University of Southern California Dissertations and Theses

Laser induced modulation of ocular renin-angiotensin system for treatment of dry age-related macular degeneration

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Laser induced modulation of ocular renin-angiotensin system for treatment of dry age-related macular degeneration

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Content Copyright 2023 Eugene Zhou

Laser Induced Modulation of Ocular Renin-Angiotensin System for Treatment of Dry Age-
Related Macular Degeneration

By

Eugene Zhou

A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
CLINICAL AND EXPERIMENTAL THERAPEUTICS (CXPT)

May 2023

ii
Acknowledgements
This dissertation would not have been possible without the training, guidance, and support I
received from the many individuals I have met throughout this experience.
I would like to thank my advisor, Dr. Stan G. Louie, for his guidance and support; shaping me
into the scientist that I am today. We have faced and overcome many challenges, which would not have
been possible without your unwavering confidence in and support for your students.
I also want to thank my committee members, Dr. Asante and Dr. Asatryan, for all their warmth,
support, and guidance throughout these many years together. Their feedback has been invaluable to
me.
I am incredibly fortunate to have had lab mates with whom I’ve shared the rollercoaster
experience that is the PhD journey. I could not have reached this level of achievement without all the
members of our lab, past and present, Hua, Tiange, Priyal, Kabir, Brandon, Andrew, Angela, Rita, Yahya,
Lindsey, Darryl, Roland, and Cindy. I cannot wait for our paths to intersect again.
Finally, I would like to thank my friends and family, who have always pushed me to excel and
were my source of motivation. I am forever grateful for their support.

iii
Table of Contents
Acknowledgements ........................................................................................................................ ii
List of Tables ................................................................................................................................. vi
List of Figures .............................................................................................................................. viii
Abstract ........................................................................................................................................ xiii
Chapter 1: Introduction to Age-Related Macular Degeneration, Subthreshold Nano-lasers, and
Wound Healing ............................................................................................................................... 1
1.1 Epidemiology of AMD ........................................................................................................................ 1
1.2 Risk Factors and Pathogenesis of AMD .............................................................................................. 2
1.3 Retinal Pigment Epithelium and Age-Related Dysfunction ................................................................ 3
1.4 Drusen as a Clinical Biomarker of AMD .............................................................................................. 5
1.4.1 Lipids in Drusen .......................................................................................................................................... 5
1.4.2 Proteins in Drusen ...................................................................................................................................... 6
1.5 Current Therapies In Development for Dry AMD ............................................................................... 6
1.5.1 HMG-CoA Reductase Inhibitors (Statins) .................................................................................................... 7
1.5.2 Metformin .................................................................................................................................................. 7
1.5.3 Complement Inhibitors ............................................................................................................................... 7
1.5.4 Stem cell therapy ........................................................................................................................................ 8
1.5.5 Photocoagulation ....................................................................................................................................... 8
1.6 Selective Retinal Therapy ................................................................................................................... 8
1.7 Wound Healing and Extracellular Matrix Remodeling ....................................................................... 9
1.7.1 Metalloproteinases ..................................................................................................................................... 9
1.7.2 TGF- β and Fibrosis ..................................................................................................................................... 9
1.8 Renin-Angiotensin System and Its Role in Wound Healing .............................................................. 10
1.8.1 Classical RAS ............................................................................................................................................. 10
Protective RAS ................................................................................................................................................... 11
Ocular RAS ......................................................................................................................................................... 12
1.9 Hypothesis and Specific Aims ........................................................................................................... 12
1.10 Outline of the Dissertation ............................................................................................................. 13
1.11 Chapter 1 References ..................................................................................................................... 14
Chapter 2: Characterization of Selective Retinal Therapy and Wound Healing Response in Hard
Drusen Mouse Model of Age-Related Macular Degeneration ..................................................... 19
2.1 Introduction ..................................................................................................................................... 19
2.2 Materials and Methods .................................................................................................................... 21
2.2.1 Animals ..................................................................................................................................................... 21
2.2.2 Preparation of Blood Plasma for LC-MS/MS Analysis ............................................................................... 23
2.2.3 Preparation of Eyes ................................................................................................................................... 24
2.2.4 Flat mount Staining ................................................................................................................................... 26
2.2.5 Histopathology ......................................................................................................................................... 27
2.2.6 RT-PCR Analysis ........................................................................................................................................ 27
iv
2.2.7 LC-MS/MS Analysis ................................................................................................................................... 29
........................................................................................................................................................................... 31
2.2.8 Statistical Analysis ..................................................................................................................................... 31
2.3 Results .............................................................................................................................................. 32
2.3.1 Applied Laser Settings .............................................................................................................................. 32
2.3.2 Fundus Images .......................................................................................................................................... 32
2.3.3 Fluorescein Angiography .......................................................................................................................... 33
........................................................................................................................................................................... 35
2.3.5 Flat Mount Staining for Tight Junctions .................................................................................................... 35
2.3.5 H&E Assessment of R:GEN Lesions ........................................................................................................... 36
........................................................................................................................................................................... 37
2.3.6 Fundus Autofluorescence as Measure of Drusen Regression ................................................................... 37
2.3.7 RT-PCR ...................................................................................................................................................... 38
2.3.8 LC-MS/MS ................................................................................................................................................. 41
2.4 Discussion ........................................................................................................................................ 54
2.4.1 R:GEN upregulates ECM turnover in hard drusen (Cx3Cr1 -/-) mouse model .......................................... 55
2.4.2 R:GEN shifts ocular RAS towards pro-regeneration in hard drusen mouse model .................................. 55
2.5 Conclusion ........................................................................................................................................ 58
2.6 Chapter 2 References ....................................................................................................................... 60
Chapter 3: Characterization of Selective Retinal Therapy and Wound Healing Response in Soft
Drusen Mouse Model of Age-Related Macular Degeneration ..................................................... 63
3.1 Introduction ..................................................................................................................................... 63
3.2 Materials and Methods .................................................................................................................... 65
3.2.1 Animals ..................................................................................................................................................... 65
3.2.2 Preparation of Blood Plasma for LC-MS/MS Analysis ............................................................................... 67
3.2.3 Preparation of Eyes ................................................................................................................................... 68
3.2.6 RT-PCR Analysis ........................................................................................................................................ 70
3.2.7 RAS Metabolomics Using Quantitive LC-MS/MS Analysis ........................................................................ 72
3.2.8 Statistical Analysis ..................................................................................................................................... 73
3.3 Results .............................................................................................................................................. 74
3.3.1 Applied Laser Settings .............................................................................................................................. 74
3.3.2 Fundus Images .......................................................................................................................................... 75
3.3.3 Fluorescein Angiography .......................................................................................................................... 76
3.3.4 Flat Mount Staining for Tight Junctions .................................................................................................... 78
3.3.5 H&E Assessment of R:GEN Lesions ........................................................................................................... 79
3.3.6 RT-PCR ...................................................................................................................................................... 80
3.3.7 LC-MS/MS ................................................................................................................................................. 83
3.4 Discussion ...................................................................................................................................... 100
3.4.1 R:GEN upregulates ECM turnover in soft drusen (CxCr5 -/-) mouse model ........................................... 101
3.4.2 R:GEN shifts ocular RAS towards pro-regeneration in soft drusen (CxCr5 -/-) mouse model ................ 102
3.5 Conclusion ...................................................................................................................................... 105
3.6 Chapter 3 References ..................................................................................................................... 106
Chapter 4: Cell-Based Assays for Characterization of Ocular Damage Model ........................... 108
4.1 Introduction ................................................................................................................................... 108
v
4.2 Materials and Methods .................................................................................................................. 109
4.2.1 Cells ........................................................................................................................................................ 109
4.2.2 Angiotensin II Dose Escalation ................................................................................................................ 110
4.2.3 Renin-Angiotensin Inhibition and Stress Induction ................................................................................ 110
4.2.4 RT-PCR Analysis ...................................................................................................................................... 111
4.2.5 LC-MS/MS Analysis ................................................................................................................................. 113
4.2.6 Statistical Analysis ................................................................................................................................... 114
4.3 Results ............................................................................................................................................ 114
4.3.1 Angiotensin II Dose Escalation ................................................................................................................ 114
4.3.2 Cell Injury and RAS Inhibition ................................................................................................................. 117
4.4 Discussion ...................................................................................................................................... 136
4.4.1 Angiotensin II Upregulates Genes in Dose-Response Relationship ........................................................ 137
4.4.2 Hydrogen Peroxide Treatment Alters RAS in ARPE-19 Cells ................................................................... 137
4.5 Conclusion ...................................................................................................................................... 140
4.6 Chapter 4 References ..................................................................................................................... 141
Chapter 5: Summary and Concluding Remarks .......................................................................... 143
References ................................................................................................................................. 149

