1 the Mechanobiology of the Crystalline Lens Dissertation

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The Mechanobiology of the Crystalline Lens

Dissertation

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy

in the Graduate School of The Ohio State University

Bharat Kumar

Graduate Program in Biomedical Engineering

The Ohio State University

2020

Dissertation Committee:

Matthew A. Reilly, Advisor

Cynthia Roberts

Heather Chandler

Copyrighted by

Bharat Kumar

2020

Abstract

The lens is a pivotal organ in the eye; playing a crucial role in the process of accommodation, by which the eye is able to alter its focal distance. The lens continuously grows in size throughout the lifetime, unlike the globe which maintains a constant size from adulthood. This growth is a result of lens epithelial cell (LEC) proliferation, which ultimately leads to an increase in the number of fiber cells.

Changes in the size, stiffness, and shape of the lens contribute to the etiology of age- related refractive issues in the lens, namely presbyopia, and cataracts. Additionally understanding the forces that control the proliferation of LECs has implications in developing therapies for posterior capsule opacification (PCO) and translational research in clinical applications for lens regeneration. The processes governing the growth of the lens are therefore of great clinical interest; however, they are not fully understood.

This dissertation considers the broadest context of the translational utility of understanding lens growth, beginning with the long-term goal of regenerating a lens following cataract extraction. This review is followed by the first basic science studies investigating mechanobiological regulation of lens growth. This represents a significant step towards understanding lens biology since, to date, all such studies have been conducted without consideration for the refractive state of the lens.

In the first study, and for the first time, LECs were found to be mechanosensitive in vitro using a bespoke stretching device. The LEC proliferation rate was found to depend strongly on the amplitude and frequency of stretching. This dependence was effectively eliminated via chemical inhibition of yes-associated protein (YAP) activation using verteporfin. These findings suggest that zonular tension is a major driving force for lens growth.

The second study investigated the localization of proliferative activity. This spatial distribution of proliferating LECs was altered significantly depending on whether static or cyclic strains were applied. Prior studies on cellular mechanisms of lens growth have used non-accommodating species (i.e. mouse), finding that adult lens growth is driven by

LEC proliferation in the “germinative zone” near the equator. When lenses were not stretched or stretched to a fixed extent, proliferation was observed primarily in the equatorial region. When oscillatory stretching was performed, proliferation was more uniform across the epithelium. The strong influence of equatorial stretching therefore must be considered when considering the growth and morphogenesis of the human lens.

These novel findings have a significant impact in the understanding of the lens growth and are foundational in the field of lens mechanobiology.

By demonstrating that the LECs are mechanosensitive, that YAP is involved in this mechanosensation, and that location of these changes depends on the dynamic nature of the applied stretching, many new research questions are available and may be studied using the methods described herein.

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Dedication

This work is dedicated to my parents, Vijay and Lakshmi for the support that they have

provided through these many years of work.

Acknowledgments

I would like to thank my advisor, Dr. Matthew Reilly for his help and guidance throughout my graduate school career. Without his support none of this would have been possible.

I would also like to thank the other members of my committee, Dr. Cynthia Roberts and

Dr. Heather Chandler. As well as the other members of the Reilly lab: Wade Rich, Sam

Croarkin, and Daniel Mackessey. And the faculty at OSU, namely: Dr. Timothy

Plageman, Dr. Swindle-Reilly, and Dr. Keith Gooch.

The material support provided by The Ohio State University was invaluable in completing this work, specifically the Campus Microscopy & Imaging Facility and the

James Comprehensive Cancer Center. And finally, our partners at Delaware Meats, and

Bay Foods, and the Pig King.

Vita

2013...... B.Eng. Biomedical Engineering, Vanderbilt University

2018...... M.S Biomedical Engineering, The Ohio State University

2014-2015 ...... Graduate Research Associate, Department of Biomedical

Engineering, University of Texas at San Antonio

2016-Present ...... Graduate Research Associate, Department of Biomedical

Engineering, The Ohio State University

Publications

1. Kumar B, Chandler HL, Plageman T, Reilly MA. Lens Stretching Modulates

Lens Epithelial Cell Proliferation via YAP Regulation. Investigative

Ophthalmology & Visual Science 2019;60:3920-3929.

2. Kumar B, Reilly MA. The Development, Growth, and Regeneration of the

Crystalline Lens: A Review. Current Eye Research 2020;45:313-326.

Fields of Study

Major Field: Biomedical Engineering

Table of Contents

Abstract ...... ii Dedication ...... iv Acknowledgments...... v Vita ...... vi Table of Contents ...... vii List of Tables ...... x List of Figures ...... xi Chapter 1. Introduction ...... 1 Clinical Implications of Understanding Lens Growth ...... 1 The Role of Lens Mechanobiology...... 4 The Mechanical Stimulation of Whole Lenses in vitro ...... 5 Research Objectives ...... 6 1. Conduct a Literature Review on the Development and Regeneration of the Lens . 6 2. Determine whether LECs are Mechanosenitive ...... 6 3. Determine whether LEC Mechanosenisitvity is YAP-Dependent...... 7 4. Determine whether LEC Proliferation Localization Changes in Response to Stretching ...... 7 Chapter 2. The Development, Growth, and Regeneration of the Crystalline Lens: A Review ...... 9 Introduction ...... 9 A Historical View of Lens Regeneration ...... 10 A Brief History of Regenerative Biology ...... 10 Eye of Newt: A History of Research on Lens Regeneration in Urodele ...... 12 Lens of Frog: Research on lens regeneration in Xenopus ...... 13 The History of Mammalian Lens Regeneration Research ...... 14 Anatomy and Physiology of the Lens ...... 16 vii

Lens Development ...... 16 Structure and Growth of the Lens ...... 19 De Novo Lens Regeneration ...... 23 Wolffian Lens Regeneration ...... 24 Cornea-lens Regeneration ...... 27 LEC Mediated Lens Regeneration ...... 30 Clinical Significance ...... 36 Conclusions ...... 38 Chapter 3. Lens Stretching Modulates Lens Epithelial Cell Proliferation via YAP Regulation ...... 40 Introduction ...... 40 Methods...... 43 Lens Stretching Organ Culture ...... 44 Flow Cytometry ...... 47 Fluorescent Microscopy ...... 48 Statistical Analysis ...... 49 Results ...... 50 Labelling Index of LECs Varies with Culture Time ...... 50 Increasing Stretch Amplitude Increases LEC Proliferation in Whole Lens Cultures51 LEC Proliferation Increases with Stretching Frequency in Whole Lens Cultures ... 53 Stretching Alters the Localization of LEC Proliferation Across the Lens Capsule .. 55 Inhibition of YAP Function Blocks Mechanotransductive Effects of Stretching on LEC Proliferation ...... 58 Stretching Alters YAP Localization in LECs ...... 61 Discussion ...... 63 Chapter 4. Changes in the Spatial Distribution of Lens Epithelial Cell Proliferation Resulting from Lens Stretching ...... 69 Introduction ...... 69 Methods...... 72 Lens Stretching Organ Culture ...... 72 Confocal Microscopy ...... 74 Computational Modeling Lens Capsule Strain ...... 76 Statistical Analysis ...... 80

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Results ...... 81 Labeling Index of Proliferative Activity across Whole Lens Capsules ...... 81 Localization of Proliferative Activity in Lenses Subjected to Different Stretching Regimes...... 82 Differences in LEC Labeling Index Distribution in Response to Differing Strain Regimes...... 85 Analysis of Lens Capsule Strains and Proliferation Distribution ...... 86 Discussion ...... 87 Chapter 5. Conclusions and Future Directions ...... 90 Chapter 6. Bibliography ...... 94

List of Tables

Table 3.1 Labeling Indices of Lenses Cultured Under Varying Static Strain Amplitudes 52 Table 3.2 Labeling Indices of Lenses Cultured Under Varying Cyclic Strain Frequencies ...... 54 Table 3.3 Labeling Indices of Verteporfin Treated and Paired Control Lenses Cultured Under Varying Static Strain Amplitudes ...... 58

List of Figures

Figure 2.1 An overview of mammalian eye development ...... 17 Figure 2.2 Schematic diagram of the regions of the lens epithelium ...... 21 Figure 2.3 Schematic illustrations of the process of Wolffian lens regeneration (a–e), cornea-lens regeneration (f–j), and LEC meditated regeneration (k–o) ...... 24 Figure 2.4 Wolffian lens regeneration in newts from transdifferentiation in the dorsal iris (di) ...... 26 Figure 2.5 Lens regeneration in larval Xenopus via transdifferentiation of the cornea .. 29 Figure 2.6 Overview of LEC mediated lens regeneration ...... 33 Figure 3.1 Lens Stretching Organ Culture Setup ...... 46 Figure 3.2 Variation of labeling index under null stretch with different times spent in culture ...... 51 Figure 3.3 Variation of labeling index with respect to strain amplitude ...... 53 Figure 3.4 Variation of labeling index with respect to strain frequency ...... 55 Figure 3.5 Variation in EdU labeling localization under different strain conditions...... 57 Figure 3.6 Variation of labeling index with respect to strain amplitude of lenses treated with a YAP inhibitor, verteporfin, and their paired controls ...... 59 Figure 3.7 Labeling index of cyclically stretched lenses treated with and without verteporfin ...... 60 Figure 3.8 Localization of YAP in lenses cultured with and without verteporfin under static and null stretch conditions ...... 62 Figure 4.1 Model geometry (a) and meshed model (b) of the porcine line ...... 77 Figure 4.2 Components of the stretching force ...... 79 Figure 4.3 Variation of total labeling indices of lenses subjected to different strain regimes...... 82 Figure 4.4 Representative mosaic confocal microscopy images of null (A), static (B), and cyclic (C) lenses, as well as the localized labeling indices of regions of null (n=3) (D), static (n=3) (E), and cyclic (n=3) (F) lenses and the comparison of the localized labeling indices of the three stretching regimes (G) ...... 84 Figure 4.5 Cumulative distribution function of Ki-67 labeling as a function of distance from the anterior pole in pixels in lenses subjected to different stretching regimes...... 85 Figure 4.6 : Meridional strains (left) and circumferential strains (right) as a function of arc length ...... 86 Figure 4.7 Comparison of Ki67 labeling in whole porcine lenses stretched uniformly around the circumference, resulting in an increase in equatorial diameter of 12% (left). Averaging strain anisotropy and proliferation in 10 equally-spaced increments of arc

xi length between the anterior pole and equator indicated a significant log-linear correlation (n=2; p<0.0001) (right)...... 87

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Chapter 1. Introduction

Clinical Implications of Understanding Lens Growth

The lens plays a significant role in accommodation – the process by which the eye alters its focal between objects at different distances. The lens grows in size continuously throughout a person’s lifetime while the size of the globe remains constant from adulthood1, 2. Lens growth results from the proliferation and differentiation of the lens epithelial cells (LECs) into lens fiber cells3-5. Lens fiber cells have a larger volume than

LECs, thus the total volume of the lens is increased by each instance of LEC proliferation and differentiation6. It is not yet well understood what the contribution of changes in the size and shape in the lens over time have on the onset of presbyopia and cataracts7-12.

However, establishing a link between the mechanical forces acting upon the lens during accommodation and LEC proliferation is important in understanding the relationship between presbyopia and lens growth. Additionally, elucidating the factors influencing the proliferation of LECs deepens the understanding of the etiology of posterior capsule opacification (PCO) and the regulation of lens regeneration13-16.

The loss of accommodative ability with age, known as presbyopia, is correlated with the growth of the lens. When the lens is disaccommodated to focus on distant objects, the ciliary muscle is relaxed, leading to the lens being stretched due to the tension applied by the zonules17-20. During accommodation, the ciliary muscle contracts, releasing the 1 tension on the zonules, allowing the lens to elastically recoil to a rounder shape. The diameter and force of contraction of the ciliary muscle has not been found to decrease with age21-26. The mechanical properties of the zonules, which connect the ciliary muscle to the lens are also independent of age27, suggesting that the human lens capsule may experience dynamic tension even after the onset of presbyopia. Despite the properties of the ciliary muscle and zonules not changing with age, accommodative amplitude decreases over time until it becomes difficult to focus on nearby objects. This implies that the etiology of presbyopia originates within the lens, but more work is needed to determine what part lens growth plays in its onset.

Human lenses follows a biphasic model of growth where the lens grows quickly in the first stage of life, followed by the growth rate slowing after reaching adulthood. After reaching adulthood, the lens grows linearly with about 1.38 mg of lens material being added each year5, 28-30. As the lens grows the accommodated diameter increases with age, while the disaccommodated diameter remains constant31. Therefore, the change in lens diameter between the disaccommodated and accommodated state decreases with age. As the size and shape of the lens changes with age, it has also been observed that the bulk stiffness of the lens increases32-35. The changes in size, shape, and stiffness of the lens allow us to hypothesize that as an individual ages, the accommodated state of the lens grows into more closely resembling the disaccommodated state, therefore resulting in the lens being less able to focus on nearby objects. The correlation between lens growth and the onset of presbyopia make finding a causal link of great interest.

The proliferation of LECs has a clinical relevance beyond presbyopia. Pathological migration of LECs onto the posterior capsule or intraocular lens following cataract extraction can lead to a secondary cataract36. Recent work has demonstrated that the lens epithelium is sensitive to changes in its mechanical environment37. Cataract surgery disrupts both the biochemical and biomechanical homeostasis of the LECs and alters their proliferative activity38. LEC proliferation and migration is influenced by contact inhibition, and the removal of much of the epithelium during cataract surgery induces a wound healing response in the LECs; additionally, the removal of the lens fiber cell bundle removes a physical barrier between the LECs and the posterior capsule leading to pathological proliferation39-41. PCO can be inhibited in vitro by using pharmaceutical agents targeting the cytoskeleton42; these drugs prevent the transduction of signals initiated by mechanical stresses on a cell to the nucleus resulting in changes in protein expression43. Together, these results suggest that LECs are strain-responsive (i.e. mechanical cues from their environment alter the cells’ behavior). In intact human lenses, irrespective of age, capsule stresses are predominately equibiaxial except near the equator, the strain anisotropy coincides with germinative zone (GZ) where the majority of LEC proliferation is thought to take place41, 44, 45. Departure from equibiaxial stresses normally present in the capsule following cataract extraction may drive cell migration and morphological changes leading to PCO46. Following lens fiber cell bundle removal, it is likely that the local lens capsule strains are significantly altered, although work is needed to measure these changes. It can therefore be hypothesized that the changes in local capsule strains following cataract extraction induce the pathological proliferative

3 response in LECs that precede PCO.

The Role of Lens Mechanobiology

The driving forces governing lens growth are not yet fully understood. However certain aspects of lens growth and LEC proliferation have been studied. It is well established in literature that LEC proliferation in non-accommodating species mainly occurs in the GZ near the equator, and the LECs migrate towards the equator and elongate and differentiate into lens fiber cells3, 5, 45, 47. The biological processes governing

LEC proliferation are less well understood, although it has been observed that fibroblast growth factor (FGF) signaling plays an important role in the proliferation and differentiation of LECs48-50. The Hippo and YAP signaling pathways, as well as the Wnt and Notch pathways have been implicated in regulating the proliferation LECs and their differentiation into fiber cells48-50. These studies elucidate how biochemical growth factors influence lens growth and describe some of the signaling pathways controlling this growth. They do not, however, examine the role of mechanical forces in influencing lens growth.

Other cell types have shown sensitivity to stretching, such as: mesenchymal stem cells, myoblasts, epithelial cells, and keratinocytes51-54. Proliferation was found to be upregulated in these cell types when both cyclic and static stretching regimes were applied. The proliferation response was dependent on the magnitude of the strain applied to the cells as well as the frequency of the cyclic stretching51, 52. Before this study, the effects of mechanical stretching on the proliferation of LECs were unexplored. 4

Establishing a link between the mechanical forces acting upon the lens during accommodation and the proliferation of the LECs was important in understanding the processes that govern lens growth.

Axial strains acting upon the lens, similar to those experienced during disaccommodation, have been observed in murine lenses during compression experiments to lead to a reversible increase in LEC area indicating that mechanical loading of the lens is transduced onto its cells55. This change in LEC area provides strong evidence that a lifetime of accommodation and disaccommodation and the resulting changes in LEC area would play a part in driving strain-responsive behaviors in LECs.

The Mechanical Stimulation of Whole Lenses in vitro

Technical challenges in reproducing the in vivo environment of the lens have made studying the effects of biochemical and biomechanical factors upon lens growth difficult. Lens flat mount explants often alter LEC morphology leading them to be unsuitable for studying the mechanobiology of the lens56. Animal models possessing a mechanism of accommodation similar to humans tend to be prohibitively expensive. We hypothesize that lenses of non-accommodating species are probably held under a consistent equatorial load, although that is yet to be experimentally demonstrated. It is therefore necessary to culture whole lenses ex vivo and mechanically stretch them in order to maintain the biological, biochemical, and biomechanical milieu of the natural microenvironment of the lens.

In order to study the relationship between lens stretching and the proliferation of LECs

5 while preserving the complex microenvironment present within the lens, it was necessary to culture whole lens tissues under different stretching conditions for a period of time ex vivo. A lens stretching bioreactor was developed for this purpose. Lens stretchers have been employed in previous studies to provide insight on the biomechanical57-62 and optomechanical63, 64 properties of the lens rather than the biological response of living tissue. Thus, they were unsuitable for use in the sterile environment of an incubator. This necessitated the development of a lens stretcher capable of applying both static and cyclic loading conditions on the lens ex vivo, while remaining autoclavable to ensure sterility.

Research Objectives

1. Conduct a Literature Review on the Development and Regeneration of the Lens

A literature review on the development and regeneration of the lens was conducted in order to contextualize this work with previous studies of the growth of the lens. Lens development describes how the lens develops and grows in utero and understanding what biological processes take place during this time informs our understanding of how the lens continues to grow throughout life. Lens regeneration is the repair of the lens after the removal of either the whole lens or part of the lens, of particular interest is that some mammalian lenses are capable of regenerating some of the fiber cell bundle after its removal. The processes governing this regrowth are likely linked to the mechanobiology of the lens epithelium.

2. Determine whether LECs are Mechanosenitive

This study was designed to determine whether the mechanical forces experienced by the lens during the process of accommodation contribute to lens growth by increasing

6 the proliferation rate of LECs. A lens organ culture bioreactor was developed that was capable of reproducing the biochemical milieu and cyclic tension exerted on the human lens throughout life, which overcame the technical challenges associated with studying the intact lens epithelium ex vivo. This technique allowed detailed study of the mechanobiological response of the lens epithelium to stretching ex vivo while closely mimicking that experienced by the human lens in vivo.

3. Determine whether LEC Mechanosenisitvity is YAP-Dependent

Identifying signaling pathways regulating mechanosensation and proliferation in the lens was the next logical step in understanding the mechanobiology of LECs.

Activation of YAP signaling is a potential mechanism by which the axial loading of the lens by disaccommodation is transduced onto the LECs and upregulates proliferative activity. Earlier studies have observed the expression of YAP in rodent LEC explants and the role it plays in regulating LEC proliferation50. Stretch-induced YAP activation and a resulting upregulation of cell proliferation has been observed in other cell types, including epithelial cells from other tissue types (i.e. mammary, lung, and skin epithelium

65-68) but had not yet been observed in the lens. This study sought to establish a link between YAP signaling and stretch-induced LEC proliferation by introducing a YAP inhibitor into the les culture medium and observing the effects on LEC proliferation.

4. Determine whether LEC Proliferation Localization Changes in Response to Stretching

This study attempted to examine if mechanically stretching the lens altered the distribution of LEC proliferation. The localization of proliferative activity along the capsule in lenses subjected to different types of stretching regimes: null, static, and cyclic 7 was determined. Confocal microscopy was used on the fluorescently labeled lenses to determine the positional information of proliferative activity. This allowed the hypothesis that LEC proliferation localization was altered in response to stretching to be tested.

