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Vaginal Fibroblasts Derived From Premenopausal Women With and Without Severe Pelvic Organ Prolapse:
Differential Characteristics and Effect of Mechanical Stretch
by
Hala Kufaishi
A thesis submitted in conformity with the requirements for the degree of Master of Science
Institute of Medical Science University of Toronto
© Copyright by Hala Kufaishi 2015
ii
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.
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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.
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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.
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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
vi
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
vii
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
viii
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
ix
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
x
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
TBST-T Transforming Growth Factor Beta 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
xi
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
TBST-T Transforming Growth Factor Beta Tris-buffered saline (TBS), with 0.05% tween
xii
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
1
CHAPTER 1: LITERATURE REVIEW
2
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
3
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
4
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-
5
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
6
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
7
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
8
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
9
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].
10
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
11
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
12
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].
13
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.
14
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,
15
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 pro-
peptides 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
16
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
17
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
18
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].
19
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
Metalloelastase
MMP-12 collagens, gelatins
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
20
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
21
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
22
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
23
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
24
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.
25
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 pro-
collagen 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
26
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
27
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
28
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
29
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
30
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
31
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,
32
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
33
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
34
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
35
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
36
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
37
effect of static mechanical stretch (25% elongation) on the expression of ECM proteins, and
cell adhesion proteins, their receptors and modulators.
38
CHAPTER 2: MATERIALS and METHODS
39
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
40
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.
41
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.
42
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
43
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
44
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.
45
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
hrs inside a humidified incubator with 5% CO2 at 37oC. 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
46
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.
47
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
48
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
95oC (0.1oC/sec). Gene expression values were analyzed using the ΔΔCq mode on the CFX
49
ManagerTM 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.
50
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’
Reverse 5’-AACAGCCAGGACTCAATCCCT-3’ NM_002317
LOXL1 Forward 5’-CTGTGACTTCGGCAACCTCAA-3’ Reverse 5’-TGCACGTCGGTTATGTCGAT-3’
NM_005576
LOXL2 Forward 5’-TCGAGGTTGCAGAATCCGATT-3’ Reverse 5’-TTCCGTCTCTTCGCTGAAGGA-3’
NM_002318
LOXL3 Forward 5’-CGGATGTGAAGCCAGGAAACT-3’ Reverse 5’-AGGCATCACCAATGTGGCA-3’
NM_032603
LOXL4 Forward 5’-ACCGGCATGACATTGATTGC-3’ Reverse 5’-CATCATACTTGCAGCGGCACT-3’
NM_032211
MMP1 Forward 5’-TACGAATTTGCCGACAGAGATG-3’ Reverse 5’-GCCAAAGGAGCTGTAGATGTCC-3’
NM_002421
MMP2 Forward 5’-GAATACCATCGAGACCATGCG-3’ Reverse 5’-CGAGCAAAGGCATCATCCA-3’
NM_004530
MMP3 Forward 5’TGCTGTTTTTGAAGAATTTGGGTT-3’ Reverse 5’ACAATTAAGCCAGCTGTTACTCT-3’
NM_002422.3
MMP8 Forward 5’-GCCATCCCTTCCAACTGGTAT-3’ Reverse 5’-ATCATAGCCACTCAGAGCCCA-3’
NM_002424.2
MMP10 Forward 5’-TCATGCCTACCCACCTGGAC-3’ Reverse 5’-AATTGGTGCCTGATGCATCTTC-3’
NM_002425.2
MMP12 Forward 5’-AAGGCCGTAATGTTCCCCA-3’ Reverse 5’-CAGGATTTGGCAAGCGTTG-3’
NM_002426
MMP14 Forward 5’-TGCCATGCAGAAGTTTTACGG-3’ Reverse 5’-CCTTCGAACATTGGCCTTGAT-3’
NM_004995
TIMP1 Forward 5’-TTCTGGCATCCTGTTGTTGCT-3’ Reverse 5’-CCTGATGACGAGGTCGGAATT-3’
NM_003254
TIMP2 Forward 5’-GCGTTTTGCAATGCAGATGTAG-3’ Reverse 5’-TCTCAGGCCCTTTGAACATCTT-3’
NM_003255
TIMP3 Forward 5’-CTGCTGACAGGTCGCGTCTA-3’ Reverse 5’-GCTGGTCCCACCTCTCCAC-3’
NM_000362
TIMP4 Forward 5’-TCTGAACTGTGGCTGCCAAAT-3’ Reverse 5’-GCTTTCGTTCCAACAGCCAG-3’
NM_003256
ADAMTS2 Forward 5’-GTGTGCACCTGGCAAGCATTGTTT3’ Reverse 5’-GCCAAACGGACTCCAAGCGC-3’
NM_014244.4
BMP1 Forward 5’-GCCACATTCAATCGCCCAA-3’ Reverse 5’-TGGCGCTCAATCTCAAAGGAC-3’
NM_006132
ACTB Forward 5’ACCTTCAACACCCCAGCCATGTACG-3’ Reverse 5’CTGATCCACATCTGCTGGAAGGTGG-3’
NM_001101
TBP Forward 5’-TGCACAGGAGCCAAGAGTGAA-3’ Reverse 5’-CACATCACAGCTCCCCACCA-3’
NM_003194
YWHAZ Forward 5’-ACTTTTGGTACATTGTGGCTTCAA-3’ Reverse 5’-CCGCCAGGACAAACCAGTAT-3’
NM_003406