vi
List of Tables
Table 1.1. Risk factors of AMD ………………………………………………………………………………………………………………..3
Table 1.2. Drusen deposit size as indicator of AMD disease progression…………………………………………………5
Table 2.1. R:GEN laser settings for mice retinal application…………………………………………………………………..25
Table 2.2. Study design for Cx3Cr1-/- hard drusen mice model……………………………………………………………..26
Table 2.3. Protease inhibitor cocktail……………………………………….……………………………………………………………27
Table 2.4. Preparation of buffers for flat mount (Zo-1) staining ……………………………………………………………29
Table 2.5. Primers for the specific gene targets…………………………………………………………………………………….32
Table 2.6. Liquid chromatography parameters………………………………………………………………………………………34
Table 2.7. HPLC gradient pump program……………………………………………………………………………………………….34
Table 2.8. Ion spray settings………………………………………………………………………………………………………………….35
Table 2.9. Lesion severity scoring and criteria for fluorescein angiography images……………………………….37
Table 2.10. Correlation analysis between plasma and ocular RAS peptides…………………………………………..54
Table 2.11. Correlation analysis between FAF drusen counts and ocular RAS components……………………57
Table 3.1. R:GEN laser settings for mice retinal application…………………………………………………………………..71
Table 3.2. Study design for CxCr5-/- soft drusen mice model…………………………………………………………………72
Table 3.3. Lesion severity scoring and criteria for fluorescein angiography images……………………………….81
Table 3.4. Correlation analysis between plasma and ocular RAS………………………………………………………….102
Table 3.5. Area under the curve for ocular RAS peptides…………………………………………………………………….105
Table 4.1. List of RAS inhibitors and respective IC 50’s…………………………………………………………………………..116
Table 4.2. Study design for mechanistic dissection of RAS pathway and impact of tissue injury………....116
Table 4.3. Primers for the specific gene targets…………………………………………………………………………………..118
Table 4.4. Cell media RAS peptides after H 2O 2 treatment with RAS enzyme and receptor inhibition…...138
vii
Table 4.5. One-way ANOVA of RAS peptides after H 2O 2 treatment with RAS enzyme and receptor
inhibition …………………………………………………………………………………………………………………………………………....138

viii
List of Figures
Figure 1.1. Clinical staging of AMD ………………………………………………………………………………………………………….2
Figure 1.2. Metabolic pathway of RAS following activations………………………………………………………………….11
Figure 2.1. Total energy delivered to each treated eye………………………………………………………………………….35
Figure 2.2. Fundus images of Cx3Cr1-/- hard drusen (HD) retina……………………………………………………………36
Figure 2.3. Representative fluorescein angiography of Cx3Cr1-/- hard drusen (HD) retina ……………………37
Figure 2.4. Assessment and scoring of visible lesions in Cx3Cr1-/- mouse at 100% R:GEN intensity……...38
Figure 2.5. Zo-1 staining of isolated Cx3Cr1-/- RPE……...…………………………………………………………………….....39
Figure 2.6. Representative H&E for Cx3Cr1-/- mice….……………………………………………………………………………40
Figure 2.7. Observed changes in fundus autofluorescence counts over time ….…………………………………….41
Figure 2.8. Observed changes in laser-treated RPE gene expression of metalloproteinases ………………….42
Figure 2.9. Observed changes in laser-treated RPE gene expression of RAS components…..………………….43
Figure 2.10. Observed changes in laser-treated RPE gene expression of TGF- β1 and MasR………………... 43
Figure 2.11. Plasma levels of Ang(1-10) after R:GEN treatment……………………………………………………………. 44
Figure 2.12. Plasma levels of Ang(1-9) after R:GEN treatment……………………………………………………………….45
Figure 2.13. Plasma levels of AngII after R:GEN treatment…………………………………………………………………….45
Figure 2.14. Plasma levels of Ang(1-7) after R:GEN treatment……………………………………………………………….46
Figure 2.15. Plasma levels of Ang(2-8) after R:GEN treatment……………………………………………………………….46
Figure 2.16. Plasma levels of Ang(3-8) after R:GEN treatment……………………………………………………………….47
Figure 2.17. Plasma levels of Ang(3-7) after R:GEN treatment……………………………………………………………….47
Figure 2.18. Plasma levels of Ang(1-5) after R:GEN treatment……………………………………………………………….47
Figure 2.19. Ocular metabolism of Ang(1-10) to AngII……….….……………………………………………………………...48
Figure 2.20. Ocular metabolism of Ang(1-10) to Ang(1-9)….….……………………………………………………………...49
Figure 2.21. Ocular metabolism of AngII to Ang(1-7)….….……………………………………………………………………..50
ix
Figure 2.22. Ocular metabolism of Ang(1-9) to Ang(1-7)….….…………………………………………………………….....51
Figure 2.23. Ocular binding of AngII..….………………………………………………………………………….........................52
Figure 2.24. Ocular binding of Ang(1-7)………………………………………………………………………….........................52
Figure 2.25. Ocular binding of Ang(1-9)………………………………………………………………………............................53
Figure 2.26. Aggregate fold change in RAS peptides………..…………………………………………….........................54
Figure 2.27. Correlation between ocular RAS peptide ratios and gene expression of their respective
enzymes in RPE …………………………………………………………………………………………………..…………........................55
Figure 2.28. Correlation between ocular RAS peptide ratios and gene expression of their respective
receptors in RPE …………….…………………………………………………………………………………..…………........................56
Figure 3.1. Total energy delivered to each treated eye………………………………………………………………………….79
Figure 3.2. Fundus images of CxCr5-/- soft drusen (SD) retina….……………………………………………………………80
Figure 3.3. Representative fluorescein angiography of CxCr5-/- soft drusen (SD) retina ……………………….81
Figure 3.4. Assessment and scoring of visible lesions in CxCr5-/- mouse at 100% R:GEN intensity…….....82
Figure 3.5. Assessment and scoring of visible lesions in CxCr5-/- mouse at 50% R:GEN intensity……….....83
Figure 3.6. Zo-1 staining of isolated CxCr5-/- RPE layer.…………………………………………………………………………84
Figure 3.7. Representative H&E for CxCr5-/- mice….……………………………………………………………………………..85
Figure 3.8. Observed changes in laser-treated RPE gene expression of metalloproteinases ………………….86
Figure 3.9. Observed changes in laser-treated RPE gene expression of RAS components…..………………….87
Figure 3.10. Observed changes in laser-treated RPE gene expression of TGF- β1 and MasR………………... 88
Figure 3.11. Plasma levels of Ang(1-12) after R:GEN treatment……………………………………………………………. 88
Figure 3.12. Plasma levels of Ang(1-10) after R:GEN treatment……………………………………………………………. 89
Figure 3.13. Plasma levels of Ang(1-9) after R:GEN treatment……………………………………………………………….90
Figure 3.14. Plasma levels of AngII after R:GEN treatment…………………………………………………………………….90
Figure 3.15. Plasma levels of Ang(1-7) after R:GEN treatment……………………………………………………………… 91
x
Figure 3.16. Plasma levels of Ang(2-8) after R:GEN treatment………………………………………………………….……92
Figure 3.17. Plasma levels of Ang(3-8) after R:GEN treatment………………………………………………………….……92
Figure 3.18. Plasma levels of Ang(3-7) after R:GEN treatment……………………………………………………………….93
Figure 3.19. Plasma levels of Ang(1-5) after R:GEN treatment……………………………………………………………….94
Figure 3.20. Ocular metabolism of Ang(1-10) to AngII……….….………………………………………………………………95
Figure 3.21. Ocular metabolism of Ang(1-10) to Ang(1-9)….….………………………………………………………………96
Figure 3.22. Ocular metabolism of AngII to Ang(1-7)….….……………………………………………………………………..97
Figure 3.23. Ocular metabolism of Ang(1-9) to Ang(1-7)….….…………………………………………………………….....98
Figure 3.24. Ocular binding of AngII..….………………………………………………………………………….........................99
Figure 3.25. Ocular binding of Ang(1-7)………………………………………………………………………….......................100
Figure 3.26. Ocular binding of Ang(1-9)………………………………………………………………………..........................101
Figure 3.27. Aggregate fold change in RAS peptides………..…………………………………………….......................102
Figure 3.28. Correlation between ocular RAS peptide ratios and gene expression of their respective
enzymes in RPE …………………………………………………………………………………………………..…………......................103
Figure 3.29. Correlation between ocular RAS peptide ratios and gene expression of their respective
receptors in RPE …………….…………………………………………………………………………………..…………......................104
Figure 4.1. Gene expression of RAS enzymes after AngII treatment……………………………..……………………..121
Figure 4.2. Gene expression of RAS receptors after AngII treatment…………………………………………………..122
Figure 4.3. Gene expression of metalloproteinases after AngII treatment…………………………………………..122
Figure 4.4. Gene expression changes in angiotensin converting enzymes after H 2O 2 treatment………….125
Figure 4.5. Gene expression changes in RAS receptors after H 2O 2 treatment………………………………………126
Figure 4.6. Gene expression changes in RAS components after hydrogen peroxide treatment after ACE
inhibition……………………………………………………………………………………………………………………………………………. 127
xi
Figure 4.7. Gene expression changes in RAS receptors after hydrogen peroxide treatment after ACE
inhibition……………………………………………………………………………………………………………………………………………. 128
Figure 4.8. Gene expression changes in RAS enzymes after hydrogen peroxide treatment after AT1R
inhibition……………………………………………………………………………………………………………………………………………. 129
Figure 4.9. Gene expression changes in RAS receptors after hydrogen peroxide treatment after ACE
inhibition……………………………………………………………………………………………………………………………………………. 130
Figure 4.10. Gene expression changes in RAS enzymes after hydrogen peroxide treatment after Mas
inhibition……………………………………………………………………………………………………………………………………………. 131
Figure 4.11. Gene expression changes in RAS receptors after hydrogen peroxide treatment after Mas
inhibition……………………………………………………………………………………………………………………………………………. 131
Figure 4.12. Gene expression changes in RAS enzymes after hydrogen peroxide treatment after AT2R
inhibition……………………………………………………………………………………………………………………………………………. 132
Figure 4.13. Gene expression changes in RAS receptors after hydrogen peroxide treatment after AT2R
inhibition……………………………………………………………………………………………………………………………………………. 133
Figure 4.14. Gene expression changes in RAS enzymes after hydrogen peroxide treatment after ACE2
inhibition……………………………………………………………………………………………………………………………………………. 134
Figure 4.15. Gene expression changes in RAS receptors after hydrogen peroxide treatment after ACE2
inhibition……………………………………………………………………………………………………………………………………………. 134
Figure 4.16. Gene expression signals downstream of RAS inhibitor pretreatment ………………………………135
Figure 4.17. Gene expression changes in MMP’s after hydrogen peroxide treatment…………………………136
Figure 4.18. Gene expression changes in MMP’s when inhibited prior to hydrogen peroxide treatment
…………………………………………………………………………………………………………………………………………………………….137
Figure 4.19. Cell media RAS peptide ratios after H 2O 2 treatment with RAS enzyme and receptor
inhibition……………………………………………………………………………………………………………………………………………..139
xii
Figure 4.20. RAS components gene expression and peptides levels after H 2O 2 induction…………………….140
Figure 4.21. Gene expression changes in RAS components in response to H 2O 2 induction, and the impact
of losartan on the RAS metabolism after AT1R inhibition…………………………………………………………………….141
Figure 4.22. Gene expression changes in RAS components with hydrogen peroxide treatment after ACE2
inhibition …………………………..………………………………………………………………………………………………………………..142
Figure 4.23. Gene expression changes in TGF- β with H 2O 2 treatment after RAS inhibition …….…………..143
Figure 4.24. Plot comparing TGF- β1 expression and molar Ang II concentrations of dose escalation
treatment and R:GEN treatment ……………………………………………………………………………………..…….…………..143