Chapter 2. The Development, Growth, and Regeneration of the Crystalline Lens: A Review

This chapter has been published as a review article in Current Eye Research14.

Introduction

Regeneration of the lens can be described as the repair of the lens following the removal of some or all of the organ. Lens regeneration following the total removal of the lens with its capsule, de novo lens regeneration, has been observed to occur in either one of two ways: the regenerated lens arising from the dorsal root of the iris in newts,

Wolffian regeneration69, 70, or the new lens arising from the larval cornea in Xenopus laevis69, 71, cornea-lens regeneration. While de novo lens regeneration has only been observed to occur in newts and certain species of pre-metamorphic frogs, the ocular lens is capable of spontaneously regenerating from the residual lens epithelial cells (LECs) following removal of the contents of the lens capsule, provided that the lens capsule is left relatively intact. This LEC mediated regeneration occurs in a wider number of species including mammals72.

The processes by which de novo and LEC mediated lens regeneration occur can only be understood when placed within the context of the embryologic development and subsequent growth of the lenticular organ. In early eye development the lens is formed by the invagination and closure of the surface ectoderm. The surrounding surface ectodermal

9 tissue goes on to differentiate into the cornea; and the iris and ciliary body differentiate from nearby neural ectoderm73. With the total removal of the lens organ, de novo regeneration necessitates the new lens tissue to emerge from the transdifferentiation of other surrounding tissues. The conditions necessary to induce the different forms of de novo lens regeneration and the processes by which they occur are informed by the primordial tissues from which the transdifferentiating tissue arises69, 74.

LEC mediated lens regeneration is closely linked to the process by which the intact lens grows throughout life. The lens epithelium is continuously proliferating and differentiating into lens fiber cells2, 45, 75. When the lens fiber cell bundle is removed, the remaining LECs are able to regenerate the lens through the same process of proliferation followed by differentiation. Lens regeneration occurs as either the formation of a new lens or the replacement of removed tissue from the lens. The processes governing how the lens initially forms and grows over time provide valuable insight in the mechanisms governing the processes of lens regeneration.

A Historical View of Lens Regeneration

A Brief History of Regenerative Biology

Knowledge of the ability for some living beings to regenerate lost tissue dates back to antiquity. The ancient Greeks Empedocles (490-430 BCE) and Aristotle (384-322

BCE) commented on the ability of some lizards to regenerate their tails76. However, it would take several centuries for the study of tissue regeneration to mature from remarking at a natural oddity to a more rigorous field of study.

The 18th century saw the beginning of meticulous cataloguing of tissue regeneration in various different species, and these efforts would serve as the foundation of all future work in the field of tissue regeneration research. The earliest known work in the field was a report by René Antoine Ferchault de Réaumur in 1712 on the ability of a crayfish to regenerate limbs after amputation77. In the mid-1700s, He was followed by Abraham

Trembley, Charles Bonnet, and Peter Simon Pallas who respectively added hydras

(named after mythological monster of ancient Greece with similar regenerative capabilities), annelids, and planarians to the list of animals with regenerative capabilities78. The first reported study of vertebrate regeneration was conducted in 1768 by Lazzaro Spallanzani on salamanders which were capable of regenerating their tails, limbs, and jaws following amputation79-81. The earliest known recording of eye tissue regeneration was Bonnet’s Oeuvres d’histoire naturelle et de philosophie (1781) in which the partial dismemberment of a salamander’s eye resulted in the appearance of a complete eye after several months.

In 1880 Phillipeaux confirmed the regenerative prowess of the salamander’s eye, observing that part of the eye must remain in order to be regenerated (described in

Morgan 1901 and Vergara 2018)76, 82. In 1891 Colucci described the ability of newts to regenerate their lenses after removal even during their adult stages83. In 1895 Wolff published his studies describing the mechanism of this form of lens regeneration where the iris pigmented epithelium transdifferentiated into a new lens, earning this process the name: Wolffian regeneration70, 74.

In 1901, Thomas Hunt Morgan published “Regeneration”, in which he summarized and evaluated the preceding work on regenerative biology and recontextualized the field of study as inextricably linked to developmental biology. He viewed regeneration as a fundamental developmental process widespread between species instead of cases of adaptation independently arising at different branches of the tree of life. Morgan also established that regeneration occurs in one of two main categories: epimorphosis and morphallaxis. Epimorphosis refers to the regeneration of new tissue involving cell proliferation and morphallaxis involves the remodelling of existing tissue without cellular proliferation. Wolffian regeneration in salamanders as well as the other types of lens regeneration discussed in this review are all examples of epimorphosis where the new lens arises from proliferating cells76, 82, 84.

Eye of Newt: A History of Research on Lens Regeneration in Urodele

In the 1930s, Tadao Sato studied Wolffian regeneration in newts and discovered that it is only the dorsal iris pigmented epithelium (IPE) that gives rise to a new lens and the ventral epithelium is incapable of doing so under normal conditions85-87.

In 1940, Stone and Sapir at Yale University identified the mechanism behind Wolffian regeneration relied on the transdifferentiation of the IPE, meaning the IPE cells dedifferentiated, lost their pigmentation, then redifferentiated into another cell type, primary lens fiber cells88. Eguchi used electron microscopy to characterize the histological and temporal events of newt lens regeneration89, 90.

Panagiotis A. Tsonis was mentored by Eguchi and then became a professor at the

University of Dayton, where he and his collaborators unravelled the regulatory mechanisms governing newt lens regeneration76. In 2005 the Tsonis group was able to induce lens regeneration from the ventral IPE91

Advances in genomic and proteomics allowed for a greater understanding of the machinery governing regeneration. The Tsonis lab in collaboration with Thomas Braun and Mario Looso contributed to the first transcriptome of the North American newt92

Efforts were also made to create a linkage map of the newt genome93 and to identify the genes governing cell cycle initiation94. The work done in characterizing the newt genome, proteome, and transcriptome will be foundational in future efforts to manipulate the regenerative response.

Lens of Frog: Research on lens regeneration in Xenopus

Newts are not the only animals capable of regenerating the crystalline lens. In

1963 Freeman published findings on lens regeneration in frogs71. Xenopus laevis are capable of regenerating a removed lens, but only while in their larval state. Unlike newts, which are capable of fully regenerating the lens throughout their lifespan, the regenerative ability of Xenopus decreases with age; and rather than arising from the iris, the new Xenopus lens regenerates from the corneal epithelium71, 95.

The cornea and the lens both share the same embryonic origin and the ability of the larval

Xenopus corneal epithelium to regenerate a new lens may be evidence of an extended pluripotency of the larval surface ectoderm96-100. In addition to being able to regenerate the lens, Kha et al. have shown that the eye is able to regrow after removal at the tailbud

13 stage, replacing the missing structures contained within101. Perry et al. in 2013 identified the expression of pluripotent factors expressed in the larval Xenopus corneal epithelium and may point to the presence of oligopotent epithelial stem cells present in the corneal epithelium102.

The History of Mammalian Lens Regeneration Research

The history of research into mammalian lens regeneration began in 19th century

France where Cocteau and Leroy-d’Etoille incised the lens capsule of a rabbit, removed the lens, and six months later the lens capsule was “perfectly transparent” and the lens was of similar volume and consistency compared to before removal72, 103. An attempt to reproduce these results by Backhausen in 1827 was unsuccessful; however, inflammation, short follow-up period, and less than ideal technique may have contributed to the lack of success72, 104, 105. Research into lens regeneration in rabbits continued throughout the 1800s. Loewenhardt suggested the capsule as the source of the newly regenerated lens106. Day, Middlemore, Mayer, and others all saw regeneration in rabbits, finding that the rabbit may have retained vision in the regenerated eye107, cats and dogs were unable to produce a similar regenerative response108, and that holes in the regenerated lens material corresponded to the lesions in the capsule formed during lens removal72, 109. In 1872, Milliot saw that lens regeneration was faster in younger animals and occurred near the lens equator, anterior capsule, but not the posterior capsule104. In

1900, Randolph reviewed previous studies in the field, and from his own experiments, observed that regeneration was dependant on the lens capsule, cell differentiation

14 occurred in the equatorial zone, the regenerated lens could achieve similar size as the original lens, and that outcomes improved if some of the cortex was left behind72, 105.

Regeneration in mammals other than rabbits was observed by Milliot in 1872 (dogs, sheep, cats), Agarwal in 1964 (primates), and Gwon in 1993 (cats)104, 110, 111. In the mid-

20th century mammalian lens regeneration began to be approached from an embryological and developmental perspective. During lens vesicle formation, some head ectodermal cells remain in the vesicle and cytolyse; Paul Chanturishvili hypothesized that the cytolyzing cells release products required for lens development72, 112. Chantururishvili and Sicharulidze implanted embryonic cytolyzed skin ectoderm into the capsule following lens extraction and observed better regenerative outcomes in lenses with the transplants compared to the small cataractous lenses that did not receive the transplant72,

112. Subsequent studies by Stewart and Espinasse113, Agarwal et al.110, and Angra et al.114 found improved lens regeneration following implantation of embryonic tissue. However, studies by Binder et al.115, Pettit116, and Metz117 found either no significant difference in the lenses with implants, or teratoma formation in the implant72, 115.

More recent work, including that conducted by the Gwon group has shown that the regeneration of the mammalian lens proceeds in similar fashion to the embryological development of the lens. In both cases, epithelial cells proliferate along the anterior and posterior capsule and the posterior epithelial cells elongate, anteriorly migrate, eventually lose their nuclei, and differentiate into fiber cells to form the fetal nucleus118. Further work by the Gwon group sought to refill the emptied lens capsule in order to stimulate

15 lens regeneration and improve optical performance of the regenerated lens in an effort to advance the study of mammalian lens regeneration towards its eventual clinical application15, 111, 119, 120.

A recent study by Lin et al. utilized the endogenous stem cells within the lenses of pediatric cataract patients to regenerate the lens following cataract surgery121, 122. This is the first clinical application of mammalian lens regeneration and is likely the first step in bringing lens regeneration into the clinic.

Anatomy and Physiology of the Lens

Lens Development

The embryological development of the eye in mammals begins with the formation of the optic pit as a result of the evagination of the ventral forebrain/prechordal mesenchyme (Fig. 2.1)73. The optic pits enlarge to form the optic vesicles and in parallel the overlying surface ectoderm thickens to form the lens placode123. The lens placode thickens towards the presumptive neural retina of the optic vesicle and both primordial tissues invaginate such that the presumptive neural retina and retinal pigmented epithelium form the optic cup and the folded surface ectoderm forms the lens pit. The lens vesicle forms from the closing of the lens pit and detachment from the surface ectoderm, the lens vesicle matures into the lens and the now closed surface ectoderm differentiates into the cornea123.

Figure 2.1 An overview of mammalian eye development Differentiated eye tissues are color-coded: lens/cornea (blue); neural retina (green); retinal pigmented epithelium (RPE) (red); optic stalk (orange); and mesenchyme (grey). (a) Formation of the optic vesicle by an evagination of the mesenchyme (M) resulting in the formation of the optic pit (OP). The optic pit region can be divided into two regions: The surface ectoderm (SE) containing the presumptive lens ectoderm (PLE), and the neural tube (NT) containing the presumptive neural retina (PNR), and presumptive ventral optic stalk (POS). (b) Continued growth of the optic vesicle results in close proximity between the lens placode (LP) and the neural retina (NR). The RPE and ventral optic stalk (VOS) continue to develop. (c) Invagination of the optic vesicle induces formation of the lens pit (LPit). The lens pit later closes and detaches from the surface ectoderm to form the lens vesicle. (d) Mature eye: cornea (C); lens epithelium (LE); lens fiber cells (LF); lens capsule (LC); iris (I); ciliary body (CB); neural retina (NR); optic nerve (ON).

The lens vesicle has a large central cavity, which is lined by a monolayer of lens epithelial cells. The cells at the posterior pole elongate and differentiate into primary lens fiber cells. Cells at the anterior pole of the lens remain LECs. Mitotically active epithelial 17 cells near the equatorial region elongate and differentiate into secondary fiber cells, which encircle the primary fiber cells and lose their organelles as they move inwards from the outer to the inner cortex123. This arrangement results in the anterior epithelial cells being the only mitotically active cells in the lens and the source of all new fiber cells.

In some non-mammalian vertebrates, specifically zebrafish, cavefish, and possibly

Xenopus, a lens vesicle does not form during eye development124, 125. Instead the lens placode continues to thicken forming a lens mass and eventually delaminates from the surface ectoderm124, 125

During development of the anterior segment of the eye, cells from the neural crest mesenchyme migrate between the lens and the corneal epithelium. These cells differentiate into the corneal endothelium, iris and ciliary body126-129. Studies have shown that if during this stage in eye development the lens were to be removed, the mesenchymal cells would not differentiate into the proper tissues and would instead form a disorganized mass of mesenchymal cells behind the corneal epithelium126, 130. The presence of the lens is crucial for the normal development of the cornea and iris, as well as other anterior eye tissues during development. In the absence of the lens, de novo lens regeneration occurs due to the transdifferentiation of cells in the cornea and the iris in premetamorphic Xenopus laevis and newts, respectively69. The importance of the lens in the developmental organization of the anterior tissues of the developing eye suggest that the lens produces certain signalling factors that the anterior tissues respond to; and when

18 the lens is removed, it is possible that the absence of these factors is what initiates de novo regeneration in newts and frogs126.

Structure and Growth of the Lens

The mature vertebrate lens is an oblate spheroid consisting of the lens capsule composed primarily of collagen IV and laminins. A more comprehensive composition of the capsule can be found in work by Brian P. Danysh and Melinda K Duncan131. Its inner anterior surface is covered with a monolayer of lens epithelial cells (LECs). The rest of the lens is made up of lens fiber cells which lose their nuclei and organelles after differentiating from the LECs2. The lens is unique in that it grows throughout life by the addition of new cells inside the capsule. Old cells are compacted within the center of the lens nucleus132, with newly added cells continuously proliferating and differentiating at the equatorial regions of the anterior LEC monolayer.

The lens capsule is a pivotal structure in the lens as, unlike other basement membranes, it completely encapsulates the lens and as a result mediates any signalling entering and leaving the lens. Additionally interactions between the LECs and the capsule are important in the development of a transparent lens. SPARC (secreted protein acidic and rich in cysteine) is a matricellular protein found only near the LECs; when SPARC was disrupted in murine lenses, differentiation of LECs into fiber cells were as well133.

SPARC has a high affinity for collagen IV, the primary component of the lens capsule, and has cell-binding epitopes134. It is likely that SPARC plays an important role in linking the LECs to the capsule135 and when this link is disrupted, the LECs lack vital signals allowing them to differentiate and mature into fiber cells133 19

The epithelium of the lens has a critical role in maintaining both the homeostasis and growth of the lens. The lens epithelium is also the driving force behind lens growth, as the LECs are the only mitotically active cell type within the lens5. Despite the lens epithelium’s importance in the growth of the lens, the factors governing the LECs’ proliferative activity are not well understood5. Histologically, the LEC monolayer can be classified as a simple epithelium, with all of the LECs in direct contact with the basement membrane, the lens capsule2. The lens increases in both size and mass throughout life.

This lens growth is rapid at the onset of life and slows in adulthood29, 30. The driving force behind the continuous growth of the lens is cellular proliferation which occurs exclusively in the LEC monolayer,136 resulting in increased lens volume when an LEC differentiates into a far larger fiber cell137. In neonatal animals, mitotic events can be detected throughout the lens epithelium; however, over time, the proliferative activity of the lens becomes concentrated within a region anterior of the lens equator known as the germinative zone (GZ)45, 75. Posterior to the GZ is the transitional zone (TZ) near the edge of the LEC monolayer where the LECs begin differentiating into the lens fiber cells

(Fig. 2.2). LECs in the TZ are post-mitotic and have permanently withdrawn from the cell cycle138.

Figure 2.2 Schematic diagram of the regions of the lens epithelium LECs near the anterior pole (blue) exhibit little mitotic activity. LECs in the region just anterior to the equator (green) are in the mitotically active germinative zone (GZ). LECs anterior to the equator (red) are in the post-mitotic transitional zone (TZ) where LECs begin to differentiate into lens fiber cells at the epithelial margin. The concentric shells of the fiber cell bundle are also shown (light blue).

Lens fiber cells comprise the majority of the volume of the lens and arise from differentiated LECs which lose their nucleus and organelles during differentiation139. The differentiating LECs elongate and their volume increases. These elongating fiber cells arrange themselves in concentric growth shells with each shell contributing to the expansion of the lens47, 140, 141. Due to the morphology of the lens, and its encapsulation

21 within the capsule, the lens is unique in that the proliferation of LECs and subsequent differentiation of those daughter cells into lens fiber cells is not accompanied by cell loss; therefore, the fiber cells in the center of the lens, the primary fiber cells, have been there since the formation of the lens and are some of the oldest cells in the body142.

Additionally, to ensure the optical clarity of the lens fiber cell bundle, the nucleus and organelles of the fiber cells degrade during differentiation142-144. The fiber cells lose their nucleus and organelles and are left with a high concentration of crystallin proteins and little in the way of the cellular machinery associated with the maintenance of homeostasis145. The age of the lens fiber cells, their lack of cellular machinery, and high concentration of crystallin proteins contribute to cataractogenesis, due to the accumulating risk of damage to the crystallin proteins over time resulting in the aggregation of proteins and the formation of cataracts146.

Lens regeneration is of particular importance in the context of cataracts due to the process by which cataract surgeries are performed. The lens fiber cell mass is broken up using phacoemulsification and the fragments are removed through a hole in the lens capsule147.

After removal of the fiber cells, only the incomplete capsule and epithelium remain. An intraocular lens (IOL) is implanted within the capsule to restore some of the focusing power of the lens; however, they are not yet able to completely restore the accommodative ability of the lens, making it difficult to shift focus from near to distant objects148. It is therefore an attractive option to explore the use of lens regeneration, in this case LEC-mediated regeneration, to refill the emptied lens capsule and restore visual function to the lens. Pathological behaviour of LECs following cataract extraction can 22 lead to posterior capsular opacification (PCO), a form of secondary cataract. Improved understanding of LEC wound-healing and proliferative behaviours will eventually lead to improved surgical approaches, pharmacological approaches, or IOL designs to mitigate

PCO36.

De Novo Lens Regeneration

De novo lens regeneration uses differentiated cells in tissues adjacent to the damaged lens to repair the damaged lens. Some species can replace the lens even if it is completely removed with its capsule. This happens through the process of transdifferentiation where differentiated tissues dedifferentiate and redifferentiate into new lenticular tissue (Fig. 2.3). This occurs in two forms: Wolffian regeneration, where the parent tissue originates from the dorsal root of the iris and has been observed only in newts and cornea-lens regeneration where the new lens forms from the transdifferentiation of cells in the corneal epithelium in larval frogs of the genus

Xenopus69.

Figure 2.3 Schematic illustrations of the process of Wolffian lens regeneration (a–e), cornea-lens regeneration (f–j), and LEC meditated regeneration (k–o) In (b) and (g), the intact lens is removed along with its lens capsule. (l) Shows the empty capsule with the lens fiber cell bundle removed, leaving the lens epithelium and lens capsule intact. Eye structures are labelled as: cornea epithelium (ce); dorsal iris (di); lens capsule (lc); lens epithelium (le); lens (ln); lens placode (lp); lens vesicle (lv); regenerated lens fiber cells (rlf); regenerated lens (rln); and ventral iris (vi).

Wolffian Lens Regeneration

Some newts of the order Urodela are the only vertebrates in which adult de novo lens regeneration has been observed. Research into this remarkable ability of the newt lens to regenerate after complete removal dates back to 1891 when Collucci83 observed this phenomena and the study by Wolff in 189570, after whom the process is named

Wolffian regeneration. The newly regenerated lens arises from the transdifferentiation of neuroectoderm-derived dorsal iris pigmented epithelial (IPE) cells149-151. Interestingly, the regenerative power of the lens in amphibians capable of Wolffian regeneration does not diminish with age or repeated regenerative events. One particularly long-lived newt had its lens removed 18 times over the course of 16 years and, each time, fully regenerated it while reproducing all of the normal structures152.

