 1996. 85(5): p. 683-93. 193. Borrirukwanit, K., et al., The type I collagen induction of MT1-MMP-mediated MMP-2 activation is repressed by alphaVbeta3 integrin in human breast cancer cells. Matrix Biol, 2007. 26(4): p. 291-305. 194. Al-Taher, H., et al., Vaginal pressure as an index of intra-abdominal pressure during urodynamic evaluation. Br J Urol, 1987. 59(6): p. 529-32. 195. O'Dell, K.K., et al., Vaginal pressure during lifting, floor exercises, jogging, and use of hydraulic exercise machines. Int Urogynecol J Pelvic Floor Dysfunct, 2007. 18(12): p. 1481-9. 196. Kasper, G., et al., MMP activity is an essential link between mechanical stimulus and mesenchymal stem cell behaviour. J Stem Cells Regen Med, 2007. 2(1): p. 56-7. 197. Yang, C.M., et al., Mechanical strain induces collagenase-3 (MMP-13) expression in MC3T3-E1 osteoblastic cells. J Biol Chem, 2004. 279(21): p. 22158-65. 198. Hawwa, R.L., et al., Differential expression of MMP-2 and -9 and their inhibitors in fetal lung cells exposed to mechanical stretch: regulation by IL-10. Lung, 2011. 189(4): p. 341-9. 199. Zhou, D., et al., Differential MMP-2 activity of ligament cells under mechanical stretch injury: an in vitro study on human ACL and MCL fibroblasts. J Orthop Res, 2005. 23(4): p. 949-57.

122

200. Visse, R. and H. Nagase, Matrix metalloproteinases and tissue inhibitors of metalloproteinases: structure, function, and biochemistry. Circ Res, 2003. 92(8): p. 827-39. 201. Chang, H.Y., et al., Diversity, topographic differentiation, and positional memory in human fibroblasts. Proc Natl Acad Sci U S A, 2002. 99(20): p. 12877-82. 202. Bello, Y.M., A.F. Falabella, and W.H. Eaglstein, Tissue-engineered skin. Current status in wound healing. Am J Clin Dermatol, 2001. 2(5): p. 305-13. 203. Lamme, E.N., et al., Allogeneic fibroblasts in dermal substitutes induce inflammation and scar formation. Wound Repair Regen, 2002. 10(3): p. 152-60.

123