xiii
Abstract
Age-related macular degeneration (AMD) is a leading cause of vision loss in the United States.
While VEGF inhibitors are available for treating wet AMD (wAMD), there are currently no FDA-approved
treatments for dry AMD (dAMD). The hallmark of AMD is the presence of drusen, lipid and protein
amalgamations that accumulate in the subretinal space, for which their increase in size and volume is
positively correlated with disease progression. This correlation has made drusen deposits, which are
compositionally similar to atherosclerotic plaques, a primary target in developing treatments for dAMD,
although the cause of accumulation is still unclear. One hypothesis is that Retinal Pigment Epithelium
(RPE), a single layer of cells found in the interface of the blood and photoreceptors, is critical for
maintaining retinal homeostasis, including nutrient/waste transport and its elimination, becomes
dysfunctional with age. Selective Retinal Therapy (SRT), is a treatment proposed for dAMD, for which
subthreshold nano-lasers are used to selectively destroy RPE cells and initiate mechanisms of wound
repair. SRT lasers have previously been shown to reduce drusen deposits, although the underlying
mechanism beyond extracellular matrix turnover is unclear. The Renin-Angiotensin System (RAS) has
previously been shown to play a role in wound healing from tissue (e.g., traumatic, thermal, or radiation
induced) injury and atherosclerosis, but a gap in the knowledge for whether stimulation of ocular RAS
can be beneficial for drusen reduction in dAMD. We utilized R:GEN, a proprietary SRT laser, to
demonstrate its safety and in vivo efficacy using two genetic knockout (KO) mouse models of AMD, and
explore its ability to stimulate ocular RAS as part of its drusen reduction mechanism.
Previously conducted dose-escalation studies in C57/BL mice indicated R:GEN to be safe and
capable of upregulating enzymes and receptors of the RAS pathway. Utilizing the established laser
intensities, R:GEN was shown in a time course studies to selectively destroy the RPE of AMD mouse
models, reduce drusen-like deposits, and stimulate production of RAS peptides. Negative correlations
were identified between systemic and ocular peptides, demonstrating the potential for liquid biopsies
xiv
predictive of safety and efficacy. Surprisingly, gene expression of RAS enzymes was downregulated in
the RPE of diseased mice, suggesting the laser intensities used in healthy C57/BL mice may be too high
for diseased animals. This observation was underscored by comparing the differences in RAS peptides
area under the curve (AUC) between different laser intensities. Despite these findings, reduction in
drusen was still found to be weakly negatively correlated with RAS components, although additional
“dose” refinement may be necessary to maximize efficacy in dAMD patients.
To further investigate the effect of tissue damage on ocular RAS, in vitro studies were conducted
using injury induced-ARPE-19 cell line with and without inhibitors specific to RAS enzymes or receptors.
Gene expression was correlated with RAS metabolomics to provide interactional insights as how
modulation of RAS may dictate wound healing processes and provide additional evidence to support
R:GEN’s claims of inducing minimally fibrotic injury. Finally, the effects of enalapril and losartan, two
FDA-approved therapeutics, on gene expression and RAS metabolomics after cellular injury indicate a
shift towards the protective arm of RAS.
These studies demonstrate the safety of R:GEN-induced lesions in the eye, and confirm the RAS
pathway to be activated by the laser. This body of work provides evidence that RAS modulation may be
beneficial for drusen reduction and dAMD and establishes the foundation for probing combination
therapy.