The process of IPE-dependant Wolffian regeneration follows shortly after the removal of the lens along with its capsule, followed by the growth of a new lens at the dorsal iris epithelium (Fig. 2.3)153. During the first 4 days post-lentectomy, IPE cells in the dorsal region of the iris dedifferentiate, characterized by a loss of pigmentation and initiation of cellular proliferation150, 151, 154. At the dorsal pupillary margin, IPE cells continue this dedifferentiation by proliferating and losing pigmentation until 8-10 days post- lentectomy. At this point, cell elongation and synthesis of lens-specific proteins begin. At

10 days post-lentectomy, the lens vesicle begins to form, followed by the elongation of the posterior cells and their transdifferention into lens fiber cells. The anterior cells of the nascent lens vesicle become LECs89, 90. Around 12-16 days post-lentectomy, the lens vesicle thickens and crystallin proteins begin to be produced (Fig. 2.4)69. During days 15-

19 the IPE dedifferentiation process ceases and the primary lens fiber cell bundle forms from the transdifferentiated IPE cells. The secondary lens fiber cell bundle forms from proliferating and differentiating LECs. By the 30th day post-lentectomy, lens regeneration is complete and the transdifferentiation of IPE cells has given rise to a new functional lens89, 90, 154-156. 25

Figure 2.4 Wolffian lens regeneration in newts from transdifferentiation in the dorsal iris (di) (a) Ten-days post-lentectomy, the early lens vesicle (arrow) forms. (b) Fifteen-days post- lentectomy. The cells at the posterior part of the vesicle (arrow) elongate to form lens fiber cells. (c) Twenty-days post-lentectomy. A regenerated lens with lens fibres (lf) covered by the lens epithelium (le). Reprinted from Tsonis. 200469. 26

The molecular mechanisms governing the transdifferentiation of IPEs have also been the target of much study. Differential analysis of the transdifferentiating dorsal IPE and the non-transdifferentiating ventral IPE has shown an upregulation of several hundred genes in the tissues responsible for lens regeneration; these genes perform various functions including: gene expression, protein homeostasis, extracellular matrix deposition, cell cycle, cell proliferation, and DNA replication94. The FGF (fibroblast growth factor) signalling pathway has been suggested to control Wolffian lens regeneration, FGF2 and the FGF receptors FGFR1 and FGFR3 have been found to be essential in inducing IPE transdifferentiation157-159. FGF expression has been observed to induce early lens genes such as pax6 and sox2 within a few days of the lentectomy160-162. Studies have also indicated that activation of the Wnt (wingless) signalling pathway found in the IPE undergoing lens regeneration163, 164. The role FGF plays in initiating the dedifferentiation of IPE cells provides insight on how Wolffian regeneration is induced. A FGF concentration gradient exists between the anterior and posterior chambers of the eye, with the vitreous having a much higher concentration of FGF than the aqueous48, 49, 165. The presence of the lens acts as a physical barrier between the vitreous and aqueous and, when removed, the concentration of FGF near the iris would potentially increase. This may be the initiating factor in the transdifferentiation response in newts166.

Cornea-lens Regeneration

Cornea-lens regeneration has been found to occur in pre-metamorphic frogs of the genus

Xenopus. Unlike the seemingly unlimited potential for the newt iris to produce more lenses, the ability for the larval frog to regenerate the lens from the inner layer of the

27 outer corneal epithelium declines with age. In general, cornea-lens regeneration in

Xenopus occurs through the transdifferentiation of ectodermal central cornea epithelial cells into a lens vesicle that, over time, forms a new lens71, 167. Despite the difference in location as well as the age dependence, the process by which cornea-lens regeneration produces a new lens is quite similar to what happens in Wolffian regeneration.

Cornea-lens regeneration in Xenopus has been described to occur in five phases (Fig.

2.5)69. In the first stage, 1-2 days post-lentectomy, inner corneal epithelial cells change from squamous to cuboidal epithelial cells71. LECs in an intact lens are also cuboidal epithelial cells2. The differentiating epithelial cells then begin to thicken into a placodal structure. The third stage begins 3 days post-lentectomy, when the cell aggregate begins to separate from the corneal epithelium by invaginating into the vitreous. This leads into stage 4, 5 days post-lentectomy, when a lens vesicle forms and separates from the cornea.

The lens vesicle contains primary fiber cells arising from the differentiation of the posterior cells in the lens vesicle and the anterior cells differentiate into LECs. After the fifth stage, the new LECs begin proliferating and differentiating into secondary lens fiber cells and the corneal epithelial cells return to their original squamous configuration71, 160.

This process closely resembles the initial embryological development of the lens, where the ectodermal tissue (presumptive cornea) thickens into a lens placode, invaginates, detaches to form the lens vesicle, and finally differentiates into the lens epithelium and primary fiber cell bundle73.

Figure 2.5 Lens regeneration in larval Xenopus via transdifferentiation of the cornea (a) The lens placode forms from the thickening of cells. (b) Lens vesicle formation begins with the invagination of cells and closure of the lens pit. (c) Lens fiber cells begin to differentiate (red staining with anti crystallin antibody). (d) Differentiation of lens fiber cells continues. (e) The lens has increased in size and has positioned itself by the dorsal and ventral iris. Eye structures are labelled as: cornea (c), iris (i), lens placode (lp), lens vesicle (lv), lens fiber cells (lf). Reprinted from Tsonis 200469.

The molecular signalling mechanisms that induce cornea-lens regeneration are thought to include factors in the vitreous humour that are produced by the neural retina168-170. This was demonstrated by a study where a piece of the outer cornea was implanted in the posterior chamber in contact with the vitreous and formed a new lens despite the fact that the old lens was still present171. Growth factors such as BMP (bone morphogenic protein), FGF, and those involved in the Wnt signalling pathway were identified as potential candidates for inducing cornea-lens regeneration172, 173. While in Wolffian lens regeneration, the Wnt signalling pathway needs to be activated to induce lens regeneration, it plays a different role in cornea-lens regeneration. Functional studies by

Hamilton et al. have shown that Wnt/β-catenin signalling needs to be supressed for successful lens regeneration to occur via the cornea-lens route174. When the Wnt pathway was held in a state of active signalling, there was a reduction in cases of successful lens regeneration in Xenopus. Conversely, when Wnt signalling was supressed, there was no reduction in regenerative ability174. It was also observed that Wnt was suppressed in untreated control Xenopus post-lentectomy. Interestingly, in the embryological development of the lens, the formation of the lens is also accompanied by the suppression of Wnt signalling123, 175.

LEC Mediated Lens Regeneration

Unlike the previously discussed methods of lens regeneration, LEC mediated regeneration cannot regenerate the whole lens with its capsule but instead involves replacing removed lens fiber cells after they are removed, provided the lens capsule is mostly intact. Furthermore, LEC mediated lens regeneration is not exclusive to

30 amphibians and has been studied in mammals. The spontaneous regeneration of the lens after removal of the capsular contents was first observed in 1825 in rabbits103. An incision was made in the lens capsule and the fiber cell bundle was removed; six months later the lens capsule contained lenses that were similar in size and consistency to the original lens103. It is of particular interest that the methods employed in this study to remove the lens are generally similar to the methods used in modern cataract surgery; specifically, the contents of the lens are removed while the capsule is left mostly intact147.

Further studies have elucidated the conditions necessary for LEC mediated lens regeneration to occur: the capsule as well as the LECs needed to remain in place, the posterior capsule needed to be uninjured, the anterior capsule needed to be relatively sound, and the posterior and anterior capsules could not adhere72. From these requirements, it can be concluded that the LECs are the source of new lens material and that the capsule plays a vital role as the structural framework for the new lens. Beyond the gross requirements in the structural integrity of the lens capsule post-lentectomy, LEC mediated lens regeneration is faster and more efficient in younger animals176. This agrees with clinical observations in that younger patients develop posterior capsule opacification

(PCO), caused by proliferation of LECs onto the posterior lens capsule faster and more frequently than in older patients72. This is further supported by studies showing that LEC proliferative activity is higher in younger animals and decreases with age2, 45, 75.

The mechanism by which the lens regenerates in mammals is similar to the manner in which embryological development occurs (Fig. 2.6)118. The remaining LECs on the

31 anterior capsule proliferate and migrate posteriorly during the first week post-lentectomy.

The posterior epithelial cells elongate and begin to differentiate into primary fiber cells by the end of the first month. After the second month, proliferation is only seen near the equatorial regions like in the normal adult lens118. After the initial fiber cells bundle forms and LEC proliferation becomes contained to the germinative zone (GZ), the regeneration of the lens resembles the normal growth of the lens, albeit with a higher rate of proliferation72. The new LECs produced in the GZ differentiate in the transition zone

(TZ) and move inwards and compact the fiber cells in the center.

Figure 2.6 Overview of LEC mediated lens regeneration (a): The intact lens is shown with the fiber cell bundle regularly arranged in concentric shells. (b): The LEC monolayer remains after fiber cell removal. (c): Lens epithelial cells proliferate and migrate across the posterior capsule. (d): Lens fibers begin to differentiate and elongate at the lens equator, this process is slower and more irregular along the posterior capsule. (e): The resulting irregularly arranged primary lens fibers are compacted centrally as subsequent (secondary) lens fibers differentiate and surround them, creating opacities.

Extensive work has been done by the Gwon group to best determine how to induce and control the properties of LEC mediated regeneration. In rabbits, the newly regenerated lens has been observed to fill roughly half the capsule and be relatively clear; however, as the lens matured it became opaque and slit lamp microscopy revealed that the fiber cells had grown in an irregular pattern with many whorl-like patterns containing vacuoles and opacities177, unlike the normal lens where the fiber cell bundle consists of many regularly packed concentric rings141, 178. A more normal arrangement of lens fiber cells and therefore better optical properties were observed when the hole in the anterior capsule

33 was sealed with a collagen patch and the capsule was filled with air111. However, the alignment of regenerated fiber cells was not as uniform as the normal lens. Further studies attempted to implant a disposable contact lens into the regenerating lens to determine if the structural support would improve the growth patterns in the regenerated lens. After removal of the lens and insertion of the polymer lens, the capsule was sealed with a collagen patch and distended with Healon, an ophthalmic viscoelastic material119.

Lens regenerating with the implant had poor optical clarity and cells anterior to the implant were clearer than those posterior to it119. Better results were produced when lenses were filled with a biodegradable hyaluronic scaffold with a higher clarity and more regular pattern of fiber cell growth120.

The aforementioned studies suggest that mechanically supporting the regenerating lens improves the outcome of the newly regenerated lens in terms of the arrangement of lens fiber cells and the clarity of the new lenticular organ. It can therefore be hypothesized that the lens epithelium is sensitive to changes in its mechanical environment and that the change in the mechanical loading of the LECs from the removal of the fiber cell bundle is what leads to the irregular regeneration of the lens. Bito and Harding hypothesized that mitosis in the lens epithelium is controlled by mechanical tension in the zonules179, 180.

Additionally, simulating accommodative forces in whole porcine lens cultures in vitro has been found to induce mitotic activity in the lens epithelium37. LEC proliferation and migration patterns have been shown to be influenced by contact inhibition, the sudden removal of the fiber cell bundle, and the change in the mechanical environment of the lens epithelium is what leads to their migration onto the posterior capsule and PCO39, 40. 34

Cataract surgery is known to alter LEC proliferation and disrupt both the biochemical and biomechanical homeostasis of the lens38. Departure from the equibiaxial stresses thought to be present normally in the lens capsule may drive cell migration and lead to morphological changes resulting in PCO. Such departures could potentially be initiated by removal of the fiber cell bundle41. and anisotropic strains have been found to exist in the intact human lens capsule near the equator, coinciding with the GZ44. It has also been observed that PCO can be inhibited in vitro by using pharmaceutical agents targeting the cytoskeleton42. The cytoskeleton is typically involved in transducing external mechanical cues. Thus, inhibiting this mechanosensation in LECs may disrupt cytoskeleton-based signalling activity and potentially decouple the cellular response from mechanotransductive signals. A link can therefore be drawn from the changing mechanical environment of the lens epithelium after removal of the contents of the capsule and the irregular growth of the lens fiber cells during LEC mediated lens regeneration. It is possible that the irregular mechanical environment within the lens capsule during regeneration contributes to the irregular regeneration of the lens, and that altering the mechanical loading of the epithelium to more closely match the normal lens may be beneficial in translating lens regeneration into the clinic. The rate of lens differentiation may vary in the different parts of the capsule resulting in abnormal alignment of the earliest regenerating lens fiber cells118. In order to ensure a regular alignment of regenerating fiber cells, it would be desirable to delay lens differentiation until the entire capsule is covered by a confluent monolayer of LECs. Ideally the embryological environment could be replicated such that the capsule would be supported

35 by a scaffold of amniotic fluid or similar constituents mirroring the environment present during embryological development15, 120.

Clinical Significance

One end goal of research into lens regeneration is successful regeneration of the human lens after damage to the lens or removal during cataract surgery. The nature of cataract surgeries, where the lens fiber cells are removed but the lens capsule and epithelium are left mostly intact in situ makes LEC mediated lens regeneration a particularly attractive avenue of research. Recent work by Lin et al. has made significant progress towards this goal16. In order to minimize the damage the lens capsule, a small

(1-2 mm) capsulorhexis was performed superiorly and away from the visual axis as has been done in lens refilling and all lens regeneration studies rather than centrally as is done in current cataract surgery181, 182. In vivo studies were conducted on rabbits and macaques where the surgical technique was performed to remove the lens but preserve as many of the endogenous LECs as possible. Within 7 weeks post-lentectomy, a new translucent, biconvex tissue formed and was comparable to a healthy lens16. Further in vivo studies tested the surgical technique on 1-3 month old macaques, leading to the regeneration of a biconvex, translucent lens after 5 months16. Finally, a clinical trial was conducted on infants with congenital cataracts. The current standard surgical procedure for infant cataract removal uses a large anterior continuous curvilinear capsulorhexis combined with posterior continuous curvilinear capsulorhexis and anterior vitrectomy.

The clinical trial attempted to induce human lens regeneration using the minimally invasive surgical technique. Pediatric cataract patients under 2 years of age underwent the

36 minimally invasive surgery. Within one month of surgery, the capsular opening was closed and a translucent, biconvex lens had regenerated after 3 months. Eight months after surgery, the accommodative amplitude increased to 2.5 diopters, from 0.1 diopters measured 1 week before surgery, indicating that the lenses were functional16. The results of the Lin group are a large step towards realizing the potential of human lens regeneration; however, the results achieved would unlikely extend to older patients due to the reduced proliferative activity of LECs with age and the increased size of the capsule potentially contributing to a mechanical environment not conducive to lens regeneration.

The success of Lin et al. does raise questions on how to broaden the applicability of the treatment. None of the patients in the study had traumatic, hereditary, or metabolic cataract. While the Lin study only included patients with idiopathic cataracts, non- idiopathic cataracts comprise over half of all pediatric cataract patients121, 122. Trauma to the lens may damage the capsule and reduce its ability to regenerate the lens and hereditary and metabolic disorders would likely prevent the lens from regenerating itself as well. Using cultured cells would be a potential avenue of research in order to provide source material for a healthy lens when the endogenous cells may be incapable of doing so. Additionally, refilling the emptied capsule with a scaffold would potentially speed up transdifferentiation and prevent adhesion of the anterior and posterior capsules122. The capsulorhexis technique used by Lin et al. is an important advancement in regenerative ophthalmology and by minimizing damage to the capsule allows for the conditions necessary for lens regeneration to take place and opens up avenues of research121, 122.

In addition to the promising clinical trials of Lin et al.16, a recent in vivo study by Murphy et al183 opens a new avenue of study for lens regeneration research by producing light focusing micro-lenses from pluripotent stem cells. These micro-lenses provide a new platform with which to study lens regeneration without the complications posed by animal studies while more closely matching the human regenerative response.

Conclusions

The ocular lens is able to spontaneously regenerate via one of several species- dependent mechanisms. In newts, age-independent and repeated regeneration of the lens following total removal using transdifferentiation of the dorsal iris pigmented epithelium.

In frogs of the genus Xenopus, the corneal epithelium is able to transdifferentiate into a new lens while the frog is in its larval state. Finally, mammalian regeneration requires the capsule and lens epithelial cell layer to remain mostly intact in order to refill the contents of the capsule.

All of these methods of lens regeneration are informed by the processes governing the embryological development and continued growth of the lens. In the de novo routes of lens regeneration, the new lens vesicle is formed in a manner similar to the developing mammalian lens despite occurring in amphibians. The regenerated lens vesicle in a de novo lens and the emptied lens capsule in mammalian regeneration proceed with the differentiation of cells into primary fiber cells and epithelial cells. This is succeeded by the lens growing through the usual means of lens epithelial cells in the germinative zone proliferating and differentiating into secondary fiber cells leading to the growth of the lens. 38

The ultimate end of lens regeneration research is to repair damage to the lens or refill the lens following cataract surgery. This has recently been used to good effect in infant humans, and while regenerating the lens after surgery for age-related cataracts is not yet within reach, significant progress is being made.

Chapter 3. Lens Stretching Modulates Lens Epithelial Cell Proliferation via YAP Regulation

This chapter has been published in Investigative Ophthalmology & Visual Science37.

Introduction

The lens is the pivotal tissue in accommodation – the process by which the eye alters its focal distance from far to near. The lens continues to grow in size throughout a person’s lifetime while the size of the globe of the eye stays constant through adulthood1,

2. This growth is a result of lens epithelial cell (LEC) proliferation, which ultimately leads to an increase in the number of fiber cells3. Since a lens fiber cell has a much larger volume than an LEC, the lens progressively becomes larger as a result of LEC proliferation6. The age-related changes in lens size and shape contribute to presbyopia and cataracts7, 9-11.

The driving force(s) for this continuous growth remain unknown, at least in part due to technical challenges with reproducing the lens’ complex in vivo environment which includes biochemical and biomechanical influences. Flat-mount lens explants are frequently used to probe specific aspects of LEC behavior, but may not be appropriate for examining mechanobiological behavior due to altered cell morphology184 and mechanical environment. Similarly, species which accommodate using a human-like mechanism are few and prohibitively expensive for many basic scientific studies.

In disaccommodation, tension is applied to the lens capsule via the zonules due to relaxation of the ciliary muscle20. During accommodation, this tension is released via ciliary muscle contraction, allowing the lens to elastically recoil to a rounder shape. The ciliary muscle remains active into old age18, 21, 24, 25 and the mechanical properties of zonules are independent of age27, implying that the human lens capsule may experience cyclic tension even after the onset of presbyopia. In the context of presbyopia, younger lenses exhibit a large magnitude in focal length changes during stretching, while lenses above the age of 60 showed no changes in focal length with stretching, such that the eye is only able to focus on distant objects17, 19, 20, 185. Axial strains acting upon the lens, similar to those experienced during disaccommodation, have been observed to lead to a reversible increase in LEC area indicating that mechanical loading of the lens is transduced onto its cells55. This change in LEC area provides strong evidence that a lifetime of accommodation and disaccommodation and the resulting changes in LEC area would play a part in driving strain-responsive behaviors in LECs.

Recent work has demonstrated that the lens epithelium is sensitive to changes in its mechanical environment. LEC proliferation is altered during cataract surgery which disrupts both the biochemical and biomechanical homeostasis of the lens38. Departure from equibiaxial stresses in the capsule may drive cell migration and morphological changes leading to posterior capsular opacification (PCO)46. Such anisotropic strains have been found to exist in the intact human lens capsule near the equator, coinciding with the region in which proliferation is known to occur in the mouse lens3, 44. PCO can be inhibited in vitro by using pharmaceutical agents targeting the cytoskeleton42; the 41 cytoskeleton is known to convey information about mechanical stresses on a cell to the nucleus resulting in changes in protein expression43. Together, these results suggest that

LECs are strain-responsive cells (i.e. they can alter their behavior in response to mechanical cues from their environment). However, these studies have primarily focused on pathological LEC differentiation leading to PCO and therefore did not examine whether proliferation rates or biomarker expression levels were directly influenced by mechanical stretching.