51
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.
52
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.
53
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
54
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
55
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.
56
Table 2.2: Summary of Antibodies Used in Immunoblot Analysis
Antibody Primary or
Secondary
Monoclonal or Polyclonal
Dilution Source Company
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 anti-rabbit IgG
Secondary Polyclonal 1:3000 Goat GE Healthcare, Buckinghamshire, UK
HRP-‐conjugated anti-sheep/ goat IgG
Secondary Polyclonal 1:3000 Donkey Serotec, 3200, Raleigh, NC, USA
HRP-‐conjugated anti-mouse IgG
Secondary Polyclonal 1:4000 Rabbit GE Healthcare, Buckinghamshire, UK
57
CHAPTER 3: RESULTS
58
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).
59
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
60
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.
61
A
62
B
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).
63
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).
64
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).
65
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).
66
A
B
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.
67
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.
68
A
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.
Control 24 hours
25% Stretch 24 hours
25% Stretch 48 hours
B
69
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
70
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).
71
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).
72
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).
73
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).
74
A
B
75
C
D
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).
76
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).
77
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).
78
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
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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).
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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.
81
A
B
Average of Total Protein Values
Ratio of Averages SEM of Average
POP NS Vs
Non-POP NS
14198.1 14949.5
0.94 2268.6 2668.7
Non POP S Vs.
Non-POP NS
3044.5 2897.4
1.05 1684.5 1741.7
POP S Vs.
POP NS
110146.4 94340.1
1.16 22898.69 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.
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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).
83
A
B
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).
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B
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).
* ** *
A
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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.
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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).
87
CHAPTER 4: DISCUSSION
88
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
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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
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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
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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
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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
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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
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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
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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
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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
97
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
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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
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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
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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).
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Conclusions
Non-POP NS POP-NS STRETCH
STRETCH STRETCH STRETCH
Synthesis Degradation
Synthesis
Synthesis
Degradation Synthesis
Synthesis 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.
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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.
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CHAPTER 5: FUTURE DIRECTIONS
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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
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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.
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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
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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.
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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 __
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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
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Appendix C: Vaginal wall biopsy site.
Full thickness specimen of anterior vaginal wall to be removed from the vaginal cuff after hysterectomy.
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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
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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)
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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.
114
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.
115
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.
116
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
117
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.
118
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.
119
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.
120
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.
121
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.
122
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.
123
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.