1
Chapter 1: Introduction to Age-Related Macular Degeneration, Subthreshold Nano-
lasers, and Wound Healing
1.1 Epidemiology of AMD
Age-related macular degeneration (AMD) is the leading cause of blindness for individuals 50 and
older in the developed world (Velez-Montoya et al., 2014). This retinal disorder is a debilitating disease
characterized by drusen, accumulating lipids and proteins found in the subretinal space. The growth of
these deposits is correlated with disease progression and eventual loss of central vision (Girmens, Sahel,
& Marazova, 2012). In the United States, the direct cost of AMD treatment in 2004 was approximately
$525 million, with the prevalence expected to increase to 20 million by 2050 as the population continues
to age (Rein et al., 2009).
Advanced stage AMD is classified into two forms, where approximately 10% to 20% of patients
have exudative or “wet” AMD (Ambati & Fowler, 2012). Wet AMD (wAMD) is identified by the
development of choroidal neovascularization (CNV) with fluid leakage, exudation, hemorrhaging,
detachment of retinal pigment epithelium (RPE), scarring, and fast progression towards blindness (Velez-
Montoya et al., 2014). Currently, anti-vascular endothelial growth factor (anti-VEGF) agents, which block
the formation of new blood vessels, are the standard of care for treating wAMD (Li et al., 2018; Ruan,
Jiang, & Gericke, 2021).
The “dry” or nonexudative form of AMD is characterized by the slow, progressive degeneration
of RPE, eventually leading to Geographic Atrophy (GA), the most severe form of dAMD evidenced by the
death of the photoreceptors (PR) which reside on the apical surface of the RPE layer (Buschini et al., 2015;
Velez-Montoya et al., 2014). There is currently no effective treatment for nonexudative or “dry” AMD,
which accounts for 80% of all AMD cases (Ambati & Fowler, 2012; Wright & Ambati, 2017). The clinical
progression of AMD is summarized in Figure 1.1 below.

2

1.2 Risk Factors and Pathogenesis of AMD
In a systematic review conducted by Wang et al, risk factors related with global incidence of AMD
included advancement of age, smoking, alcohol consumption, and elevated HDL-C. (Y. Wang et al., 2021).
On a molecular level, genome-wide associate studies (GWAS) indicate that variants in the complement
factor H (CFH) gene have strong association in GA development (van Lookeren Campagne, LeCouter,
Yaspan, & Ye, 2014). Further GWAS conducted by the AMD Gene Consortium identified an additional 17
other genetic loci associated with increased risk for AMD, as summarized in Table 1.1 below (van Lookeren
Campagne et al., 2014). As evidenced from Table 1.1, many of the identified pathways linked to
development of AMD are related to extracellular matrix turnover, lipid metabolism, cell death, and the
immune system. In addition, Zarbin provides a landscape overview underscoring the multiple overlapping
cellular pathways related to the pathogenesis of AMD, including oxidative stress, chronic inflammation,
altered extra-cellular matrix remodeling, and retinal pigment epithelium (RPE) dysfunction (Zarbin, 2004).

Figure 1.1: Clinical Staging of AMD where disease advancement is related to the level of
drusen disposition which correlated with the demise of photoreceptors. Adapted from
(Ruan, 2021)
Int. J. Mol. Sci.2021, 22,1296 4of22
choroidalneovascularization(CNV),whichgoesalongwithbloodandfluidleakage,lead-
ingtodetachmentoftheretinaorRPEandrapidvisionloss[41]. Figure2describesthe
clinicalmanifestationandpathologyofAMDfromearlytolatestage.
Table1. TheBeckmanclinicalclassificationofage-relatedmaculardegeneration(AMD)[41].
Classification ClinicalManifestation
NoAMD NodrusenandnoRPEabnormalities
Normalagingchanges Drusen 63µmandnoRPEabnormalities
EarlyAMD Drusen>63µmand 125µmandnoRPEabnormalities
IntermediateAMD Drusen>125µmand/orRPEabnormalities
LateAMD GAand/orneovascularAMD
Abbreviations: AMD:age-relatedmaculardegeneration;GA:geographicatrophy;RPE:retinalpigmentepithe-
lium.
Int. J. Mol. Sci. 2021, 22, x FOR PEER REVIEW 4 of 22

vessels growing from the choroid into the retina [41]. This process is known as choroidal
neovascularization (CNV), which goes along with blood and fluid leakage, leading to de-
tachment of the retina or RPE and rapid vision loss [41]. Figure 2 describes the clinical
manifestation and pathology of AMD from early to late stage.
Table 1. The Beckman clinical classification of age-related macular degeneration (AMD) [41].
Classification Clinical Manifestation
No AMD No drusen and no RPE abnormalities
Normal aging changes Drusen 63 m and no RPE abnormalities
Early AMD Drusen >63 m and 125 m and no RPE abnormalities
Intermediate AMD Drusen > 125 m and/or RPE abnormalities
Late AMD GA and/or neovascular AMD
Abbreviations: AMD: age-related macular degeneration; GA: geographic atrophy; RPE: retinal
pigment epithelium.

Figure 2. Clinical manifestation and pathology of AMD from the early to late stage. Medium-sized
drusen found in early AMD. Intermediate AMD shows the presence of large drusen. Late AMD is
classified into GA and neovascular AMD. GA is defined by the deterioration of the RPE, photore-
ceptor layer, and choroidal capillaries in the macula. The invasion of abnormal fragile choroidal
blood vessels growing from the choroid into the retina in neovascular AMD, with blood and fluid
leakage. Abbreviations: AMD: age-related macular degeneration; GA: geographic atrophy.
4. Pathogenesis of AMD
4.1. Oxidative Stress and AMD
4.1.1. The Macula—An Ideal Environment for the Generation of ROS
It is well known that the retina is one of the highest oxygen-consuming tissues in the
human body, utilizing even more oxygen per weight than the brain [42]. The local oxygen
metabolic environment in the retina plays an essential role in keeping retinal homeostasis
between the supply and consumption of retinal oxygen [42]. The retina continuously
transforms light into vision, requiring a marked amount of energy and generating reactive
oxygen species (ROS), such as the superoxide (O2
• ), the hydroxyl radical (
•
OH), hydrogen
peroxide (H2O2), and singlet oxygen (
1
O2) as normal metabolic byproducts [42]. Generally,
ROS are produced during oxidative metabolism under physiological conditions and par-
ticipate in normal cellular metabolism [43]. However, when the generation of ROS exceeds
Figure2. ClinicalmanifestationandpathologyofAMDfromtheearlytolatestage. Medium-sizeddrusenfoundinearly
AMD.IntermediateAMDshowsthepresenceoflargedrusen. LateAMDisclassifiedintoGAandneovascularAMD.GA
isdefinedbythedeteriorationoftheRPE,photoreceptorlayer,andchoroidalcapillariesinthemacula. Theinvasionof
abnormalfragilechoroidalbloodvesselsgrowingfromthechoroidintotheretinainneovascularAMD,withbloodand
fluidleakage. Abbreviations: AMD:age-relatedmaculardegeneration;GA:geographicatrophy.
4. PathogenesisofAMD
4.1. Oxidative Stress and AMD
4.1.1. TheMacula—AnIdealEnvironmentfortheGenerationofROS
Itiswellknownthattheretinaisoneofthehighestoxygen-consumingtissuesinthe
humanbody,utilizingevenmoreoxygenperweightthanthebrain[42]. Thelocaloxygen
metabolicenvironmentintheretinaplaysanessentialroleinkeepingretinalhomeostasis
between the supply and consumption of retinal oxygen [42]. The retina continuously
transformslightintovision,requiringamarkedamountofenergyandgeneratingreac-
tive oxygen species (ROS), such as the superoxide (O
2
• ), the hydroxyl radical (
•
OH),
hydrogenperoxide(H
2
O
2
),andsingletoxygen(
1
O
2
)asnormalmetabolicbyproducts[42].
Generally,ROSareproducedduringoxidativemetabolismunderphysiologicalconditions
and participate in normal cellular metabolism [43]. However, when the generation of
ROS exceeds the capacity of the antioxidant systems, ROS disrupt the balance of redox
homeostasisandcauseoxidativestress[22].