Activation of YAP/Taz signaling is a potential mechanism by which the axial loading of the lens by disaccommodation is transduced onto the LECs and upregulates proliferative activity. Earlier studies have observed the expression of YAP in rodent LEC explants and the role it plays in FGF-induced LEC proliferation50. The role of YAP in the mechanoregulation of LEC stretch-induced proliferation is not well established; however, stretch-induced YAP activation and a resulting upregulation of cell proliferation has been observed in other cell types, including epithelial cells from other tissue types (i.e. mammary, lung, and skin epithelium 65-68). It is well established that YAP has a pivotal role in transducing mechanical cues from stretching into increased cell proliferation. This study seeks to provide evidence that this same behavior is exhibited by LECs, thus identifying a contributing factor to the continuous growth of the lens throughout life.

In order to study the effects of lens stretching on the proliferation of LECs without disturbing the complex microenvironment present within the lens, it was necessary to culture whole lens tissues under different loading conditions for an extended period of

42 time ex vivo. For this purpose a lens stretching bioreactor was developed. Lens stretchers have been employed in previous studies; however, they were primarily used to provide insight on the biomechanical57-62 and optopmechanical63, 64 properties of the lens and were unsuitable for use in the sterile environment of an incubator. The lens stretcher developed for this study was capable of applying both static and cyclic loading conditions on the lens ex vivo, while remaining autoclavable to ensure sterility.

This study was designed to determine whether the mechanical forces experienced by the lens during the process of accommodation contribute to lens growth by increasing the proliferation rate of LECs as well as to identify what role YAP signaling plays in the stretch-induced proliferation of LECs. By developing a lens organ culture bioreactor capable of reproducing the biochemical milieu and cyclic tension exerted on the human lens throughout life, we have overcome the technical challenges associated with studying the intact lens epithelium ex vivo. This technique allows detailed study of the mechanobiological response of the lens epithelium to stretching ex vivo while closely mimicking that experienced by the human lens in vivo. Furthermore, agonists or inhibitors of specific molecular pathways may be used to elucidate the underlying mechanobiological mechanisms involved.

Methods

All animal tissues were used in accordance with institutionally approved protocols.

Lens Stretching Organ Culture

Freshly enucleated porcine eyes were obtained from a local abattoir (Delaware

Meats, Delaware, OH). Extra-ocular tissue was removed and the whole globe was disinfected by submersion in 0.5% povidone-iodine (Sigma-Aldrich, St. Louis, MO) in phosphate buffered saline (PBS) for 5 minutes, then transferred into PBS. The globe was removed from the PBS and partial-thickness incisions were made with a scalpel along the limbus and the equator. The cornea and the iris were carefully removed. The globe was bisected along the equatorial incision and the posterior half was discarded. The posterior portion of the vitreous was removed and the anterior portion of the vitreous was left attached to the anterior section of the eye. Eight radial cuts were made through the sclera spaced 45° apart to create eight flaps surrounding the lens. To ensure uniform stretching about the lens’ circumference a stapler was used to attach each of the scleral flaps to a 40 mm diameter silicone disk (McMaster-Carr, Elmhurst, IL) with a 10 mm diameter hole in the center with the lens positioned in the central hole (Fig. 3.1).

The silicone disks were mounted onto a bespoke stretching ring device which attached to eight equally-spaced peripheral holes in the silicone disk. Lenses mounted in this way were submerged in pre-warmed, serum-free medium 199 (M199) with Earle’s salts L- glutamine, and sodium bicarbonate, supplemented with 0.1% bovine serum albumin, 100

IU/ml penicillin, 100 mg/ml streptomycin and 2.5 mg/ml Amphotercin B (Sigma-

Aldrich). Some lenses were exposed to the Yes-associated protein (YAP) function inhibitor verteporfin (5 μM verteporfin, Sigma-Aldrich). Lens tissue was incubated at

37°C, 5% CO2 for 24 hours. After 23 hours in culture, 0.1% of 10 mM EdU (5-ethynyl- 44

2’-deoxyuridine) (Thermo-Fisher, San Jose, CA) in DMSO was added to the cell culture media for the remaining 1 hour.

Figure 3.1 Lens Stretching Organ Culture Setup The anterior portion of the eye was isolated, and the cornea and iris removed; leaving the lens, ciliary body, and a ring of sclera intact. Eight flaps were cut into the sclera and each flap was affixed to the silicone disk using staples. The silicone disk was mounted onto the stretching ring which, when expanded, would stretch the lens equally along eight axes. Disk mounted onto stretching ring in unstrained (A) and strained (B) configurations. A close up of the lens in the unstretched (C) and stretched conditions (D). The lens and accessory tissue mounted onto the motorized lens stretching device and submerged in culture media (E). The amplitude and frequency of the lens stretching regime can be customized and run for the entire duration of the tissue culture period.

Lenses were subjected to static stretching at 0% (control), 6%, and 12% strain (defined as

46 percent change in the equatorial diameter of the lens). These strain amplitudes were chosen so as to remain close to the in vivo physiological range186-188. For lenses undergoing cyclic stretching, the stretching ring was mounted onto a motorized rig and stretched at 6% strain amplitude with a frequency of 0 (control), 0.05, 0.1, and 0.2 Hz.

These frequencies were chosen with 0.2 Hz as the highest because previous studies have observed proliferation in other cell types is inhibited at higher frequencies189. The strain amplitude was validated by comparing images of the lenses in the stretched and unstretched configuration in ImageJ (NIH, Bethesda, MD)190.

Flow Cytometry

Lenses were removed from culture, immediately isolated from the surrounding tissue, and rinsed in PBS. The lens capsule was isolated by peeling the capsule open from the posterior pole using jeweler’s forceps then the fiber cell bundle removed. The lens capsules were rinsed twice with PBS and submerged in 0.25% trypsin with 0.04% ethylenediaminetetraacetic acid (EDTA) (VWR, Radnor, PA) at 4°C for 18 hours. Excess trypsin was removed and the capsules were placed in a 37°C water bath for 30 minutes.

Ten milliliters of M199 supplemented with 0.1% BSA was added to quench the trypsin activity. Capsule fragments were filtered using a 70 μm cell strainer and the solution was centrifuged for 10 minutes at 180 rcf to collect the LECs.

The LECs were immediately fixed in 10% neutral buffered formalin (Sigma-Aldrich) and stained for EdU detection with AlexaFluor 488-azide using a Click-iTTM kit for 30 minutes according to manufacturer instructions (Invitrogen, Carlsbad, CA). LECs were also stained for nuclear detection using NucRed TM Live 647 ReadyProbes TM Reagent for 47

30 minutes (Invitrogen). LECs were analyzed using a BD LSR II flow cytometer using the 488 nm and 640 nm lasers (BD Biosciences, Franklin Lakes, NJ). The labelling index was calculated from the percentage of cells that had a positive signal for the Alexa 488 stain using Flowing Software (Turku, Finland).

The resulting raw data were analyzed by collecting measurements from unstained cells to establish autofluorescence thresholds and cells from the same lens stained with both dyes.

The dataset from each stained sample was gated such that the threshold would exclude

95% of the signal from the unstained control.

Fluorescent Microscopy

In order to image the intact LEC monolayer in situ, a flat-mounting technique191 rather than imaging an intact lens192. This avoids the potential complications arising from imaging a curved surface. Lenses were removed from culture, and immediate isolated from surrounding tissue. The intact lens was fixed in 10% neutral buffered formalin for 10 minutes in order prevent sloughing off of the LEC monolayer during flat-mounting.

Lenses were then dissected and flat-mounts of the lens capsule were prepared191. Lens flat-mounts were then fixed in 10% neutral buffered formalin for a further 10 minutes.

The LECs were permeabilized in 0.05% (v/v) Triton X-100 and blocked in 1% (w/v) bovine serum albumin for 30 minutes191. Flat-mounts were rinsed in PBS and then incubated with fluorescent stains. AlexaFluor 488-azide using a Click-iTTM kit (Invitrogen) was used to visualize proliferative activity in the lens. To visualize YAP localization, flat mounts were incubated with primary and secondary antibodies for 60 minutes each.

Primary antibodies included YAP1 Rabbit Polyclonal Antibody (Thermo-Fisher). The secondary antibody used was Goat anti-Rabbit, Alexa Fluor 488 (Thermo-Fisher). For

48 both tests, following incubation with the antibodies and commercially available kits, samples were then counterstained with Hoechst (Thermo-Fisher). Lenses stained for proliferation were imaged using a Nikon Eclipse Ti2-E confocal microscope and those stained for YAP localization were imaged using a Nikon Eclipse Ts2 fluorescent microscope (Nikon Instruments Inc., Melville, NY).

Confocal images were processed to objectively remove background signal, assumed to arise from either autofluorescence or residual dye, as follows. Z-stacks were flattened by keeping the maximum channel intensity value for each pixel location in the horizontal plane. Pixels corresponding to Hoechst-positive nuclei were determined using Otsu’s threshold method to produce a binary mask193 Morphological closing and opening operations were applied to avoid the loss of real connections or development of spurious connections between pixels. Areas with an area >50px were excluded.

EdU-positive nuclei were detected as follows. Autofluorescence or residual dye intensity in the green channel was estimated on the basis of the green channel intensity for all pixels not corresponding to a nucleus (as defined above). An empirical (Kaplan-Meier) cumulative distribution function was determined by including all such pixels. A probability that the mean pixel intensity within a given nucleus was not a result of autofluorescence or residual dyes was then calculated. If this probability exceeded 95%, the corresponding nucleus was considered to be EdU-positive.

Statistical Analysis

A paired t-test was conducted on paired lenses cultured for either 1 or 24 hours to determine whether the labelling indices of LECs, defined as the number of EdU-labeled 49

LECs to the total number of LECs, varied with respect to time spent in culture. Simple linear regression analysis was used to determine the effects of stretch amplitude and frequency on LEC proliferation. Analysis of covariance (ANCOVA) was performed to investigate the effects of verteporfin on LEC proliferation across different stretching conditions. A post-hoc Tukey’s Honestly Significant Difference (HSD) test was used to compare the verteporfin group with the control. Statistical analysis was performed using

JMP Pro 13 (SAS Institute, Cary, SC).

Results

Labelling Index of LECs Varies with Culture Time

The microenvironment LECs experience within the eye in vivo differs from that experienced in vitro. In order to investigate whether the stresses caused by a change in environment affect the labelling index of LECs paired whole lenses mounted on silicon rings and were cultured for either one hour or 24 hours, under null stretch conditions with each lens exposed to EdU for one hour. LECs were then analyzed using flow cytometry to determine labelling index.

A paired t-test was used to determine if the labelling index of LECs cultured for 1 hour immediately following dissection was significantly different than those cultured for 24 hours (Fig. 3.2). A significant difference was found (p=0.0055). The data show that the labelling index of LECs in whole lens cultures decreased with time in culture. The higher initial labelling index followed by a decrease over time may be due to a stress response induced by changing the LEC microenvironment and the subsequent acclimation of the

LECs to the new environment. Longer culture times could therefore be preferable for

50 later studies in order to avoid any initial confounding cellular response to a new microenvironment.

Figure 3.2 Variation of labeling index under null stretch with different times spent in culture A paired t-test showed a significant difference in means (P = 0.0055). The labeling indices of lenses cultured without strain for one hour was higher than those cultured for 24 hours.

Increasing Stretch Amplitude Increases LEC Proliferation in Whole Lens Cultures

To determine if the amplitude of static stretching affected LEC proliferation, a total of 8 pairs of whole lenses were cultured for a period of 24 hours under various static stretch conditions. For each pair, one was subjected to 6% (4 pairs) or 12% (4 pairs) 51 strain while the other (control) was held at 0%. During the final hour of the culture period the lenses were exposed to EdU, which would be incorporated into any newly synthesized DNA. The LECs were analyzed using flow cytometry and the labelling index was calculated from the percentage of the total population of cells that had synthesized new DNA during the hour-long EdU pulse. The labelling index values for the different stretch conditions are presented in Table 1.

Strain Amplitude (%) Labelling Index (%) n

0 1.14 ± 0.37 8

6 3.23 ± 0.27 4

12 6.57 ± 0.46 4

Table 3.1 Labeling Indices of Lenses Cultured Under Varying Static Strain Amplitudes

Simple linear regression analysis was used to determine if stretch amplitude was a significant predictor of LEC proliferation (Fig. 3.3). A significant regression equation was found (Labelling Index = 1.03 % + 0.44 % * Percent Stretch, R2 =0.963, p <0.0001).

The data show that the proliferation of LECs increased proportionally to stretch amplitude. These findings indicate LEC proliferation is driven, at least in part, by mechanotransduction. Therefore, the accommodative process may contribute to the growth of the lens.

Figure 3.3 Variation of labeling index with respect to strain amplitude Linear regression analysis predicted the relationship to follow: Labeling Index = 1.03 + 0.44 × Percent Strain Amplitude (R2 = 0.963, P < 0.0001). Lenses were cultured for 24 hours under varying static strain amplitudes and exposed to a one-hour EdU pulse before LECs were isolated and analyzed using flow cytometry. The labeling index increased proportionally with strain amplitude, suggesting a strong relationship between lens stretching and LEC proliferation.

LEC Proliferation Increases with Stretching Frequency in Whole Lens Cultures

Once a link between LEC proliferation and stretch amplitude was established it was necessary to determine if a change in labelling index occurred in response to changes in stretching frequency as well as stretch amplitude. Whole lens tissues were cultured and analyzed as described above. Stretch amplitude was held at 6% and the triangular stretch waveform was applied cyclically, oscillating from 0% to 6%, at frequencies of 0, 0.05,

0.1, and 0.2 Hz. A total of 16 unpaired lenses were used. The labelling index values are described in Table 2. 53

Strain Frequency (Hz) Labelling Index (%) n

0 3.23 ± 0.27 4

0.05 3.80 ± 0.51 4

0.10 5.52 ± 0.27 4

0.20 7.56 ± 0.23 4

Table 3.2 Labeling Indices of Lenses Cultured Under Varying Cyclic Strain Frequencies

Linear regression analysis was used to determine if LEC labelling index is predicted by changes in stretching frequency (Fig. 3.4). A significant regression was found (Labelling

Index = 3.05 % + 22.62 % * Stretching Frequency (Hz), R2 = 0.954, p<0.0001). Results show that the proliferation of LECs increased proportionally to stretching frequency. As such, lens growth is not only affected by the amplitude but also the frequency of the stretch applied.

Figure 3.4 Variation of labeling index with respect to strain frequency Linear regression analysis predicted the relationship to follow: Labeling Index = 3.05 + 22.62 × Strain Frequency (Hz; R2 = 0.954, P < 0.0001). Lenses were cultured for 24 hours under 6% cyclic strain amplitude at varying strain frequencies with one hour of exposure to EdU. Labeling index was highly correlated to strain frequency.

Stretching Alters the Localization of LEC Proliferation Across the Lens Capsule

Qualitative analysis of the effects of different stretching regimes on the localization of LEC proliferative activity was performed by staining flat-mounted lens capsules for the thymine analog, EdU, and counterstaining LEC with the nuclear stain

Hoechst. Representative images of lenses cultured under null strain (A), 12% static stretch (B), and cyclic stretch at 6% amplitude and 0.20 Hz (C) were used (Fig. 3.5). Low levels of EdU staining were observed in the null stretch lens (A), primarily near the equator. The static stretch lens appeared to have a higher labeling index (B), with the

55 majority of EdU staining also occurring near the equator. In the cyclic lens, EdU labeling was observed near the anterior pole (C) and at different points along the equator (D-F).

Figure 3.5 Variation in EdU labeling localization under different strain conditions Qualitative analysis of representative images of lenses cultured under null strain (A), static strain (B), and cyclic strain (C–F); and stained for the EdU proliferative marker (green). The null (A) and static (B) stretch lenses are shown as mosaics going from one side of the equator to the other. The cyclic stretch lens (C–F) shows discrete images taken at different points along the anterior capsule: the anterior pole (C) and points along the equator (D–F). Lenses cultured under null strain (A) showed little reactivity with the EdU stain. In static strain (B), and cyclic strain (C) lenses, EdU labeling was primarily in the germinative zones (GZ). Magnified images of the GZ of null (A), static (B), and cyclic (F) stretched lenses are shown.

Inhibition of YAP Function Blocks Mechanotransductive Effects of Stretching on LEC Proliferation

The effects of the YAP function inhibitor verteporfin on LEC proliferation was determined by exposing both paired eyes to identical static stretching conditions: 0%,

6%, or 12%. Both members of each pair were cultured in enhanced M199 and the treatment group was supplemented with verteporfin. A total of 10 pairs of eyes were used with 4 pairs of eyes cultured under null stretch, and 3 pairs each with 6% and 12% static strain. The labelling index values are described in Table 3.

Strain Amplitude (%) Labelling Index (%) n

Verteporfin Control

0 0.925 ± 0.24 1.21 ± 0.10 4

6 1.27 ± 0.20 3.86 ± 0.21 4

12 1.14 ± 0.14 5.92 ± 0.59 4

Table 3.3 Labeling Indices of Verteporfin Treated and Paired Control Lenses Cultured Under Varying Static Strain Amplitudes

Linear regression analysis was performed on both treatment groups. A significant correlation was found for the control group (Labelling Index = 1.28 % + 0.39 % * Stretch

Amplitude, R2 = 0.975, p<0.0001), but no significant relationship was found for the group treated with verteporfin (Labelling Index = 0.99 % + 0.02 % * Stretch Amplitude,

R2 = 0.292, p = 0.107) (Fig. 3.6). To confirm the difference between the treatment groups, ANCOVA was used to determine whether there was a significant difference 58 between groups; a significant difference was observed (F=282.34, p<0.0001). A post-hoc

Tukey’s HSD test indicated a significant difference between the regression lines of the verteporfin and control groups (p <0.0001).

Figure 3.6 Variation of labeling index with respect to strain amplitude of lenses treated with a YAP inhibitor, verteporfin, and their paired controls Linear regression analysis predicted the relationships to follow: Labeling Index (%) = 0.99 + 0.02 × Strain Amplitude (R2 = 0.292, P = 0.107) and Labeling Index = 1.28 + 0.39 × Strain Amplitude (R2 = 0.975, P < 0.0001) for the treatment and control groups, respectively. Lenses were cultured for 24 hours under varying static strain amplitudes. After a 1-hour EdU pulse, LECs were analyzed using flow cytometry. The labeling index of the control group increased with static strain amplitude while the group treated with a YAP inhibitor showed no statistically significant relationship.

Analysis of the effects of YAP function inhibition on cyclic stretch proliferative response 59 was also performed by culturing lenses at 6% stretch amplitude at 0.20 Hz with a verteporfin treated and untreated paired control (Fig. 3.7). The mean labelling index of the untreated control was 6.30±1.58. The verteporfin treated group had a mean labelling index of 1.30±0.33. A paired t-test showed a significant difference between the means

(p<0.0001).

Figure 3.7 Labeling index of cyclically stretched lenses treated with and without verteporfin Comparison of LEC labeling index of verteporfin and control lenses under a cyclic strain (6% strain amplitude, 0.20 Hz) regime. The control group had a mean labeling index of 6.30% ± 1.58%. The verteporfin treated group had a mean labeling index of 1.30% ± 0.33%. The results of a paired t-test had a P < 0.0001.

The data shows that when YAP function was inhibited by verteporfin, the correlation between LEC labelling index and stretch amplitude was effectively eliminated. This suggests that YAP plays a crucial role in the transduction of mechanical signals into an upregulation of LEC proliferation. Furthermore, when YAP function is inhibited those signals are blocked and LEC proliferation does not increase with mechanical stretching.

Stretching Alters YAP Localization in LECs

Qualitative analysis of the effects of YAP function inhibition by verteporfin on the localization of intracellular YAP was performed by staining flat-mounted lens capsules for YAP, and counterstaining them with the nuclear stain Hoechst. Paired verteporfin treated and untreated control lenses were cultured under static and null stretch conditions and were then stained and imaged using fluorescent microscopy (Fig. 3.8). All images were taken in the germinative zone near the equator of the lens. YAP nuclear localization was observed in the static stretched untreated lenses (C), suggesting the activation of YAP and translocation into the nucleus. YAP nuclear localization was not observed in the YAP function inhibited static stretched lens (F). YAP activation was not observed in both the null stretched untreated (I) and treated (L) lenses. The staining and microscopy techniques used were only able to visualize the nuclear YAP, this was potentially due to the more diffuse localization of cytoplasmic YAP being unable to generate a strong enough signal to be visualized over the autoflourescent background.

These results suggest YAP can be activated by stretch in LECs and that its function can be inhibited by verteporfin.

Figure 3.8 Localization of YAP in lenses cultured with and without verteporfin under static and null stretch conditions Lenses were stained with the nuclear stain Hoechst (A, D, G, J) and for YAP (B, E, H, K). Nuclear localization of YAP was observed in the statically stretched untreated lens (C). This nuclear localization response was not observed in the statically stretched lens treated with verteporfin (F). YAP nuclear localization was also not detected in the untreated (I) or verteporfin treated (L) lenses cultured under null stretch conditions.

Discussion

This study was designed to determine if radially stretching the lens had an effect on LEC proliferation and whether YAP was involved in the mechanotransductive signaling pathway driving stretch-induced LEC proliferation. The stretch response was found to be dependent on both stretching amplitude and frequency; qualitative analysis of the localization of LEC proliferative activity showed differences between static and cyclic stretch. YAP was found to play an important role in the signaling pathway.

These results have important implications for understanding lens growth and morphogenesis, as well as approaches for modulating LEC proliferation. Controlling

LEC proliferation could allow for retarding lens growth as a means for delaying presbyopia or cataract, as well as prevention of PCO (i.e. regeneration of the lens material following cataract surgery). Our data suggest that behavioral, environmental, and therapeutic approaches may be feasible for limiting or encouraging lens growth.

The human lens continues to grow throughout life, with an apparent bi-phasic growth pattern194. It may be that the initial, very rapid, pre-natal growth phase is driven by a rapid increase in lens capsule surface area and constant stretching forces, whereas the later, much slower, growth phase is retarded by the partial relief of lens stretching during accommodation. This is supported by Augusteyn’s observation that the transition between growth phases occurs near the time of birth, as does the ability to accommodate194. In non- or minimally-accommodating species, age-matched lenses tend to be much larger (e.g. a six-month-old pig lens may be ~400 mg, whereas an infant

63 human lens is ~150 mg), possibly due to persistent disaccommodation.

Earlier studies have identified YAP as playing an important role in the regulation of tissue size, including the lens49, 50, 65. When YAP is unphosphorylated and active, it is localized in the nucleus, acting as a transcriptional coactivator promoting the expression of genes inducing cell proliferation, survival, and migration195. YAP is primarily regulated by the Hippo signaling pathway, which when activated, phosphorylates YAP and inhibits its activity. The Hippo pathway is regulated by various mechanisms, including cell-cell contact, cell polarity, cellular energy status, hormonal signals, and, most relevant to this study, mechanical cues65, 195, 196. Studies on other cell types have observed that Hippo pathway activity is downregulated by stretching of those cells via the phosphorylation of the LATS1 kinase, which is the primary negative regulator of

YAP65, 197. While the role of YAP in mechanosensing and cell proliferation has been studied in other cell types65, 195-197, and its expression in the lens is well documented49, 50, this study is the first establish the link between mechanical cues and the regulation of the

YAP protein in the lens.

The data presented in this study support the hypothesis that stretching the lens results in the activation of YAP and a subsequent increase in proliferative activity. However, YAP regulation is controlled by several different pathways with a significant amount of crosstalk between them and several other mechanosensing pathways may also be at play198, 199. In addition to the Hippo signaling pathway, RHO and MAPK/ERK signaling have also been implicated in the mechanoregulation of YAP198, 200-203. In other tissue

64 types the signaling pathways Wnt, TGF-β, and Notch have been also been shown to increase cell proliferation in reaction to shear stress without the mediation of YAP199.

Additionally, p38 and JNK signaling pathways have been reported to respond to stretch and increase cell proliferation204, 205. Further studies will be necessary to determine what role the different mechanotransduction pathways play in the regulation of LEC growth.

This study demonstrates that stretching the porcine lens and connective tissues ex vivo results in increased LEC proliferation. There are several factors which could modulate

LEC behavior during stretching. First, the capsule experiences increased tension due to the increase of zonular tension; this increase in capsule surface area will necessarily increase the footprint of LECs. LECs will also presumably experience increased apical pressure from the fiber cell bundle in the stretched state which could vary with position.

LEC-LEC tensile forces may increase as well. Stretching could also drive fluid flow in and out of the lens or drive an increased rate of transport via convection due to relative motion of the capsule to the surrounding media206, 207. Finally, it is also possible that alternate signaling molecule(s) are activated or transported due to the stretching motion.

Further investigation is required to pinpoint the underlying mechanism(s) of the stretch- induced change in proliferation.

The microscopy results presented in this study suggest increased labeling in cyclic and static stretch lenses compared to static lenses. The labeling indices measured using flow cytometry for the different stretch conditions were in qualitative agreement with microscopy findings. When visualizing the distribution of proliferative activity, the

65 majority of EdU staining was observed at the GZ (Fig. 3.5). Future work will quantitatively assess spatial variations in proliferation.

Fluorescent microscopy showed activated nuclear YAP in LECs in the GZ (Fig. 3.8).

This activation of YAP was only observed in stretched lenses uninhibited with verteporfin; YAP activation was observed in neither the unstretched lenses nor lenses treated with verteporfin. In some regions of the GZ of the statically stretched, EdU stained LEC proliferative activity (Fig. 3.5 B, E) was observed in a similar density as the static stretched lens stained for YAP nuclear localization (Fig. 3.8 B); suggesting a correlation between YAP nuclear localization and DNA synthesis. The microscopy technique used was unable to visualize cytoplasmic YAP and as a result provides no information on the total YAP content of the LECs. Additionally, the distribution of YAP nuclear localization likely changes depending on the position on lens capsule, and potentially corresponds to the local strain profile of the LECs. Further studies are needed to more completely characterize the behavior of YAP in response to stretching the lens.

While mechanotransduction pathways are highly conserved and every effort was made to replicate the microenvironment of the lens in vitro, caution should be used in extrapolating these findings to predict the in vivo behavior of human LECs. We kept the lens and connective tissues intact and retained the anterior vitreous attached to the lens.

Still, the conditions experienced by the lens during the study differed from those in vivo.

For example, cell culture media composition, including oxygen content, could alter the magnitude of the change in proliferation that was observed. Stretch amplitudes were

66 chosen to match physiological extents of stretching: the maximal stretch-induced change in equatorial radius in a young human lens has been found to be between 5-10%62, 208-210.

In the present study, porcine lenses were used, which have different geometric and mechanical properties when compared to human lenses11, 58, 209, 211. Therefore, the distribution of mechanical stresses and the deformation from stretching likely differs between species. Additionally, pigs do not accommodate, resulting in a smaller and less robust ciliary muscle than that found in a human or primate eye212. However, the biomolecular composition and crystallin distribution in both the human and porcine lenses are very similar213, 214. Further, mechanotransduction signaling and gene expression are highly conserved between species195. Thus, if the LECs in the porcine lens have an increased rate of proliferation in response to mechanical stretch, a similar response, albeit possibly with a different magnitude, would likely be observed in humans.

There are several possible explanations for the higher labelling index immediately after dissection relative to 24 hours of culturing after dissection. A stress response induced by changing the LEC microenvironment, subsequent acclimation of the LECs to the new environment, strain acting on the porcine lens in vivo and post mortem, and mechanical stimulation during the dissection process may contribute to the difference in labelling indices59. Longer culture times could therefore be preferable for later studies in order to avoid any initial confounding cellular response to a new microenvironment or effects from mechanical loading prior to the culture period.

The results of the study establish a link between mechanical stretching and the

67 upregulation of LEC proliferative activity, as well as identifying a target which, when inhibited, would reduce the growth of the lens. Overall this study provides new insights into the processes controlling lens growth and opens novel avenues by which to study the etiology of age-related vision disorders of the lens. Future work will map the localized proliferation changes with corresponding mechanical strains in the capsule.

Chapter 4. Changes in the Spatial Distribution of Lens Epithelial Cell Proliferation Resulting from Lens Stretching

This chapter has been submitted for publication.

Introduction

The lens plays a crucial role in the process of accommodation, by which the eye is able to alter its focal distance. The lens continuously grows in size throughout the lifetime, unlike the globe which maintains a constant size from adulthood1, 2. The growth of the lens is driven by the proliferation and subsequent differentiation of the lens epithelial cells (LECs) into lens fiber cells3-5. Recent work has found LEC proliferative behavior to be mechanosensitive37 The volume of lens fiber cells is larger than that of a

LEC, thus the increase of the total volume of the lens is further impacted by each instance of LEC proliferation and differentiation6. The relationship between lens size and stiffness and the loss of accommodative ability with age is not fully understood11, 12. However, these changes in size and stiffness may contribute to the onset of presbyopia and cataracts7-11. Understanding the factors influencing the proliferation of LECs is crucial in opening avenues of research for developing treatments for posterior capsule opacification

(PCO) and for translating lens regeneration into the clinic13-16.

Presbyopia is the progressive loss of accommodative ability with age and is potentially linked to the growth of the lens. When the lens is disaccommodated to focus on distant

69 objects, the ciliary muscle is relaxed and has a larger diameter, leading to the lens being stretched due to tension applied by the zonules17-20. When the lens accommodates to shift the focus to nearby objects, the ciliary muscle contracts, the tension on the zonules releases and the lens elastically recoils to its more relaxed state. With age the amplitude of accommodation decreases until it becomes difficult to focus on nearby objects, a condition known as presbyopia. The diameter and force of contraction of the ciliary muscle,21-26 as well as the mechanical properties of the zonules have also been found to be largely independent of age27. This suggests that the etiology of presbyopia originates within the lens.

In humans, the lens follows a biphasic growth pattern where the lens grows rapidly in the first stage of life, after which growth slows and increases linearly with about 1.38 mg of lens material being added each year5, 28-30. As the lens grows, the accommodated diameter increases with age, while the disaccommodated diameter remains constant31. Therefore, the change in lens diameter between the disaccommodated and accommodated state decreases with age. As the size and shape of the lens change with age, the bulk stiffness of the lens also increases32-35. It can be hypothesized that these changes contribute to the onset of age-related refractive errors, in which case defining the relationship between the forces acting upon the lens during accommodation and the growth of the lens is important in understanding the etiology of presbyopia.

The driving forces governing lens growth are not yet fully understood. Technical challenges in reproducing the in vivo environment of the lens have long prevented the

70 study of the effects of biochemical and biomechanical factors upon lens growth. Lens flat mount explants often result in altered LEC morphology leading them to be inappropriate in studying the mechanobiology of the lens56. Additionally, animal models that employ a human-like mechanism of accommodation tend to be prohibitively expensive. Recent work has found success in using an in vitro stretching device to culture whole porcine lenses to study the effects of mechanical stimulation on the growth of the lens37. By culturing whole lenses, the biomechanical microenvironment is preserved and the response of the LECs to stretching likely mirrors the response that would have been observed in vivo. It has been established that LEC proliferation is impacted by stretch and that both the magnitude of stretching and the frequency of a cyclic stretching stimulus does impact the growth rate of the LECs37. The relationship between LEC proliferation and both the frequency and magnitude of stretching was found to be proportional37.

However, the quantification of LEC proliferation was accomplished using flow cytometry, and as a result all spatial resolution was lost; therefore, it is unknown what effect, if any, stretching had on the localization of LEC proliferative events.

The spatial distribution of LEC proliferation has been previously studies in non- proliferating species such as mice, rats, cows, and rabbits2, 5, 30, 45, 216-218. In those species,

LEC proliferation was primarily contained in the GZ. In a non-accommodating lens, the strains acting upon the lens remain relatively stable, with the local capsule strains being higher near the equator, in the GZ, than anywhere else in the lens11, 12, 209. In an accommodating species, the localized strains along the lens capsule are more dynamic, and may result in a difference in the spatial distribution of LEC proliferation. 71

This study was designed to determine the localization of proliferative activity along the capsule in lenses subjected to different types of stretching regimes: null, static, and cyclic. A lens organ culture bioreactor was employed to mechanically stimulate the lenses ex vivo37. And confocal microscopy was used on the fluorescently labeled lenses to determine the positional information of proliferative activity.

Methods

All animal tissues were used in accordance with institutionally approved protocols.

Lens Stretching Organ Culture

Eye dissections and lens stretching was performed based on a previously published procedure37. Porcine eyes were obtained from a nearby abattoir (Bay Packing,

Lancaster, OH) after being freshly enucleated. Extra-ocular tissue was removed using scissors allowing the whole globe to be submerged in a solution of 0.5% povidone-iodine

(Sigma-Aldrich, St. Louis, MO) in phosphate buffered saline (PBS) for a period of 5 minutes in order to be disinfected. Following disinfection, the globe was transferred into

PBS. The globe was removed from PBS and small incisions were made in the limbus and equator of the globe using a scalpel. Using a pair of micro scissors, the cornea and iris were carefully removed in order to prevent dragging along the lens during stretching. The globe was then bisected using a pair of dissection scissors and the posterior half of the globe was discarded. The vitreous was cut in half, leaving the anterior portion of the vitreous attached to the lens and ciliary body and the posterior vitreous was discarded.

This resulted in a portion of tissue consisting of the lens still attached to the ciliary body

72 via the zonules, and the anterior section of the vitreous, and a ring of sclera surrounding it.

To mount the lens onto the stretching device, eight radial cuts were made 45° apart to create eight flaps surrounding the lens. To ensure uniform stretching, the lens and surrounding tissue were attached to a 40 mm diameter silicone disk with a 10 mm diameter hole placed in the center, with the lens placed over the central hole. The silicone disk had eight equally spaced peripheral holes to facilitate mounting onto the stretching ring device. Each flap that was made in the sclera was attached to the silicone disk using a staple such that there was one flap affixed between every two peripheral holes. The silicone disks were attached to the stretching ring using a screw going through the peripheral holes (Fig. 3.1)37.

The lenses mounted to the stretching device were submerged in pre-warmed, serum-free medium 199 (M199) with Earle’s salts L-glutamine, and sodium bicarbonate, supplemented with 0.1% bovine serum albumin, 100 IU/ml penicillin, 100 mg/ml streptomycin and 2.5 mg/ml Amphotercin B (Sigma-Aldrich). Lens tissue was incubated at 37°C, 5% CO2 for 18 hours.

Lenses were subjected to one of three stretching regimes throughout the culture period: null stretching, 12% static stretch, and 6% cyclic stretch at 0.2 Hz. The stretching magnitude was defined as percent change in the equatorial diameter in the lens and were chosen to remain close to the in vivo physiological range37, 186-188. The stretch magnitude was validated by comparing images of stretched and unstretched lenses in ImageJ (NIH,

Bethesda, MD)190. The cyclic stretching was achieved using a motorized rig to manipulate the stretching ring moving the lens from 0% to 6% stretch along a triangular waveform. The frequency of 0.2 Hz was chosen as previous work found that stretching the lens at that frequency has an effect on LEC proliferation37.

Confocal Microscopy

In order to preserve spatial information of proliferative activity of the lens, the

LEC monolayer was imaged in situ. Due to the large size and radius of curvature, a flat- mounting technique was used37, 217, 219 rather than imaging the intact lens126. After the culture period ended, lenses were removed from the stretching device and immediately isolated from the surrounding tissue and rinsed in PBS. Intact lenses were fixed in 10% neutral buffered formalin for 10 minutes in order to prevent the LEC monolayer from sloughing off of the capsule. Lenses were then rinsed in PBS and flat-mounted by making a small incision in the posterior pole using a scalpel and then using micro scissors to make 6 pie cuts from the posterior pole to the equator. After which the fiber cell bundle was removed. The lens flat mounts were then fixed in formalin for an additional 10 minutes. The lens capsules were rinsed in PBS and then submerged in a blocking solution of PBS containing 0.5% (v/v) triton X-100, 1% (w/v) bovine serum albumin (Sigma-

Aldrich), and 1% (v/v) goat serum (Thermo-Fisher, San Jose, CA) for 1 hour. The blocked flat-mount lenses were then incubated with the primary antibody overnight at 4°

C for 18 hours by submerging the lens capsule in a staining buffer containing the primary antibody, 0.5% (v/v) triton X-100, 1% (w/v) bovine serum albumin (Sigma-Aldrich), and

5% (v/v) goat serum (Thermo-Fisher) in PBS. The final dilution for the monoclonal rabbit anti-Ki67 primary antibody was 1:50 in staining buffer (Thermo-Fisher).

The lens capsules were removed from the staining buffer and rinsed in fresh blocking solution three times. The lenses were then washed in blocking solution for 2 hours at room temperature. After which the lenses were incubated with the secondary antibody in staining buffer at room temperature for 1 hour. The dilution of the goat anti-rabbit highly cross-adsorbed secondary antibody conjugated with Alexa-488 was 1:250 in staining buffer. The lens capsule was again removed from the staining buffer and rinsed in fresh blocking solution three times and washed in blocking solution for 1 hour. After which the lenses were counterstained with Hoechst.

Lenses were imaged using an Olympus FV300 (Olympus, Tokyo, Japan) confocal microscope. For mosaics of lens proliferative activity, a 10x objective was used.

Confocal images were processed to objectively remove background signal. Z-stacks were flattened by keeping the maximum channel intensity value for each pixel location.

Hoechst-positive nuclei were determined using Otsu’s threshold method to produce a binary mask193. Ki-67-positive nuclei were by estimating autofluorescence in the green channel on the basis of the green channel intensity for all pixels not corresponding to a nucleus (as defined above). An empirical (Kaplan-Meier) cumulative distribution function was determined by including all such pixels. A probability that the mean pixel intensity within a given nucleus was not a result of autofluorescence or residual dyes was then calculated. If this probability exceeded 95%, the corresponding nucleus was

75 considered to be Ki-67-positive37.

Computational Modeling Lens Capsule Strain

A computational model of a six-month-old pig lens was constructed in COMSOL

Multiphysics v5.5 (COMSOL Inc., Burlington, MA). Figure 4.1a shows the geometry represented by two oblate spheroids connected at the equator with dimensions of the lens taken from literature (equatorial radius: 5.02mm; anterior minor radius 2.46mm; posterior minor radius 5.41mm)58. The same photo was used to measure the upper and lower boundaries of the zonular insertions to the lens, beginning 0.515mm posterior to the equator and extending to 0.992mm anterior to the equator.

Figure 4.1 Model geometry (a) and meshed model (b) of the porcine line Blue nodes indicate the boundaries of zonular attachments which were modeled as linear springs

The lens fiber cells were treated as an incompressible hydrogel which was subjected to swelling encapsulated by a membrane. Both the membrane and lens fibers were modeled as isotropic, neo-Hookean materials with properties obtained from the literature (lens fibers: 퐸(푃푎) = 1952푒−0.268푑 where d is the distance from the center of the lens (mm), assumed incompressible211; capsule: E=260 kPa220, =0.47221, thickness=57µm). The lens capsule was represented by 203 quartic Lagrange membrane elements, while the lens fibers were represented by 1999 triangular elements (Fig. 4.1b).