3

1.3 Retinal Pigment Epithelium and Age-Related Dysfunction
Retinal pigment epithelium (RPE) are hexagonal cells that forms “cobble stone” morphology, is a
cellular barrier playing multiple roles in maintaining retinal homeostasis (Kim, Lee, & Won, 2021; Sharma,
Bose, Maminishkis, & Bharti, 2020). Because RPE are responsible wide spectrum of cellular functions as
listed below, where RPE dysfunction can lead to AMD development:
1. Maintenance of the blood-retinal-barrier via the formation of tight junctions
Chr. Gene/Locus Pathway Function
10 ARMS2/HTRA1 ECM
ARMS2 - Linked to extracellular matrix; HTRA1 –
regulator of cell growth
1 CFH Complement Negative regulation of complement pathway
6 C2/CFB Complement CFB - Associated with C3b; C2 cleaved to create C3
19 C3 Complement Regulation of complement pathway
22 TIMP3 ECM Inhibitor of matrix metalloproteinase-3
19 APOE Lipid metabolism Lipid transporter
16 CETP Lipid metabolism Transfers cholesterol esters between lipoproteins
6 VEGFA Angiogenesis
Induces angiogenesis and vasculogenesis,
promotes cell migration
8 TNFRSF10A Apoptosis Induces apoptosis
15 LIPC Lipid metabolism
Functions in triglyceride hydrolase and receptor-
mediated lipid lipoprotein uptake
4 CFI Complement
Regulation of complement pathway via
inactivation of C3b and C4b
6 COL10A1 ECM Encodes alpha chain of Type X collagen
3 COL8A1/FILIIP1L ECM; Angiogenesis
COL8A1 – Major component of basement
membrane of corneal epithelium; FILIP1L –
regulator of angiogenesis on epithelial cells
6 IER3/DDR1 Apoptosis; ECM
IER3 – protection of cells from TNF-a induced
apoptosis; DDR1 – receptor tyrosine kinase
involved in regulation of cell growth,
differentiation, and metabolism
22 SLC16A8 Other Mediates lactate transport across cell membrane
9 TGFBR1 Other
Signal transduction of TGF-B from cell surface to
cytoplasm
14 RAD51B Apoptosis Essential for DNA repair
3 ADAMTS9 Apoptosis Implicated in the cleavage of proteoglycans
13 B3GALTL Other Transfer of glucose to O-linked fucosylglcans
Table 1.1: Risk factors of AMD ( adapted from van Lookeren Campagne, 2014)

4
2. Transportation of nutrients into the retinal in exchange for waste into the vasculature
3. Phagocytosis of photoreceptors molecular and cellular waste
4. Absorption of light and protection from oxidative stress
5. Secretion of cytokines to maintain blood vessel confluency and choroid permeability
6. Secretion of metalloproteinases and metalloproteinase inhibitors
Aging is commonly associated with the accumulation of free radicals and oxidative damage
(Harman, 1956). Reactive oxygen species (ROS) and advanced glycation end (AGE) products associated
with aging can induce cell injury and dysregulated tissue remodeling by increasing the deposits of extra-
cellular matrix, resulting in the thickening Bruch’s membrane characteristic of AMD (Burns & Feeney-
Burns, 1980). Because of this observed thickening, it is believed that the aged Bruch’s membrane retains
its ability to synthesize extracellular matrix, but unable to degrade the matrix (Karwatowski et al., 1995).
In addition to the Bruch’s membrane, the RPE may be susceptible to oxidative damage with age,
interfering with lysosomal activity and increasing the buildup of toxic waste products (Zarbin, 2004). A
major contributor to the RPE’s defense against oxidative damage include the super-oxide dismutase
(SOD), glutathione peroxidase (GPx), and catalase (Cat), where decreased expression has been associated
in aged RPE and AMD (Zarbin, 2004). The inability to effectively neutralize ROS has been linked to the
peroxidation of lipids that accumulate in aged RPE, reducing the photoreceptor outer segment (OS) from
eliminating waste, and creating a positive feedback loop that continues accumulate debris (Kaemmerer,
Schutt, Krohne, Holz, & Kopitz, 2007).

5
1.4 Drusen as a Clinical Biomarker of AMD
AMD pathogenesis is often attributed to a combination of the previously mentioned factors.
However, these risk factors commonly manifest in the formation of drusen deposits, which are subretinal
amalgamations of extracellular material (Burns & Feeney-Burns, 1980). Drusen accumulation has been
highly correlated with progression of AMD, in particular dAMD, as summarized in Table 1.2 (Burns &
Feeney-Burns, 1980).

Drusen accumulation is an important factor in AMD disease progression which can cause hypoxia
and triggering ocular inflammation in the affected region (Nita, Grzybowski, Ascaso, & Huerva, 2014).
Drusen accumulation is thought to disrupt tissue oxygenation, reduce oxygen-carbon dioxide exchange,
and ocular metabolism leading to oxidative stress capable of triggering tissue injury and inflammatory
processes. All of these factors have been associated with AMD pathogenesis and disease progression
(Donoso, Kim, Frost, Callahan, & Hageman, 2006). As such, research efforts in trying to understand the
molecular events leading to drusen formulation and how cellular debris can interact with the retinal
microenvironment.
1.4.1 Lipids in Drusen
Insights into these aspects will allow for strategic therapeutic development and tactics to prevent
disease progression. The lipid composition of drusen deposits have long been confirmed using Oil red O
staining and confirmed with electron microscopy analysis revealing the presence of both esterified and
unesterified cholesterol (Curcio, Millican, Bailey, & Kruth, 2001; Wolter & Falls, 1962). Oxidized lipids
Disease Severity Drusen Size Number of Drusen Location
Early AMD 63-124 µm More than 5, less than 20 Does not extend to macula
Intermediate AMD ³ 125 µm At least 1 Does not extend to macula
Advance (Late) AMD Extends to macula
Table 1.2: Drusen deposit size as indicator of AMD disease progression (Ferris et al., 2005)

6
increased within the photoreceptors (PRs) and RPE with age causing lipid accumulation in combination
with oxidative stress (M. Suzuki et al., 2007). Because oxidation of lipoproteins is a critical catalyst in the
formation of atherosclerotic-like plaques, it has been suggested that the mechanism of drusen formation
mirror that of plaque formation; embedding lipoproteins in the Bruch’s membrane (Curcio, Johnson,
Huang, & Rudolf, 2010).
1.4.2 Proteins in Drusen
Although lipids, cholesterols, and their metabolites have been associated with AMD, proteomic
analyses have identified a variety of proteins, such as vitronectin, lipoproteins, tissue metalloproteinase
inhibitor-3 (TIMP-3), complement C9, clusterin, and serum albumin (Crabb et al., 2002). The presence of
complement system components in drusen, it is possible that the debris accumulated in drusen deposits
can amplify the inflammatory cascade and creating a positive feedback loop to drive disease progression.
The complement system is a complex series of protein assembly leading to inflammatory responses
against infectious intrusion (Janeway CA Jr, 2001). Comprised of three arms, the complement system
provides innate immune response: 1) the classical pathway in which antibodies bind to antigens; 2) the
lectin pathway activated by carbohydrates and oxidative stress and; 3) the alternative pathway triggered
by pathogens and cellular debris (Sivaprasad & Chong, 2006).
1.5 Current Therapies In Development for Dry AMD
Currently, there are no FDA approved therapies for the management of dAMD. The only
therapeutic options to prevent disease progression include lifestyle modifications, such as smoking
cessation and dietary supplements (Wright & Ambati, 2017). In the Age-Related Eye Disease Study
(AREDS), a multicenter randomized clinical trial involving more than 3,600 patients who received
antioxidant vitamins and mineral supplementation, including vitamin C, E and A. In addition, zinc oxide,
and cupric oxide was also included in the supplementation study, where risk for developing advanced
AMD for individuals with at least one large druse was reduced (Zarbin, 2004).

7

1.5.1 HMG-CoA Reductase Inhibitors (Statins)
There are parallel phenotypes similarities between atherosclerotic plaques and AMD-related
drusen. These findings led to evaluating the impact of pharmacological interventions to prevent plague
formation as potential AMD therapies. One open-label clinical study involving 26 patients receiving high
dose atorvastatin (80 mg daily), an HMG-CoA reductase inhibitor that reduce systemic cholesterol
biosynthesis, was able to regress ocular drusen in 10 patients (38%) corresponding to visual acuity
improvements (Vavvas et al., 2016). While statistical significance of the study was not met, the results
underscored the critical role cholesterol plays in the development of drusen and subsequent vision loss.
1.5.2 Metformin
Metformin has been proposed to delay age-related diseases associated with its antioxidant and
anti-inflammatory activity (Barzilai, Crandall, Kritchevsky, & Espeland, 2016; Fang et al., 2018). A meta-
analysis of five studies conducted by Romdhoniyyah et al suggested that metformin use was associated
with a decreased risk of developing AMD. While three of the studies found odds ratios to support the use
of metformin for reducing AMD risk, statistical significance was not met. The trends suggest some benefit
with metformin-mediated suppression of the respiratory chain and inflammation, identifying a possible
treatment to be further studied for slowing the development of AMD (Romdhoniyyah, Harding, Cheyne,
& Beare, 2021).
1.5.3 Complement Inhibitors
As previously mentioned, complement factor subunits have been found embedded in drusen in
proteomic analysis. This has led the field to believe that controlling the alternative complement pathway
might be critical to prevent damage to “self” tissues in controlling progression of the disease (Gehrs,
Anderson, Johnson, & Hageman, 2006). As of this writing, inhibitors for complement factors H and C5
have progressed in clinical trials as potential therapies for dAMD. Currently, at least two complement

8
inhibitor candidates have shown reductions in GA lesions, while other study results lacked efficacy,
highlighting the multifactored complexity of this disease (Qin et al., 2021).
1.5.4 Stem cell therapy
More recently, the replacement of damaged RPE with stem cells have been an intense area of
interest for AMD. Currently, there are two main methods for cellular implantation pursued in clinical
studies, 1) cell suspension and 2) polarized cellular sheets (Baradaran-Rafii et al., 2020). Although
preliminary results in clinical studies have been promising, major concerns regarding cell-based therapies
primarily involve cell proliferation and tissue rejection, as at least one clinical trial has reported a high rate
of retinal detachment from their experimental treatment (Rubner, Li, & Canto-Soler, 2022).
1.5.5 Photocoagulation
Commonly used for diabetic retinopathy, photocoagulation raises the temperature of RPE cells
through the absorption of laser light energy, leading to cellular lyses and coagulation of RPE, PRs, and the
Bruch’s Membrane (Querques et al., 2018). While such damage is fairly severe, drusen regression has
commonly been reported as unintended effect, suggesting it to be a potential therapy for dAMD (Virgili,
Michelessi, Parodi, Bacherini, & Evans, 2015).
1.6 Selective Retinal Therapy
While previous photocoagulation results would suggest laser-induced lesions to the back of the
eye are a potential method to treat dAMD, macular scarring is also a known adverse effect of
photocoagulation, resulting in a sub-optimal treatment (Kotoula, Chatzoulis, Tahmitzi, & Tsironi, 2009;
Maeshima, Utsugi-Sutoh, Otani, & Kishi, 2004). To mitigate the multitude of side effects related to the
excessive thermal damage associated with photocoagulation, conventional laser lesions were applied to
the periphery to avoid the central macula when exploring drusen regression techniques (Rodanant et al.,
2002).

9
Modifications have since been made to laser technology to improve the safety for ophthalmic
applications. By utilizing subthreshold laser technology, selective retinal therapy (SRT) targets RPE without
inducing trauma to the surrounding photoreceptors or Bruch’s membrane (Querques et al., 2018). SRT
utilizes a 532 nm wavelength to selectively target the melanosomes in the RPE, quickly pulsing light to
form microbubbles that selectively disrupt cell membranes without inducing visible signs of damage
(Neumann & Brinkmann, 2005).
1.7 Wound Healing and Extracellular Matrix Remodeling
1.7.1 Metalloproteinases
Metalloproteinases (MMPs) are a family of gelatinases known to engage in remodeling of the
extracellular matrix (ECM) (Salo, Makela, Kylmaniemi, Autio-Harmainen, & Larjava, 1994). Wound healing
is a coordinated and complex series of processes, during which 1) clotting of the wound occurs, 2) followed
by inflammation, 3) clearance of pathogens and necrotic debris, and 4) repair through ECM synthesis and
remodeling (Martin, 1997). RPE produce MMPs, with inactive MMP-2 and MMP-9 accumulating in the
Bruch’s membrane with aging, resulting in the thickening observed in AMD (Zarbin, 2004). It is
hypothesized that the observed drusen regression commonly reported after photocoagulation treatment
is due to MMP activation secreted by the RPE during tissue reparative and regeneration processes (Richert
et al., 2018). SRT showed specific upregulation of MMP-2 and MMP-3 in the RPE of treated mice,
potentially mitigating one of the underlying causes of AMD (Jobling et al., 2015).
1.7.2 TGF- β and Fibrosis
Transforming growth factor-beta (TGF- β), a family of cytokines, is a master regulator of fibrosis,
where dysregulation of this signaling pathway can drive the normal wound healing response to fibrosis
(Meng, Nikolic-Paterson, & Lan, 2016). While TGF-β is produced as part of the normal wound healing
response, excessive amounts contribute to pathological fibrosis seen in diseases of different organ
systems (Branton & Kopp, 1999). It is understood that the failure of negative feedback mechanisms to

10
terminate the TGF-β-driven proliferation and production of ECM leads to the endless healing that leads
to fibrosis (Eickelberg, 2001). Therefore, the persistent upregulation of TGF-β may promote pro-fibrotic
healing. Additionally, it is known that Angiotensin II (Ang II) modulates fibrosis in kidney disease and also
induces TGF- β when it binds and activate Angiotensin II Receptor Sub-type 1 (AT1R) (Daniel, 2008). These
findings would indicate that the Renin-Angiotensin System (RAS) may play a role in determining either
fibrotic or non-fibrotic wound healing.
1.8 Renin-Angiotensin System and Its Role in Wound Healing
1.8.1 Classical RAS
The RAS has been associated with maintaining hemodynamics balance. Within the system,
Angiotensin I (Ang I) is cleaved by angiotensin converting enzyme (ACE) to form Ang II where its binding
onto AT1R promotes vasoconstriction, oxidative stress, and inflammation (Mehta & Griendling, 2007).
Less is known about the role RAS plays in wound healing. Ang II was found to accelerate the wound
healing process when activating AT1R (Takeda, Katagata, Hozumi, & Kondo, 2004). However, because Ang
II is commonly known to participate in pathogenesis of diseases, including chronic kidney disease, where
activation of AT1R can promote pro-fibrotic healing (Y. Suzuki, Ruiz-Ortega, & Egido, 2000). Furthermore,
Ang II have been found to promote atherosclerosis, while the administration of angiotensin receptor
blockers (ARBs) can reduce atherosclerosis (Kon & Jabs, 2004; Ramadan et al., 2016). It could then be
surmised from the literature the ACE/Ang II/AT1R axis may play a role in drusen formation. This hypothesis
is further underscored by the evidence that long-term use of RAS inhibitors was found to be beneficial for
early and late stage AMD patients (Ren, Liu, Yin, Zhang, & Lu, 2020).