The effects of lens growth on strains within the capsule were estimated by simulating 77 growth as volumetric swelling using the Hygroscopic Swelling node in COMSOL, producing a uniform volumetric strain of 8%. This growth state was assumed to represent the residual loading on the lens capsule. The effect of stretching was then computed by applying a zonular load related to the radial and axial components estimated for the human lens by Hermans et al222. Specifically, the ratios of each force vector component was proportional to those found by Hermans et al.222, with the magnitude scaled until the equatorial displacement increased by 12%. Additionally, these forces were distributed over the zonular attachment area according to the superposition of three Gaussian distributions roughly corresponding to the nominal groupings of anterior, equatorial, and posterior zonules (Fig. 4.2). This distribution of forces avoided singularities arising from sharp stress transitions.

Figure 4.2 Components of the stretching force

The local capsule strains were then computed for the growth state (G) and the growth + stretching state (GS). We assumed that LECs were in homeostasis in state G, and that this state corresponded to the null loading condition in the lens culture experiments. The local strain energy density W within the lens epithelium was calculated assuming it behaves as an incompressible, isotropic, neo-Hookean layer and that it stretched in the meridional/circumferential directions by the same amount as the underlying capsule.

Thus, W was given by: 79

휇 2 2 1 푊 = (휆푚 + 휆푐 + 2 2 − 3), (1) 2 휆푚휆푐

where 휆푚 and 휆푐 are respectively the local meridional and circumferential stretch ratios in the capsule, and µ is the shear modulus of the epithelium (arbitrarily selected as 100

Pa). Note that the stretch ratios 휆푚 and 휆푐 were computed on the basis of stretch-induced changes in local strains, i.e. 휆푚 = √2Ε푚 + 1 and 휆푐 = √2Ε푐 + 1, where Ε푚 and Ε푐 are the local Green strain values in state GS relative to those in state S (e.g. Ε푚 = Ε푚,퐺푆 −

Ε푚,퐺). This difference was used to account for the assumption that LECs were in homeostasis in state G prior to the application of stretching.

Statistical Analysis

To determine whether the total labeling indices of lenses in different stretching groups were significantly different a Welch’s ANOVA was used along with a post-hoc

Games-Howell test to compare the groups due to the unequal variances found between the stretching regimes. To compare the labeling indices between the regions of each of the stretching groups, a nonparametric Kruskal-Wallis test and post-hoc Wilcoxon comparison was used. A cumulative distribution function for the proliferative events for the cyclically and statically stretched lenses was created and a non-parametric

Kolmogorov-Smirnov two-sample test was employed to determine if they were statistically significantly different. Correlation between the local LEC Ki67 labeling index and the computational model-predicted local epithelial strain energy density was

80 also evaluated. Statistical analysis was performed using Matlab (MathWorks, Natick,

MA) and JMP Pro 13 (SAS Institute, Cary, SC).

Results

Labeling Index of Proliferative Activity across Whole Lens Capsules

The labeling index, defined here as the percentage total LECs that were labeled with the anti-Ki-67 antibody, of a whole lens capsule was determined by imaging a strip of lens capsule starting at the equator, passing through the anterior pole, and ending at the antipodal region of the equator. The percentage of the total number of LECs that were positively labeled with the Ki-67 antibody was calculated for each lens subjected to each of the 3 strain regimes (3 lenses per group) (Fig. 4.3). Welch’s ANOVA was used to compare the means of the 3 groups and found a significant difference (F=13.89, p=

0.0204). A post-hoc Games-Howell test was performed and found the statically stretched lenses to be significantly different from the null lenses (p=0.0175). Despite having a higher calculated mean than the null lenses, the variability in the total labeling index cyclic in cyclic lenses resulted in the differences not being statistically significant.

Figure 4.3 Variation of total labeling indices of lenses subjected to different strain regimes. The total labeling indices of lenses subjected to the different stretching regimes: Null (n=3), 12% static (n=3), and 6% cyclic at 0.20 Hz (n=3). A Welch’s ANOVA and post- hoc Games-Howell test were performed and found a significant difference in the labeling indices between the static and null lenses (p=0.0175).

Localization of Proliferative Activity in Lenses Subjected to Different Stretching Regimes

Lenses were flat-mounted and incubated with anti-ki-67 primary and Alexa 488 conjugated secondary antibodies as well as stained with the general nuclear stain

Hoechst. A strip of the lens capsule was imaged using a confocal microscope and the image was analyzed by dividing the total length of the capsule into ten regions based on their distance from the anterior pole in terms of the percentage of the total arc length of the lens (i.e. 0-10%, 10-20%, 20-30%, etc.).The labeling index within each region was calculated for each lens. 82

The lenses were subjected to one of three stretching regimes: null, 12% static stretch, and

6% cyclic stretch at 0.2 Hz. Three lenses were imaged in each group and the localized labeling indices were compared (Fig. 4.4). A Kruskal-Wallis non-parametric test was used to compare the localized labeling indices by group and found a significant difference (H=21.62, p<0.0001). A post-hoc Wilcoxon comparison found the cyclically stretched lens to be significantly different from the static (p=0.0006) and null lenses

(p<0.0001). The static lenses were not found to have a statistically significant difference compared to the null strain lenses, likely due to the similar labeling indices in regions of the lens outside of the germinative zone (Fig. 4.4g).

Figure 4.4 Representative mosaic confocal microscopy images of null (A), static (B), and cyclic (C) lenses, as well as the localized labeling indices of regions of null (n=3) (D), static (n=3) (E), and cyclic (n=3) (F) lenses and the comparison of the localized labeling indices of the three stretching regimes (G)

The LECs were labeled with the Hoechst general nuclear stain (gray), and for Ki-67 (green). A Kruskal-Wallis non-parametric test and a post-hoc Wilcoxon comparison found the cyclically stretched lens to be significantly different from the static (p=0.0006) and null lenses (p<0.0001).

Differences in LEC Labeling Index Distribution in Response to Differing Strain Regimes

In order to determine if the distribution of proliferative events changed between the static and cyclic strain regimes, the location of each proliferative event imaged as a function of distance in pixels from the anterior pole was calculated and plotted and used to calculate a cumulative distribution function (CDF) for the two strain regimes (Fig.

4.5). The two CDFs were compared using a non-parametric Kolmogorov-Smirnov two- sample test, and a significant difference was found (p<0.0001).

Figure 4.5 Cumulative distribution function of Ki-67 labeling as a function of distance from the anterior pole in pixels in lenses subjected to different stretching regimes A Kolmogorov-Smirnov test was used to compare the distributions of (n=3) (solid line) and cyclic (n=3) (dashed line) lenses and found a significant difference in distributions (p<0.0001)

Analysis of Lens Capsule Strains and Proliferation Distribution

Figure 4.6 : Meridional strains (left) and circumferential strains (right) as a function of arc length Blue: residually loaded capsule (i.e. stresses due to growth only); black: strains including growth + 12% stretching; red: strain assumed to be experienced by LECs during one stretching cycle (black line minus blue line)

A model of localized lens capsule strains on a statically stretched lens was created in COMSOL. The strains in the meridional and circumferential directions were calculated as a function of distance from the anterior pole for both model states (Fig. 4.6). The epithelial strain energy density was then computed using Eqn. 1. The log of the resulting local strain energy density was found to strongly correlate with the local Ki67 labeling index of statically stretched lenses (Fig. 4.7a; R2=0.914; p<0.0001). This suggests a potential relationship between local capsule strains and LEC proliferation. Additionally, it is possible that the high local strain at the periphery of the lens may play a role in the

GZ being the region of highest LEC proliferative activity in non-accommodating species.

Figure 4.7 Comparison of Ki67 labeling in whole porcine lenses stretched uniformly around the circumference, resulting in an increase in equatorial diameter of 12% (left). Averaging strain anisotropy and proliferation in 10 equally-spaced increments of arc length between the anterior pole and equator indicated a significant log-linear correlation (n=2; p<0.0001) (right).

Discussion

This study was designed to determine whether radially stretching the lens altered the spatial distribution of proliferating cells in the lens. The response of the LECs is highly sensitive to the type of stretching stimulation applied to the lens (Fig. 4.4).

Unstretched lenses appeared to have a low baseline level of proliferative activity evenly spread across the whole lens capsule. Statically stretched lenses had a similar baseline of proliferative activity across the lens along with a large peak of proliferative activity near the equator in the GZ. Cyclically stretched lenses did not have a large peak in the GZ but instead had elevated levels of proliferative activity spread across the lens capsule. The total labeling indices found in this study (Fig. 4.3) as well as previous studies measuring the labeling index using flow cytometry37 found there to be no statistically significant difference in the labeling of 12% static stretched lenses and 6% cyclic stretched lenses at 87

0.20 Hz. However, the spatial distribution of proliferative activity between the two groups were significantly different (Fig. 4.5).

It is not possible to directly compare the flow cytometry results from previous studies37 of the proliferation response due to stretching and the results obtained in this study. Earlier studies used an hour long EdU pulse to measure LEC proliferation by quantifying the number of LECs containing EdU in newly synthesized DNA using flow cytometry37.

This study uses Ki-67 as a proliferative marker instead of EdU, and employs confocal microscopy rather than flow cytometry. During mitosis, DNA synthesis occurs during the

S-phase, and it is only while the LEC is in the S-phase that EdU can be incorporated during the one hour EdU pulse37, 223, 224. Ki-67 is present in the cell during all stages of mitosis, and is absent in resting cells in the G0 phase225; however, the LEC needs to be in the active phases of the cell cycle (G1, S, G2, and M) at the time of fixation to be positively labelled for Ki-67. In addition to the differences in time periods where the

LECs can be positively labelled, the methodologies used to quantify the labelling index likely have different sensitivities to fluorescence making comparing the results of this and previous studies difficult226. In order to mathematically model the relationship between the two labelling methodologies, it would be necessary to co-stain lenses for both Ki-67 and EdU and use both flow cytometry and confocal microscopy to define the relationship.

Together, these findings suggest that LECs are sensitive to the changes in local capsule strains arising from zonular tension. This suggests that the spatial distribution of LEC proliferation in accommodating species may differ from the non-accommodating species

88 previously used as animal models of lens growth4, 45, 218. The proposed computational approach for determining cell behavior is novel in the field of mechanobiology. Since mechanotransduction pathways are highly conserved227, the combination of these data and a computational model of lens growth and stretching may allow construction of species-dependent models of lens growth.

We have previously observed that non-accommodating species tend to have more spherical lenses than do accommodating species11, leading us to hypothesize that the dynamic forces experienced by the epithelium during accommodation may contribute to altered lens geometry.

The results of this study establish a link between mechanically stretching the lens and the localization of LEC proliferation. Overall this study provides insights into the processes governing the growth of the lens and the information obtained on the localization of proliferation along the lens capsule will prove invaluable in modeling the growth of the lens and studying age-related refractive issues of the lens.

Chapter 5. Conclusions and Future Directions

While extensive study has been made regarding the cell and molecular biology of the lens, all prior studies have neglected the importance of mechanobiological effects.

This has been necessary due to technical challenges in (i) maintaining the viability of the lens and connective tissues, and (ii) applying physiologically relevant biomechanical stimuli to the lens. Accommodation is a fundamentally biomechanical process involving not only the lens but its connective tissues (zonules and ciliary body). In this study, we overcame both of these technical challenges to enable long-term culture of the lens in a bioreactor mimicking the biochemical and biomechanical milieu of the in vivo human lens, including cyclic zonular tension relevant to accommodation.

This project aimed to determine the mechanobiological response of the lens to stretching.

To accomplish this goal, novel techniques were developed to culture whole lens organs ex vivo and a whole lens organ culture bioreactor was designed to mechanically stimulate the lens in vitro. For the first time, lens epithelial cells (LECs) were found to be mechanosensitive. The proliferative activity of LECs increased proportionally to the strain magnitude applied to the lens, and when the lens was stretched cyclically, the proliferation rate increased proportionally to the frequency. This mechanosensitive response was found to be mediated by the YAP signaling pathway. When YAP was inhibited, the proliferative response was decoupled from the mechanical forces applied by 90 stretching.

Further studies revealed that not only did the proliferation rate of LECs increase in response to stretching, but that the localization of proliferative activity was dependent on the type of stretching that was used. Statically stretched lenses had a low level of proliferative activity across the capsule but had high peaks near the equatorial region, where the local capsule strains were highest. This distribution was drastically altered in cyclically stretched lenses, where the proliferative activity was elevated across the entire capsule.

The work presented in this study is foundational in developing the field of lens mechanobiology, as it is the first time LECs were observed to be mechanosensitive.

These findings open a new avenue of research on the lens, which would have possible impacts on studying the etiology of age-related refractive issues involving the lens, namely presbyopia, and cataracts. Additionally, characterizing the forces governing LEC proliferation is necessary for developing therapies for preventing posterior capsule opacification (PCO) after cataract extraction. Furthermore, understanding and eventually modulating LEC proliferation has the potential of opening the door for bringing research on lens regeneration into the clinic by inducing the LECs to proliferate and differentiate in such a way as to regenerate a transparent lens after cataract surgery. This study represents a first step towards that ultimate goal.

This basic science study is somewhat limited in the scope in which the results can be applied to lenses in vivo. The porcine eyes used were all from young subjects due to

91 originating from an abattoir, therefore it is unknown what influence the age of the animal has on LEC proliferation and mechanotransduction. Additionally the porcine lens capsule is stiffer and thicker than a human capsule58, 228. The contribution of the mechanical properties of the capsule to mechanotransductive signaling in the lens are unknown. The lens is also highly unlikely to be constantly disaccommodated or cyclically undergoing constant accommodation and disaccommodation at regular intervals for 24 hours; as a result the stretching regimes do not completely reflect physiological conditions. Despite these limitations, this project is significant in showing that the lens does respond to mechanical stretching with increased LEC proliferation and that the distribution of this proliferation is altered by whether the stretching is static or cyclic.

Future work will seek to more fully understand the interactions between mechanical forces and the biology of the lens. Further understanding of the signaling pathways activated by stretching, and the degree to which the local capsule strains affect signaling is needed and can be obtained by targeting different components of signaling pathways hypothesized to regulate mechanotransduction and LEC proliferation. Target pathways include: Yap/Hippo signaling, Notch/Wnt, and the MAPK/ERK pathways. By inhibiting a component of one of those pathways and observing the effects on mechanotransduction, a more complete picture of what processes govern lens growth can be drawn.

Additionally, the effects of stretching on solute transport in the lens are of considerable interest. It can be hypothesized that the dynamic mechanical forces experienced by the lens during accommodation play a part in driving the transport of solutes in and out of the

92 nucleus of the lens. When the lens no longer is able to accommodate with age, this may slow the transport of solutes and lead to oxidative damage of crystallin proteins in the lens nucleus, eventually contributing to cataract formation.

Future projects may seek to determine the role of the actin cytoskeleton in mediating mechanotransductive signaling in the lens. The cytoskeleton is a pivotal component in transducing mechanical cues into a cellular response. Mechanical stresses are able to modulate cellular functions by activating or inhibiting signal transduction pathways. This is accomplished at the interfaces between the internal cytoskeleton, which is mainly composed of actin filaments, and the extracellular membrane229-233. Dynamic reorganization of the actin cytoskeleton has been found to occur in response to changes in a cell’s microenvironment, one such change being cyclic strain234-239. Additionally, transduction of mechanical forces by the actin cytoskeleton have been implicated in modulating YAP signaling240-242. The mechanosensitive behavior displayed by the LECs is likely mediated by the actin cytoskeleton, and the different stretching regimes may result in different organizations of the cytoskeletons between groups, and it is possible that these different cytoskeletal organizations may influence the proliferative behavior of

LECs differently under different stretching regimes.

Further investigation is necessary to broaden the understanding of how the mechanical environment of the lens affects its biology. The significance of the work included in this dissertation lies in opening this field of study and allowing for the mechanobiology of the lens to direct the development of treatments for age-related refractive issues in the lens.

Chapter 6. Bibliography

1. Levin LA, Adler FH. Adler's physiology of the eye. 2011. 2. Bassnett S. Cell Biology of Lens Epithelial Cells. In: Saika S, Werner L, Lovicu FJ (eds), Lens Epithelium and Posterior Capsular Opacification: Springer Japan; 2014:25-38. 3. Shi Y, De Maria A, Lubura S, Šikić H, Bassnett S. The Penny Pusher: A Cellular Model of Lens GrowthThe Penny Pusher: A Cellular Model of Lens Growth. Investigative Ophthalmology & Visual Science 2015;56:799-809. 4. Šikić H, Shi Y, Lubura S, Bassnett S. A stochastic model of eye lens growth. Journal of theoretical biology 2015;376:15-31. 5. Bassnett S, Šikić H. The lens growth process. Progress in retinal and eye research 2017;60:181-200. 6. Bassnett S. Three-dimensional reconstruction of cells in the living lens: the relationship between cell length and volume. Experimental eye research 2005;81:716- 723. 7. Truscott RJW, Zhu X. Presbyopia and cataract: A question of heat and time. Progress in Retinal and Eye Research 2010;29:487-499. 8. Truscott RJW, Friedrich MG. Molecular Processes Implicated in Human Age- Related Nuclear Cataract. Investigative Ophthalmology & Visual Science 2019;60:5007- 5021. 9. Shui Y-B, Beebe DC. Age-Dependent Control of Lens Growth by Hypoxia. Investigative Ophthalmology & Visual Science 2008;49:1023-1029. 10. Patel I, West SK. Presbyopia: prevalence, impact, and interventions. Community Eye Health 2007;20:40-41. 11. Reilly MA. A quantitative geometric mechanics lens model: Insights into the mechanisms of accommodation and presbyopia. Vis Res Vision Research 2014;103:20- 31. 12. Wang K, Pierscionek BK. Biomechanics of the human lens and accommodative system: Functional relevance to physiological states. Progress in Retinal and Eye Research 2019;71:114-131. 13. Sofija A. The role of lens epithelial cells in the development of the posterior capsule opacifcation and in the lens regeneration after congenital cataract surgery. Zdravniški Vestnik 2017. 14. Kumar B, Reilly MA. The Development, Growth, and Regeneration of the Crystalline Lens: A Review. Current Eye Research 2020;45:313-326.

15. Gwon A. Tissue engineering of the lens: Fundamentals. In: Chirila TV (ed), Biomaterials and Regenerative Medicine in Ophthalmology. Cambridge, UK: Woodhead Publishing Limited; 2010:243-262. 16. Lin H, Ouyang H, Zhu J, et al. Lens regeneration using endogenous stem cells with gain of visual function. Nature 2016;531:323-328. 17. Glasser A. Accommodation: mechanism and measurement. Ophthalmol Clin N Am 2006;19. 18. Glasser A. Restoration of accommodation: surgical options for correction of presbyopia. CXO Clinical and Experimental Optometry 2008;91:279-295. 19. Glasser A, C.W. Campbell M. Biometric, optical and physical changes in the isolated human crystalline lens with age in relation to presbyopia. Vision Research 1999;39:1991-2015. 20. Glasser A, Campbell MCW. Presbyopia and the optical changes in the human crystalline lens with age. Vision Research 1998;38:209-229. 21. Fisher RF. The force of contraction of the human ciliary muscle during accommodation. The Journal of physiology 1977;270:51-74. 22. Fisher RF. Presbyopia and the changes with age in the human crystalline lens. The Journal of physiology 1973;228:765-779. 23. Stachs O, Martin H, Kirchhoff A, Stave J, Terwee T, Guthoff R. Monitoring accommodative ciliary muscle function using three-dimensional ultrasound. Graefe's Archive for Clinical and Experimental Ophthalmology 2002;240:906-912. 24. Bacskulin A, Gast R, Bergmann U, Guthoff R. Ultrasound biomicroscopy monitoring of human ciliary body changes during accommodation in presbyopia. OPHTHALMOLOGE -BERLIN- 1996;93:199-204. 25. Strenk SA, Semmlow JL, Strenk LM, Munoz P, Gronlund-Jacob J, DeMarco JK. Age-Related Changes in Human Ciliary Muscle and Lens: A Magnetic Resonance Imaging Study. INVESTIGATIVE OPHTHALMOLOGY AND VISUAL SCIENCE 1999;40:1162-1169. 26. Tabernero J, Chirre E, Hervella L, Prieto P, Artal P. The accommodative ciliary muscle function is preserved in older humans. Scientific Reports 2016;6:25551. 27. Michael R, Mikielewicz M, Gordillo C, Montenegro GA, Pinilla Cortés L, Barraquer RI. Elastic Properties of Human Lens Zonules as a Function of Age in Presbyopes. Invest Ophthalmol Vis Sci Investigative Opthalmology & Visual Science 2012;53:6109. 28. Augusteyn RC. Growth of the human eye lens. Molecular vision 2007;13:252- 257. 29. Augusteyn RC. Growth of the lens: in vitro observations. Clinical and Experimental Optometry 2008;226. 30. Augusteyn RC. On the growth and internal structure of the human lens. Experimental Eye Research 2010;90:643-654. 31. Wendt M, Croft MA, McDonald J, Kaufman PL, Glasser A. Lens diameter and thickness as a function of age and pharmacologically stimulated accommodation in rhesus monkeys. Experimental eye research 2008;86:746-752.