11
Protective RAS
Additional research into RAS identified an alternative pathway, also known as the “Protective”
arm. Ang II had a second receptor, AT2R, where Ang II binding triggers non-fibrotic healing (Chiu et al.,
1989). Furthermore, Ang II can also be metabolized by ACE2, forming Ang(1-7), which binds to Mas
receptor (Rabelo, Alenina, & Bader, 2011). It was found that the Ang II/AT2R and ACE2/Ang(1-7)/MasR
axes counter-balance the classical or “Pathogenic” RAS arm, and thus called the “Protective Arm” that
promotes vasodilation, anti-inflammation, and anti-fibrotic wound healing (R. A. S. Santos et al., 2019).
Previous studies have shown that RAS peptides are protective when upregulated in dermal tissue from
radiation and thermal burns, for which tissue regeneration has been observed through the activity of its
metabolites Ang II and Ang(1-7) (Jadhav et al., 2013). Because lasers are focused beams of radiation
energy, the findings by Jadhav et al lend credence to the possibility that SRT might stimulate an increase
in ocular RAS peptides.

Figure 1.2: Metabolic pathway of RAS following activations (Adapted from Reddy et al)
AT1 AT2
Angiotensinogen
Angiotensin I (1-10)
Angiotensin II (1-8)
Renin
Prorenin
• Vasoconstriction
• Proliferative
• Hypertrophy
• Aldosterone release
• Oxidative stress
• Insulin resistance
• Vasodilation
• Neurite outgrowth
• Anti-proliferative
• Anti-remodeling
• Neuronal excitability
• Cerebral protective
ACE
Prorenin
Receptor
• Vasodilation
• Anti-proliferative
• Antigrowth
• Diuresis
• Anti-inflammatory
• Anti-remodeling
• Learning
• Memory
• Long-term potentiation
• Glucose uptake
• Blood flow
APA-N
Angiotensin (1-7)
AT4
Angiotensin (1-9)
Angiotensin III (2-8)
Angiotensin IV (3-8)
ACE2 APA-A
Angiotensin (3-7)
ACE2
Mas

12
Ocular RAS
While viewed as the hormonal system regulating blood pressure and electrolyte/fluid balance, it
is also believed to play a role in maintaining osmotic pressure in the eye. Studies have confirmed that
there are significant levels of Ang II in the eye, and surprisingly, experimental results indicate that
angiotensin peptides are produced locally at levels greater than that found in plasma, suggesting local
production (Danser et al., 1994; Van Haeringen, 1996). Ang II is able to induce MMPs to destabilize arterial
plaques (Browatzki et al., 2005). If the similarities between arterial plagues and drusen can be drawn, this
observed MMP activity in the eye may be explained by laser induced upregulation of RAS peptides,
suggesting that RAS peptides may be a potential method for determining SRT‘s ability to promote wound
healing, tissue regeneration, and treatment of AMD.
1.9 Hypothesis and Specific Aims
The RAS is known to play a role within the wound healing process as well as in the promotion and
regression of atherosclerotic plaques, thus leading to the hypothesis that R:GEN upregulates the pro-
regenerative arm of RAS, which will promote the drusen regression phenomenon observed after
photocoagulation and SRT.
R:GEN is an SRT laser with unique dual-dosimetry technology, utilizing opto-acoustics and
reflectometry to identify the exact moment of microbubble formation, resulting in tightly regulated
energy delivery proposed to further improve the safety and efficacy of SRT. In prior studies, R:GEN was
shown to selectively target the RPE of healthy C57B/L mice, as well as upregulate the gene expression of
RAS enzymes and receptors, although the changes in the actual peptides is not known.
This project focused on determining the safety and efficacy of R:GEN in AMD, using both hard and
soft drusen rodent models. Throughout this project, I intended to identify an upper threshold of laser
energy that could be used safely in the eye using a combination of histological and molecular techniques,
and characterize the impact that R:GEN would have on producing pro-regenerative RAS peptides

13
hypothesized to promote efficacy and drusen regression. The project was culminated in the following
specific aims (SA):
SA1: Evaluate safety and efficacy of R:GEN in hard drusen (Cx3Cr1 -/-) mouse model.
SA2: Evaluate safety and efficacy of R:GEN in soft drusen (CxCr5 -/-) mouse model.
SA3: Characterize the molecular events in ARPE-19 cell damage model with and without RAS inhibition.
1.10 Outline of the Dissertation
This dissertation characterized the ocular molecular events after SRT treatment in a hard drusen
mouse model (Chapter 2), followed by the characterization of SRT in a soft drusen mouse model (Chapter
3). To confirm these in vivo wound healing responses, we designed an RPE cell injury model with or
without the use of RAS inhibitors (Chapter 4).

14
1.11 Chapter 1 References

Ambati, J., & Fowler, B. J. (2012). Mechanisms of age-related macular degeneration. Neuron, 75(1), 26-
39. doi:10.1016/j.neuron.2012.06.018

Baradaran-Rafii, A., Sarvari, M., Alavi-Moghadam, S., Payab, M., Goodarzi, P., Aghayan, H. R., . . .
Arjmand, B. (2020). Cell-based approaches towards treating age-related macular degeneration.
Cell Tissue Bank. doi:10.1007/s10561-020-09826-3

Barzilai, N., Crandall, J. P., Kritchevsky, S. B., & Espeland, M. A. (2016). Metformin as a Tool to Target
Aging. Cell Metab, 23(6), 1060-1065. doi:10.1016/j.cmet.2016.05.011

Branton, M. H., & Kopp, J. B. (1999). TGF-beta and fibrosis. Microbes Infect, 1(15), 1349-1365.
doi:10.1016/s1286-4579(99)00250-6

Browatzki, M., Larsen, D., Pfeiffer, C. A., Gehrke, S. G., Schmidt, J., Kranzhofer, A., . . . Kranzhofer, R.
(2005). Angiotensin II stimulates matrix metalloproteinase secretion in human vascular smooth
muscle cells via nuclear factor-kappaB and activator protein 1 in a redox-sensitive manner. J
Vasc Res, 42(5), 415-423. doi:10.1159/000087451

Burns, R. P., & Feeney-Burns, L. (1980). Clinico-morphologic correlations of drusen of Bruch's
membrane. Trans Am Ophthalmol Soc, 78, 206-225.

Buschini, E., Fea, A. M., Lavia, C. A., Nassisi, M., Pignata, G., Zola, M., & Grignolo, F. M. (2015). Recent
developments in the management of dry age-related macular degeneration. Clin Ophthalmol, 9,
563-574. doi:10.2147/OPTH.S59724

Chiu, A. T., Herblin, W. F., McCall, D. E., Ardecky, R. J., Carini, D. J., Duncia, J. V., . . . Timmermans, P. B.
M. W. M. (1989). Identification of angiotensin II receptor subtypes. Biochemical and Biophysical
Research Communications, 165(1), 196-203. doi:https://doi.org/10.1016/0006-291X(89)91054-1

Crabb, J. W., Miyagi, M., Gu, X., Shadrach, K., West, K. A., Sakaguchi, H., . . . Hollyfield, J. G. (2002).
Drusen proteome analysis: an approach to the etiology of age-related macular degeneration.
Proc Natl Acad Sci U S A, 99(23), 14682-14687. doi:10.1073/pnas.222551899

Curcio, C. A., Johnson, M., Huang, J. D., & Rudolf, M. (2010). Apolipoprotein B-containing lipoproteins in
retinal aging and age-related macular degeneration. J Lipid Res, 51(3), 451-467.
doi:10.1194/jlr.R002238

Curcio, C. A., Millican, C. L., Bailey, T., & Kruth, H. S. (2001). Accumulation of cholesterol with age in
human Bruch's membrane. Invest Ophthalmol Vis Sci, 42(1), 265-274.