32. Besner S, Scarcelli G, Pineda R, Yun S-H. In Vivo Brillouin Analysis of the Aging Crystalline Lens. Investigative Ophthalmology & Visual Science 2016;57:5093- 5100. 33. Cheng C, Parreno J, Nowak RB, et al. Age-related changes in eye lens biomechanics, morphology, refractive index and transparency. Aging (Albany NY) 2019;11:12497-12531. 34. Wang K, Venetsanos D, Wang J, Pierscionek BK. Gradient moduli lens models: how material properties and application of forces can affect deformation and distributions of stress. Scientific Reports 2016;6:31171. 35. Heys KR, Cram SL, Truscott RJ. Massive increase in the stiffness of the human lens nucleus with age: the basis for presbyopia? Molecular vision 2004;10:956-963. 36. Wormstone IM, Eldred JA. Experimental models for posterior capsule opacification research. Experimental Eye Research 2016;142:2-12. 37. Kumar B, Chandler HL, Plageman T, Reilly MA. Lens Stretching Modulates Lens Epithelial Cell Proliferation via YAP Regulation. Investigative Ophthalmology & Visual Science 2019;60:3920-3929. 38. Wang E, Reid B, Lois N, Forrester JV, McCaig CD, Zhao M. Electrical inhibition of lens epithelial cell proliferation: an additional factor in secondary cataract? FASEB journal : official publication of the Federation of American Societies for Experimental Biology 2005;19:842-844. 39. Nishi O, Yamamoto N, Nishi K, Nishi Y. Contact inhibition of migrating lens epithelial cells at the capsular bend created by a sharp-edged intraocular lens after cataract surgery. Journal of cataract and refractive surgery 2007;33:1065-1070. 40. Nagamoto T, Hara E. Lens epithelial cell migration onto the posterior capsule in vitro. Journal of cataract and refractive surgery 1996;22 Suppl 1:841-846. 41. Pedrigi RM, Dziezyc J, Kalodimos HA, Humphrey JD. Ex vivo quantification of the time course of contractile loading of the porcine lens capsule after cataract-like surger. Experimental Eye Research 2009;89:869-875. 42. Sureshkumar J, Haripriya A, Muthukkaruppan V, Kaufman PL, Tian B. Cytoskeletal drugs prevent posterior capsular opacification in human lens capsule in vitro. Graefe's archive for clinical and experimental ophthalmology = Albrecht von Graefes Archiv fur klinische und experimentelle Ophthalmologie 2012;250:507-514. 43. Humphrey JD, O'Rourke SL. An introduction to biomechanics : solids and fluids, analysis and design. 2015. 44. Burd HJ, Montenegro GA, Panilla Cortés L, Barraquer RI, Michael R. Equatorial wrinkles in the human lens capsule. Experimental Eye Research 2017;159:77-86. 45. Shi Y, De Maria A, Lubura S, Šikić H, Bassnett S. The penny pusher: a cellular model of lens growth. Investigative ophthalmology & visual science 2015;56:799-809. 46. Pedrigi RM, Dziezyc J, Kalodimos HA, Humphrey JD. Ex vivo quantification of the time course of contractile loading of the porcine lens capsule after cataract-like surgery. Experimental eye research 2009;89:869-875. 47. Kuszak JR, Zoltoski RK, Sivertson C. Fibre cell organization in crystalline lenses. Experimental Eye Research 2004;78:673-687.

48. Lovicu FJ, McAvoy JW. Growth factor regulation of lens development. Developmental Biology 2005;280:1-14. 49. McAvoy JW, Dawes LJ, Sugiyama Y, Lovicu FJ. Intrinsic and extrinsic regulatory mechanisms are required to form and maintain a lens of the correct size and shape. Experimental eye research 2017;156:34-40. 50. Dawes LJ, Shelley EJ, McAvoy JW, Lovicu FJ. A role for Hippo/YAP-signaling in FGF-induced lens epithelial cell proliferation and fibre differentiation. Experimental Eye Research 2018;169:122-133. 51. Sun L, Qu L, Zhu R, et al. Effects of Mechanical Stretch on Cell Proliferation and Matrix Formation of Mesenchymal Stem Cell and Anterior Cruciate Ligament Fibroblast. Stem Cells International Stem Cells International 2016;2016:1-10. 52. Feng Y, Tian XY, Sun P, Cheng ZP, Shi RF. Simultaneous Study of Mechanical Stretch-Induced Cell Proliferation and Apoptosis on C2C12 Myoblasts. Cells, tissues, organs 2018;205:189-196. 53. Gudipaty SA, Lindblom J, Loftus PD, et al. Mechanical stretch triggers rapid epithelial cell division through Piezo1. Nature 2017;543:118-121. 54. Yano S, Komine M, Fujimoto M, Okochi H, Tamaki K. Mechanical Stretching In Vitro Regulates Signal Transduction Pathways and Cellular Proliferation in Human Epidermal Keratinocytes. Journal of Investigative Dermatology 2004;122:783-790. 55. Parreno J, Cheng C, Nowak RB, Fowler VM. The effects of mechanical strain on mouse eye lens capsule and cellular microstructure. Molecular biology of the cell 2018;29:1963-1974. 56. O'Connor MD, Wederell ED, de Iongh R, Lovicu FJ, McAvoy JW. Generation of transparency and cellular organization in lens explants. Experimental eye research 2008;86:734-745. 57. Ehrmann K, Ho A, Parel JM, Ehrmann K, Ho A, Parel J-M. Biomechanical analysis of the accommodative apparatus in primates. Clinical & Experimental Optometry 2008;91:302-312. 58. Reilly MA, Hamilton PD, Perry G, Ravi N. Comparison of the behavior of natural and refilled porcine lenses in a robotic lens stretcher. Experimental Eye Research 2009;88:483-494. 59. Reilly MA, Hamilton PD, Ravi N. Dynamic multi-arm radial lens stretcher: a robotic analog of the ciliary body. Experimental eye research 2008;86:157-164. 60. Sharma PK, Busscher HJ, Terwee T, Koopmans SA, van Kooten TG. A comparative study on the viscoelastic properties of human and animal lenses. 2011:681- 688. 61. Ziebarth NM, Borja D, Arrieta E, et al. Role of the lens capsule on the mechanical accommodative response in a lens stretcher. Investigative ophthalmology & visual science 2008;49:4490-4496. 62. Pellegrino A, Burd HJ, Pinilla Cortés L, et al. Anterior lens capsule strains during simulated accommodation in porcine eyes. Experimental eye research 2018;168:19-27. 63. Augusteyn RC, Mohamed A, Nankivil D, et al. Age-dependence of the optomechanical responses of ex vivo human lenses from India and the USA, and the

97 force required to produce these in a lens stretcher: The similarity to in vivo disaccommodation. 2011:1667-1678. 64. Borja D, Manns F, Ho A, et al. Optical power of the isolated human crystalline lens. Investigative ophthalmology & visual science 2008;49:2541-2548. 65. Codelia VA, Sun G, Irvine KD. Regulation of YAP by mechanical strain through Jnk and Hippo signaling. Current biology : CB 2014;24:2012-2017. 66. Panciera T, Azzolin L, Cordenonsi M, Piccolo S. Mechanobiology of YAP and TAZ in physiology and disease. Nature Reviews Molecular Cell Biology 2017;758. 67. Lin C, Yao E, Zhang K, et al. YAP is essential for mechanical force production and epithelial cell proliferation during lung branching morphogenesis. eLife 2017. 68. Mendoza-Reinoso V, Beverdam A. Epidermal YAP activity drives canonical WNT16/β-catenin signaling to promote keratinocyte proliferation in vitro and in the murine ski. Stem Cell Research 2018;29:15-23. 69. Tsonis PA, Del Rio-Tsonis K. Lens and retina regeneration: transdifferentiation, stem cells and clinical applications. Experimental Eye Research 2004;78:161-172. 70. Wolff G. Entwickelungsphysiologische Studien. Archiv für Mikroskopische Anatomie 1895;1:380-390. 71. Freeman G. Lens regeneration from the cornea in Xenopus laevis. Journal of Experimental Zoology 1963;154:39-65. 72. Gwon A. Lens Regeneration in Mammals: A Review. Survey of Ophthalmology 2006;51:51-62. 73. Chow RL, Lang RA. EARLY EYE DEVELOPMENT IN VERTEBRATES. Annual Review of Cell & Developmental Biology 2001;17:255. 74. Call MK, Grogg MW, Tsonis PA. Eye on regeneration. Anatomical record Part B, New anatomist 2005;287:42-48. 75. Sikic H, Shi Y, Lubura S, Bassnett S. A stochastic model of eye lens growth. Journal of theoretical biology 2015;376:15-31. 76. Vergara MN, Tsissios G, Del Rio-Tsonis K. Lens regeneration: a historical perspective. Int J Dev Biol 2018;62:351-361. 77. Dinsmore CE, Zoologists ASo, Meeting ASoZ. A History of Regeneration Research: Milestones in the Evolution of a Science: Cambridge University Press; 1991. 78. Elliott SA, Sánchez Alvarado A. The history and enduring contributions of planarians to the study of animal regeneration. Wiley Interdiscip Rev Dev Biol 2013;2:301-326. 79. Spallanzani L. Prodromo di un opera da imprimersi sopra la riproduzioni animali (An Essay on Animal Reproduction). Mati M translation T Becket and de Hondt, London 1769. 80. Tsonis PA. Limb regeneration: Cambridge university press; 1996. 81. Tsonis PA, Fox TP. Regeneration according to Spallanzani. Developmental dynamics : an official publication of the American Association of Anatomists 2009;238:2357-2363. 82. Morgan TH, Moszkowski M. Regeneration: Wilhelm Engelmann; 1907. 83. Colucci V. Sulla rigenerazione parziale dell'occhio nei Tritoni. Istogenesi e svilluppo Studio sperimentale Mem R Accad Bologna Sez Sci Nat 1891;1:167-203. 98

84. Sunderland ME. Regeneration: Thomas Hunt Morgan's window into development. Journal of the history of biology 2010;43:325-361. 85. Okada TS. Experimental embryology in Japan, 1930-1960. A historical background of developmental biology in Japan. The International journal of developmental biology 1994;38:135-154. 86. Okada T. A brief history of regeneration research—For admiring Professor Niazi’s discovery of the effect of vitamin A on regeneration. Journal of biosciences 1996;21:261-271. 87. Eguchi G. A life in research on lens regeneration and transdifferentiation. An interview with Goro Eguchi. The International journal of developmental biology 2004;48:695-700. 88. Stone L, Sapir P. Experimental studies on the regeneration of the lens in the eye of anurans, urodeles and fishes. Journal of Experimental Zoology 1940;85:71-101. 89. Eguchi G. ELECTRON MICROSCOPIC STUDIES ON LENS REGENERATION. Embryologia 1963;8:45-62. 90. Eguchi G. ELECTRON MICROSCOPIC STUDIES ON LENS REGENERATION. Embryologia 1964;8:247-287. 91. Grogg MW, Call MK, Okamoto M, Vergara MN, Del Rio-Tsonis K, Tsonis PA. BMP inhibition-driven regulation of six-3 underlies induction of newt lens regeneration. Nature 2005;438:858-862. 92. Looso M, Preussner J, Sousounis K, et al. A de novo assembly of the newt transcriptome combined with proteomic validation identifies new protein families expressed during tissue regeneration. Genome Biol 2013;14:R16-R16. 93. Keinath MC, Voss SR, Tsonis PA, Smith JJ. A linkage map for the Newt Notophthalmus viridescens: Insights in vertebrate genome and chromosome evolution. Developmental biology 2017;426:211-218. 94. Sousounis K, Bhavsar R, Looso M, et al. Molecular signatures that correlate with induction of lens regeneration in newts: lessons from proteomic analysis. Human genomics 2014;8:22-22. 95. Henry JJ, Tsonis PA. Molecular and cellular aspects of amphibian lens regeneration. Progress in retinal and eye research 2010;29:543-555. 96. Schaefer JJ, Oliver G, Henry JJ. Conservation of gene expression during embryonic lens formation and cornea-lens transdifferentiation in Xenopus laevis. Developmental dynamics : an official publication of the American Association of Anatomists 1999;215:308-318. 97. Mizuno N, Mochii M, Takahashi TC, Eguchi G, Okada TS. Lens regeneration in Xenopus is not a mere repeat of lens development, with respect to crystallin gene expression. Differentiation; research in biological diversity 1999;64:143-149. 98. Henry JJ, Carinato ME, Schaefer JJ, et al. Characterizing gene expression during lens formation in Xenopus laevis: evaluating the model for embryonic lens induction. Developmental dynamics : an official publication of the American Association of Anatomists 2002;224:168-185. 99. Henry JJ. The cellular and molecular bases of vertebrate lens regeneration. International review of cytology 2003;228:195-265. 99

100. Cannata SM, Arresta E, Bernardini S, Gargioli C, Filoni S. Tissue interactions and lens-forming competence in the outer cornea of larval Xenopus laevis. Journal of experimental zoology Part A, Comparative experimental biology 2003;299:161-171. 101. Kha CX, Son PH, Lauper J, Tseng KA. A model for investigating developmental eye repair in Xenopus laevis. Experimental eye research 2018;169:38-47. 102. Perry KJ, Thomas AG, Henry JJ. Expression of pluripotency factors in larval epithelia of the frog Xenopus: evidence for the presence of cornea epithelial stem cells. Developmental biology 2013;374:281-294. 103. PHYSIOLOGY.: Experiments relating to the reproduction of the Crystalline lens, by MM. Cocteau and Leroy d'Etoille, Doctors of Physic. Account read at the Academy of Surgery, 10th Feb. 1825. The Lancet 1827;8:117-118. 104. Milliot B. Experiments on the restoration of a normalcrystalline lens in some mammals after its removal. J Anat Physiol (Paris) 1872;8:1. 105. RL R. The regeneration of the crystallin lens: an experimental study. Johns Hopkins Hosp Rep 1900;237-263. 106. Loewenhardt. Einige Versuche um die Regeneration der Krystallinse. Neune Notizen von Froriep 1841;19. 107. H D. Re-production of the lens. The Lancet 1828;2. 108. R M. On the reproduction of the crystalline lens. Lond Med Gaz 1832;344-348. 109. Mayer. Uber die reproduktion der Krystallinse. J Chirurgie Augenheilkunde 1832;524. 110. Agarwal L, Angra S, Khosla P, Tandon H. Lens regeneration in mammals. II- monkeys. Orient A Ophth 1964;2:47-59. 111. Gwon A, Gruber LJ, Mantras C. Restoring lens capsule integrity enhances lens regeneration in New Zealand albino rabbits and cats. Journal of Cataract & Refractive Surgery 1993;19:735-746. 112. Sikharuldze T. Exchange of crystallin lens in rabbits by embryonic skin ectoderm. Bull Acad Sci Georg SSR 1956;14:337-356. 113. Stewart DS, ESPINASSE PG. Regeneration of the lens of the eye in the rabbit. Nature 1959;183:1815. 114. Angra S, Agarwal L, Khosla P. Lens regeneration in mammals-I. East Arch Ophtalmol 1973;1:214-224. 115. Binder HF, Binder RF, Wells AH, Katz RL. Experiments on Lens Regeneration in Rabbits*: I. Lens Regrowth after Extracapsular Extraction with Keyhole Iridectomy. American journal of ophthalmology 1961;52:919-922. 116. PETTIT TH. A Study of Lens Regeneration in the Rabbit. Investigative Ophthalmology & Visual Science 1963;2:243-251. 117. METZ HS, LIVINGSTON W, ZIGMAN S, LERMAN S. Studies on the metabolism of the regenerating rabbit lens. Archives of Ophthalmology 1965;74:244-247. 118. Gwon AE, Gruber LJ, Mundwiler KE. A histologic study of lens regeneration in aphakic rabbits. Investigative ophthalmology & visual science 1990;31:540-547. 119. Gwon A, Kuszak J, Gruber LJ. Intralenticular implant study in pigmented rabbits: opacity lensmeter assessment. Journal of cataract and refractive surgery 1999;25:268- 277. 100

120. Gwon A, Gruber L. Engineering the crystalline lens with a biodegradable or non- degradable scaffold. Experimental Eye Research 2010;91:220-228. 121. Lin H, Ouyang H, Zhu J, et al. Lens regeneration using endogenous stem cells with gain of visual function. 2016:323-+. 122. Sukhija J, Kaur S. Nature nurtures: lens regeneration, a breakthrough in ophthalmology. Annals of Eye Science 2017;2. 123. Graw J. Chapter Ten - Eye Development. In: Koopman P (ed), Current Topics in Developmental Biology: Academic Press; 2010:343-386. 124. Richardson R, Tracey-White D, Webster A, Moosajee M. The zebrafish eye-a paradigm for investigating human ocular genetics. Eye (Lond) 2017;31:68-86. 125. Bibliowicz J, Tittle RK, Gross JM. Toward a better understanding of human eye disease insights from the zebrafish, Danio rerio. Progress in molecular biology and translational science 2011;100:287-330. 126. Beebe DC, Coats JM. The Lens Organizes the Anterior Segment: Specification of Neural Crest Cell Differentiation in the Avian Eye. Developmental Biology 2000;220:424-431. 127. Hay ED. Development of the Vertebrate Cornea; 1980:263-322. 128. Hay ED. An overview of epithelio-mesenchymal transformation. Acta anatomica 1995;154:8-20. 129. Johnston MC, Noden DM, Hazelton RD, Coulombre JL, Coulombre AJ. Origins of avian ocular and periocular tissues. Experimental Eye Research 1979;29:27-43. 130. Zinn KM. Changes in Corneal Ultrastructure Resulting from Early Lens Removal in the Developing Chick Embryo. Investigative Ophthalmology & Visual Science 1970;9:165-182. 131. Danysh BP, Duncan MK. The lens capsule. Experimental Eye Research 2009;88:151-164. 132. Bassnett S, Costello MJ. The cause and consequence of fiber cell compaction in the vertebrate lens. Experimental eye research 2017;156:50-57. 133. Bassuk JA, Birkebak TED, Rothmier JD, et al. Disruption of theSparcLocus in Mice Alters the Differentiation of Lenticular Epithelial Cells and Leads to Cataract Formation. Experimental Eye Research 1999;68:321-331. 134. Sasaki T, Göhring W, Timpl R, Hohenester E. Crystal structure and mapping by site-directed mutagenesis of the collagen-binding epitope of an activated form of BM- 40/SPARC/osteonectin. EMBO Journal 1998;17:1625-1634. 135. Termine JD, Whitson SW, Conn KM, Kleinman HK, McGarvey ML, Martin GR. Osteonectin, a bone-specific protein linking mineral to collagen. Cell 1981;26:99-105. 136. Hanna C, O'Brien JE. Cell production and migration in the epithelial layer of the lens. Archives of ophthalmology (Chicago, Ill : 1960) 1961;66:103-107. 137. Bassnett S, Winzenburger PA. Morphometric analysis of fibre cell growth in the developing chicken lens. Experimental eye research 2003;76:291-302. 138. Zhou M, Leiberman J, Xu J, Lavker RM. A Hierarchy of Proliferative Cells Exists in Mouse Lens Epithelium: Implications for Lens Maintenance. Investigative Ophthalmology & Visual Science 2006;47:2997-3003.

101

139. Kuwabara T. The maturation of the lens cell: a morphologic study. Experimental Eye Research 1975;20:427-443. 140. Kuszak JR, Zoltoski RK, Tiedemann CE. Development of lens sutures. The International journal of developmental biology 2004;48:889-902. 141. al-Ghoul KJ, Costello MJ. Light microscopic variation of fiber cell size, shape and ordering in the equatorial plane of bovine and human lenses. Molecular vision 1997;3:2. 142. Wride MA. Lens fibre cell differentiation and organelle loss: many paths lead to clarity. Philosophical transactions of the Royal Society of London Series B, Biological sciences 2011;366:1219-1233. 143. Vrensen GF, Graw J, De Wolf A. Nuclear breakdown during terminal differentiation of primary lens fibres in mice: a transmission electron microscopic study. Experimental eye research 1991;52:647-659. 144. Bassnett S. On the mechanism of organelle degradation in the vertebrate lens. Experimental Eye Research 2009;88:133-139. 145. Graw J. Genetics of crystallins: cataract and beyond. Experimental eye research 2009;88:173-189. 146. Truscott RJ, Zhu X. Presbyopia and cataract: a question of heat and time. Progress in retinal and eye research 2010;29:487-499. 147. Allen D, Vasavada A. Cataract and surgery for cataract. BMJ (Clinical research ed) 2006;333:128-132. 148. Ale J, Manns F, Ho A. Evaluation of the performance of accommodating IOLs using a paraxial optics analysis. Ophthalmic & Physiological Optics 2010;30:132-142. 149. Stone LS. Further experiments on lens regeneration from retina pigment cells in adult newt eyes. The Journal of experimental zoology 1957;136:75-87. 150. Eguchi G, Shingai R. Cellular analysis on localization of lens forming potency in the newt iris epithelium. Development, growth & differentiation 1971;13:337-349. 151. Reyer RW, Woolfitt RA, Withersty LT. Stimulation of lens regeneration from the newt dorsal iris when implanted into the blastema of the regenerating limb. Developmental biology 1973;32:258-281. 152. Eguchi G, Eguchi Y, Nakamura K, Yadav MC, Millán JL, Tsonis PA. Regenerative capacity in newts is not altered by repeated regeneration and ageing. Nature Communications 2011;2:384. 153. Henry JJ. Diverse Evolutionary Origins and Mechanisms of Lens Regeneration. Molecular biology and evolution 35:1563-1575. 154. Reyer RW. Repolarization of reversed, regenerating lenses in adult newts, Notophthalmus viridescens. Experimental eye research 1977;24:501-509. 155. Yamada T. Control mechanisms in cell-type conversion in newt lens regeneration. Monographs in developmental biology 1977;13:1-126. 156. Tsonis PA. Regeneration in vertebrates. Developmental biology 2000;221:273- 284. 157. McDevitt DS, Brahma SK, Courtois Y, Jeanny JC. Fibroblast growth factor receptors and regeneration of the eye lens. Developmental dynamics : an official publication of the American Association of Anatomists 1997;208:220-226. 102

158. Del Rio-Tsonis K, Trombley MT, McMahon G, Tsonis PA. Regulation of lens regeneration by fibroblast growth factor receptor 1. Developmental dynamics : an official publication of the American Association of Anatomists 1998;213:140-146. 159. Hayashi T, Mizuno N, Ueda Y, Okamoto M, Kondoh H. FGF2 triggers iris- derived lens regeneration in newt eye. Mechanisms of development 2004;121:519-526. 160. Barbosa-Sabanero K. Lens and retina regeneration: new perspectives from model organisms. Biochem J 447:321-334. 161. Madhavan M, Haynes TL, Frisch NC, et al. The role of Pax-6 in lens regeneration. Proceedings of the National Academy of Sciences of the United States of America 2006;103:14848-14853. 162. Del Rio-Tsonis K, Washabaugh CH, Tsonis PA. Expression of pax-6 during urodele eye development and lens regeneration. Proceedings of the National Academy of Sciences of the United States of America 1995;92:5092-5096. 163. Hayashi T, Mizuno N, Takada R, Takada S, Kondoh H. Determinative role of Wnt signals in dorsal iris-derived lens regeneration in newt eye. Mechanisms of development 2006;123:793-800. 164. Grigoryan EN. Molecular Factors of the Maintenance and Activation of the Juvenile Phenotype of Cellular Sources for Eye Tissue Regeneration. Biochemistry Biokhimiia 2018;83:1318-1331. 165. Chamberlain CG, McAvoy JW. Fibre differentiation and polarity in the mammalian lens: a key role for FGF. Progress in Retinal and Eye Research 1997;16:443- 478. 166. Henry JJ, Thomas AG, Hamilton PW, Moore L, Perry KJ. Cell Signaling Pathways in Vertebrate Lens Regeneration. In: Heber-Katz E, Stocum DL (eds), New Perspectives in Regeneration. Berlin, Heidelberg: Springer Berlin Heidelberg; 2013:75- 98. 167. Reeve JG. Studies on the Regeneration of Lens from Cornea in larval Xenopus laevis. University of Southampton; 1978. 168. Filoni S, Bernardini S, Cannata SM, apos, alessio A. Lens Regeneration in Larval Xenopus laevis: Experimental Analysis of the Decline in the Regenerative Capacity during Developmen. Developmental Biology 1997;187:13-24. 169. Filoni S, Bosco L, Cioni C. The role of neural retina in lens regeneration from cornea in larval Xenopus laevis. Acta embryologiae et morphologiae experimentalis ("Halocynthia" Association") 1982;3:15-28. 170. Bosco L, Testa O, Venturini G, Willems D. Lens fibre transdifferentiation in cultured larval Xenopus laevis outer cornea under the influence of neural retina- conditioned medium. Cellular and molecular life sciences : CMLS 1997;53:921-928. 171. Reeve JG, Wild AE. Lens regeneration from cornea of larval Xenopus laevis in the presence of the lens. Journal of embryology and experimental morphology 1978;48:205-214. 172. Day RC, Beck CW. Transdifferentiation from cornea to lens in Xenopus laevis depends on BMP signalling and involves upregulation of Wnt signalling. BMC Developmental Biology 2011;11:54.

103

173. Fukui L, Henry JJ. FGF signaling is required for lens regeneration in Xenopus laevis. The Biological bulletin 2011;221:137-145. 174. Hamilton PW, Sun Y, Henry JJ. Lens regeneration from the cornea requires suppression of Wnt/β-catenin signaling. Experimental Eye Research 2016;145:206-215. 175. Fuhrmann S. Wnt signaling in eye organogenesis. Organogenesis 2008;4:60-67. 176. Gwon AE, Jones RL, Gruber LJ, Mantras C. Lens regeneration in juvenile and adult rabbits measured by image analysis. Investigative ophthalmology & visual science 1992;33:2279-2283. 177. Gwon A, Enomoto H, Horowitz J, Garner MH. Induction of de novo synthesis of crystalline lenses in aphakic rabbits. Experimental eye research 1989;49:913-926. 178. Taylor VL, al-Ghoul KJ, Lane CW, Davis VA, Kuszak JR, Costello MJ. Morphology of the normal human lens. Invest Ophthalmol Vis Sci 1996;37:1396-1410. 179. Bito LZ, Davson H, Snider N. The effects of autonomic drugs on mitosis and DNA synthesis in the lens epithelium and on the composition of the aqueous humour. Experimental Eye Research 1965;4:54-61. 180. Bito LZ, Harding CV. Patterns of cellular organization and cell division in the epithelium of the cultured lens. Experimental eye research 1965;4:146-161. 181. Gindi J, WL W, Schanzlin D. Endocapsular Cataract Surgery-1. Surgical Technique. Cataract. International Journal of Cataract Surgery 1985;2:5-10. 182. Tan X, Liu Z, Zhu Y, et al. The Fate of In Situ Lens Regeneration is Determined by Capsulorhexis Size. Current molecular medicine 2017;17:270-279. 183. Murphy P, Kabir MH, Srivastava T, et al. Light-focusing human micro-lenses generated from pluripotent stem cells model lens development and drug-induced cataract in vitro. Development (Cambridge, England) 145:dev155838. 184. O'Connor MD, Wederell ED, de Iongh R, Lovicu FJ, McAvoy JW. Generation of transparency and cellular organization in lens explants. Experimental Eye Research 2008;86:734-745. 185. Glasser A, Hilmantel G, Calogero D, MacRae S, Masket S, Stark W. Special Report: American Academy of Ophthalmology Task Force Recommendations for Test Methods to Assess Accommodation Produced by Intraocular Lenses. Ophthalmology 2017;124. 186. Jackson E. Amplitude of Accommodation at Different Periods of Life. California state journal of medicine 1907;5:163-166. 187. Ziebarth NM, Arrieta E, Feuer WJ, Moy VT, Manns F, Parel J-M. Primate lens capsule elasticity assessed using Atomic Force Microscopy. 2011:490-494. 188. Marussich L, Manns F, Nankivil D, et al. Measurement of Crystalline Lens Volume During Accommodation in a Lens Stretcher. Investigative ophthalmology & visual science 2015;56:4239-4248. 189. Cui Y, Hameed FM, Yang B, et al. Cyclic stretching of soft substrates induces spreading and growth. Nature Communications 2015;6:6333. 190. Rueden CT, Schindelin J, Hiner MC, et al. ImageJ2: ImageJ for the next generation of scientific image data. BMC Bioinformatics 2017;18:529. 191. Wu JJ, Wu W, Tholozan FM, Saunter CD, Girkin JM, Quinlan RA. A dimensionless ordered pull-through model of the mammalian lens epithelium evidences 104 scaling across species and explains the age-dependent changes in cell density in the human lens. Journal of the Royal Society, Interface 2015;12:20150391. 192. Wiley LA, Shui YB, Beebe DC. Visualizing lens epithelial cell proliferation in whole lenses. Molecular vision 2010;16:1253-1259. 193. Otsu N. A Threshold Selection Method from Gray-Level Histograms. IEEE Transactions on Systems, Man, and Cybernetics 1979;9:62-66. 194. Augusteyn RC. On the growth and internal structure of the human lens. YEXER Experimental Eye Research 2010;90:643-654. 195. Meng Z, Moroishi T, Guan KL. Mechanisms of Hippo pathway regulation. Genes & development 2016;30:1-17. 196. Nardone G, Oliver-De La Cruz J, Vrbsky J, et al. YAP regulates cell mechanics by controlling focal adhesion assembly. Nature communications 2017;8:15321-15321. 197. Aragona M, Panciera T, Manfrin A, et al. A mechanical checkpoint controls multicellular growth through YAP/TAZ regulation by actin-processing factors. Cell 2013;154:1047-1059. 198. Dupont S. Review Article: Role of YAP/TAZ in cell-matrix adhesion-mediated signalling and mechanotransduction. Experimental Cell Research 2016;343:42-53. 199. Abuammah A, Maimari N, Towhidi L, et al. New developments in mechanotransduction: Cross talk of the Wnt, TGF-β and Notch signalling pathways in reaction to shear stres. Current Opinion in Biomedical Engineering 2018;5:96-104. 200. Halder G, Dupont S, Piccolo S. Transduction of mechanical and cytoskeletal cues by YAP and TAZ. Nature Reviews Molecular Cell Biology 2012;13:591-600. 201. Wada K-I, Itoga K, Okano T, Yonemura S, Sasaki H. Hippo pathway regulation by cell morphology and stress fibers. Development 2011;138:3907. 202. Feng X, Degese Maria S, Iglesias-Bartolome R, et al. Article: Hippo-Independent Activation of YAP by the GNAQ Uveal Melanoma Oncogene through a Trio-Regulated Rho GTPase Signaling Circuitry. Cancer Cell 2014;25:831-845. 203. You B, Yang Y-L, Xu Z, et al. Inhibition of ERK1/2 down-regulates the Hippo/YAP signaling pathway in human NSCLC cells. Oncotarget 2015;6:4357-4368. 204. Kippenberger S, Bernd A, Loitsch S, et al. Signaling of Mechanical Stretch in Human Keratinocytes via MAP Kinase. Journal of Investigative Dermatology 2000;114:408-412. 205. Hofmann M, Žaper J, Bernd A, Bereiter-Hahn J, Kaufmann R, Kippenberger S. Mechanical pressure-induced phosphorylation of p38 mitogen-activated protein kinase in epithelial cells via Src and protein kinase. Biochemical and Biophysical Research Communications 2004;316:673-679. 206. Sheppard AL, Evans CJ, Singh KD, Wolffsohn JS, Dunne MCM, Davies LN. Three-Dimensional Magnetic Resonance Imaging of the Phakic Crystalline Lens during Accommodation. Investigative Ophthalmology & Visual Science 2011;52:3689-3697. 207. Candia OA. Surface and Volume Changes in the Lens during Accommodation. Investigative Ophthalmology & Visual Science 2011;52:3698-3698. 208. Manns F, Parel J-M, Denham D, et al. Optomechanical Response of Human and Monkey Lenses in a Lens Stretcher. Investigative Ophthalmology & Visual Science 2007;48:3260-3268. 105

209. Reilly MA, Ravi N. A geometric model of ocular accommodation. VR Vision Research 2010;50:330-336. 210. Burd HJ, Judge SJ, Cross JA. Numerical modelling of the accommodating lens. VR Vision Research 2002;42:2235-2251. 211. Reilly M, Ravi N. Microindentation of the Young Porcine Ocular Lens. Journal of Biomechanical Engineering 2009;131:044502-044502-044504. 212. Kammel R, Ackermann R, Mai T, Damm C, Nolte S. Pig lenses in a lens stretcher: implications for presbyopia treatment. Optometry and vision science : official publication of the American Academy of Optometry 2012;89:908-915. 213. De Korte CL, Van Der Steen AFW, Thijssen JM, Duindam JJ, Otto C, Puppels GJ. Relation Between Local Acoustic Parameters and Protein Distribution in Human and Porcine Eye Lenses. Experimental Eye Research 1994;59:617-627. 214. Reilly MA, Rapp B, Hamilton PD, Shen AQ, Ravi N. Material characterization of porcine lenticular soluble proteins. Biomacromolecules 2008;9:1519-1526. 215. Sureshkumar J, Haripriya A, Muthukkaruppan V, Kaufman PL, Tian B. Cytoskeletal drugs prevent posterior capsular opacification in human lens capsule in vitro. 2012:507-514. 216. Rafferty NS, Rafferty KA, Jr. Cell population kinetics of the mouse lens epithelium. Journal of cellular physiology 1981;107:309-315. 217. Wu JJ, Wu W, Tholozan FM, Saunter CD, Girkin JM, Quinlan RA. A dimensionless ordered pull-through model of the mammalian lens epithelium evidences scaling across species and explains the age-dependent changes in cell density in the human lens. Journal of the Royal Society Interface 2015;12:20150391. 218. Wiley LA, Shui YB, Beebe DC. Visualizing lens epithelial cell proliferation in whole lenses. Molecular vision 2010;16:1253-1259. 219. Ong MD, Payne DM, Garner MH. Differential protein expression in lens epithelial whole-mounts and lens epithelial cell cultures. Experimental eye research 2003;77:35-49. 220. Krag S, Andreassen TT. Biomechanical Measurements of the Porcine Lens Capsule. Experimental Eye Research 1996;62:253-260. 221. Fisher RF. Elastic constants of the human lens capsule. The Journal of physiology 1969;201:1-19. 222. Hermans EA, Dubbelman M, van der Heijde GL, Heethaar RM. Estimating the external force acting on the human eye lens during accommodation by finite element modelling. Vision Res 2006;46:3642-3650. 223. Salic A, Mitchison TJ. A chemical method for fast and sensitive detection of DNA synthesis in vivo. Proceedings of the National Academy of Sciences 2008;105:2415-2420. 224. Kaiser CL, Kamien AJ, Shah PA, Chapman BJ, Cotanche DA. 5-Ethynyl-2'- deoxyuridine labeling detects proliferating cells in the regenerating avian cochlea. Laryngoscope 2009;119:1770-1775. 225. Scholzen T, Gerdes J. The Ki-67 protein: from the known and the unknown. J Cell Physiol 2000;182:311-322.

106

226. Basiji DA, Ortyn WE, Liang L, Venkatachalam V, Morrissey P. Cellular image analysis and imaging by flow cytometry. Clin Lab Med 2007;27:653-viii. 227. Meng Z, Moroishi T, Guan K-L. Mechanisms of Hippo pathway regulation. Genes & development 2016;30:1-17. 228. Hermans EA, Pouwels PJW, Dubbelman M, Kuijer JPA, van der Heijde RGL, Heethaar RM. Constant Volume of the Human Lens and Decrease in Surface Area of the Capsular Bag during Accommodation: An MRI and Scheimpflug Study. Investigative Ophthalmology & Visual Science 2009;50:281-289. 229. Alenghat FJ, Ingber DE. Mechanotransduction: all signals point to cytoskeleton, matrix, and integrins. Science's STKE : signal transduction knowledge environment 2002;2002:pe6. 230. Shafrir Y, Forgacs G. Mechanotransduction through the cytoskeleton. American journal of physiology Cell physiology 2002;282:C479-486. 231. Forgacs G, Yook SH, Janmey PA, Jeong H, Burd CG. Role of the cytoskeleton in signaling networks. Journal of cell science 2004;117:2769-2775. 232. Maxwell CA, Hendzel MJ. The integration of tissue structure and nuclear function. Biochemistry and cell biology = Biochimie et biologie cellulaire 2001;79:267- 274. 233. Harris AR, Jreij P, Fletcher DA. Mechanotransduction by the Actin Cytoskeleton: Converting Mechanical Stimuli into Biochemical Signals. Annual Review of Biophysics 2016;47:617-631. 234. Letort G, Ennomani H, Gressin L, Théry M, Blanchoin L. Dynamic reorganization of the actin cytoskeleton. F1000Res 2015;4:F1000 Faculty Rev-1940. 235. Lou H-Y, Zhao W, Li X, et al. Membrane curvature underlies actin reorganization in response to nanoscale surface topography. Proceedings of the National Academy of Sciences 2019;116:23143-23151. 236. Yamazaki D, Kurisu S, Takenawa T. Regulation of cancer cell motility through actin reorganization. Cancer science 2005;96:379-386. 237. Geiger RC, Taylor W, Glucksberg MR, Dean DA. Cyclic stretch-induced reorganization of the cytoskeleton and its role in enhanced gene transfer. Gene Ther 2006;13:725-731. 238. Goldyn AM, Kaiser P, Spatz JP, Ballestrem C, Kemkemer R. The kinetics of force-induced cell reorganization depend on microtubules and actin. Cytoskeleton (Hoboken) 2010;67:241-250. 239. Livne A, Bouchbinder E, Geiger B. Cell reorientation under cyclic stretching. Nature Communications 2014;5:3938. 240. Gao J, He L, Zhou L, et al. Mechanical force regulation of YAP by F-actin and GPCR revealed by super-resolution imaging. Nanoscale 2020;12:2703-2714. 241. Mana-Capelli S, Paramasivam M, Dutta S, McCollum D. Angiomotins link F- actin architecture to Hippo pathway signaling. Molecular biology of the cell 2014;25:1676-1685. 242. Seo J, Kim J. Regulation of Hippo signaling by actin remodeling. BMB Rep 2018;51:151-156.

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