Daniel, C. (2008). Blocking of angiotensin II is more than blocking of transforming growth factor-beta.
Kidney Int, 74(5), 551-553. doi:10.1038/ki.2008.290

Danser, A. H., Derkx Fh Fau - Admiraal, P. J., Admiraal Pj Fau - Deinum, J., Deinum J Fau - de Jong, P. T.,
de Jong Pt Fau - Schalekamp, M. A., & Schalekamp, M. A. (1994). Angiotensin levels in the eye.
(0146-0404 (Print)).

15
Donoso, L. A., Kim, D., Frost, A., Callahan, A., & Hageman, G. (2006). The role of inflammation in the
pathogenesis of age-related macular degeneration. Surv Ophthalmol, 51(2), 137-152.
doi:10.1016/j.survophthal.2005.12.001

Eickelberg, O. (2001). Endless healing: TGF-beta, SMADs, and fibrosis. FEBS Lett, 506(1), 11-14.
doi:10.1016/s0014-5793(01)02875-7

Fang, J., Yang, J., Wu, X., Zhang, G., Li, T., Wang, X., . . . Wang, L. (2018). Metformin alleviates human
cellular aging by upregulating the endoplasmic reticulum glutathione peroxidase 7. Aging Cell,
17(4), e12765. doi:10.1111/acel.12765

Ferris, F. L., Davis, M. D., Clemons, T. E., Lee, L. Y., Chew, E. Y., Lindblad, A. S., . . . Age-Related Eye
Disease Study Research, G. (2005). A simplified severity scale for age-related macular
degeneration: AREDS Report No. 18. Arch Ophthalmol, 123(11), 1570-1574.
doi:10.1001/archopht.123.11.1570

Gehrs, K. M., Anderson, D. H., Johnson, L. V., & Hageman, G. S. (2006). Age-related macular
degeneration--emerging pathogenetic and therapeutic concepts. Ann Med, 38(7), 450-471.
doi:10.1080/07853890600946724

Girmens, J. F., Sahel, J. A., & Marazova, K. (2012). Dry age-related macular degeneration: A currently
unmet clinical need. Intractable Rare Dis Res, 1(3), 103-114. doi:10.5582/irdr.2012.v1.3.103

Harman, D. (1956). Aging: a theory based on free radical and radiation chemistry. J Gerontol, 11(3), 298-
300. doi:10.1093/geronj/11.3.298

Jadhav, S. S., Sharma, N., Meeks, C. J., Mordwinkin, N. M., Espinoza, T. B., Roda, N. R., . . . Rodgers, K. E.
(2013). Effects of combined radiation and burn injury on the renin-angiotensin system. Wound
Repair Regen, 21(1), 131-140. doi:10.1111/j.1524-475X.2012.00867.x

Janeway CA Jr, T. P., Walport M, et al. (2001). The complement system and innate immunity. In G.
Science (Ed.), Immunobiology: The Immune System in Health and Disease. 5th edition. New
York.

Jobling, A. I., Guymer, R. H., Vessey, K. A., Greferath, U., Mills, S. A., Brassington, K. H., . . . Fletcher, E. L.
(2015). Nanosecond laser therapy reverses pathologic and molecular changes in age-related
macular degeneration without retinal damage. FASEB J, 29(2), 696-710. doi:10.1096/fj.14-
262444

Kaemmerer, E., Schutt, F., Krohne, T. U., Holz, F. G., & Kopitz, J. (2007). Effects of lipid peroxidation-
related protein modifications on RPE lysosomal functions and POS phagocytosis. Invest
Ophthalmol Vis Sci, 48(3), 1342-1347. doi:10.1167/iovs.06-0549

Karwatowski, W. S., Jeffries, T. E., Duance, V. C., Albon, J., Bailey, A. J., & Easty, D. L. (1995). Preparation
of Bruch's membrane and analysis of the age-related changes in the structural collagens. Br J
Ophthalmol, 79(10), 944-952. doi:10.1136/bjo.79.10.944

16
Kim, J., Lee, Y. J., & Won, J. Y. (2021). Molecular Mechanisms of Retinal Pigment Epithelium Dysfunction
in Age-Related Macular Degeneration. Int J Mol Sci, 22(22). doi:10.3390/ijms222212298

Kon, V., & Jabs, K. (2004). Angiotensin in atherosclerosis. Curr Opin Nephrol Hypertens, 13(3), 291-297.
doi:10.1097/00041552-200405000-00005

Kotoula, M. G., Chatzoulis, D. Z., Tahmitzi, S., & Tsironi, E. E. (2009). Detection of macular
photocoagulation scars with confocal infrared reflection imaging. Ophthalmic Surg Lasers
Imaging, 40(4), 385-388. doi:10.3928/15428877-20096030-06

Li, M., Dolz-Marco, R., Messinger, J. D., Wang, L., Feist, R. M., Girkin, C. A., . . . Freund, K. B. (2018).
Clinicopathologic Correlation of Anti-Vascular Endothelial Growth Factor-Treated Type 3
Neovascularization in Age-Related Macular Degeneration. Ophthalmology, 125(2), 276-287.
doi:10.1016/j.ophtha.2017.08.019

Maeshima, K., Utsugi-Sutoh, N., Otani, T., & Kishi, S. (2004). Progressive enlargement of scattered
photocoagulation scars in diabetic retinopathy. Retina, 24(4), 507-511. doi:10.1097/00006982-
200408000-00002

Martin, P. (1997). Wound healing--aiming for perfect skin regeneration. Science, 276(5309), 75-81.
doi:10.1126/science.276.5309.75

Mehta, P. K., & Griendling, K. K. (2007). Angiotensin II cell signaling: physiological and pathological
effects in the cardiovascular system. Am J Physiol Cell Physiol, 292(1), C82-97.
doi:10.1152/ajpcell.00287.2006

Meng, X. M., Nikolic-Paterson, D. J., & Lan, H. Y. (2016). TGF-beta: the master regulator of fibrosis. Nat
Rev Nephrol, 12(6), 325-338. doi:10.1038/nrneph.2016.48

Neumann, J., & Brinkmann, R. (2005). Boiling nucleation on melanosomes and microbeads transiently
heated by nanosecond and microsecond laser pulses. J Biomed Opt, 10(2), 024001.
doi:10.1117/1.1896969

Nita, M., Grzybowski, A., Ascaso, F. J., & Huerva, V. (2014). Age-related macular degeneration in the
aspect of chronic low-grade inflammation (pathophysiological parainflammation). Mediators
Inflamm, 2014, 930671. doi:10.1155/2014/930671

Qin, S., Dong, N., Yang, M., Wang, J., Feng, X., & Wang, Y. (2021). Complement Inhibitors in Age-Related
Macular Degeneration: A Potential Therapeutic Option. J Immunol Res, 2021, 9945725.
doi:10.1155/2021/9945725

Querques, G., Cicinelli, M. V., Rabiolo, A., de Vitis, L., Sacconi, R., Querques, L., & Bandello, F. (2018).
Laser photocoagulation as treatment of non-exudative age-related macular degeneration: state-
of-the-art and future perspectives. Graefes Arch Clin Exp Ophthalmol, 256(1), 1-9.
doi:10.1007/s00417-017-3848-x

Rabelo, L. A., Alenina, N., & Bader, M. (2011). ACE2–angiotensin-(1–7)–Mas axis and oxidative stress in
cardiovascular disease. Hypertension Research, 34(2), 154-160. doi:10.1038/hr.2010.235

17
Ramadan, R., Dhawan, S. S., Binongo, J. N., Alkhoder, A., Jones, D. P., Oshinski, J. N., & Quyyumi, A. A.
(2016). Effect of Angiotensin II Type I Receptor Blockade with Valsartan on Carotid Artery
Atherosclerosis: A Double Blind Randomized Clinical Trial Comparing Valsartan and Placebo
(EFFERVESCENT). Am Heart J, 174, 68-79. doi:10.1016/j.ahj.2015.12.021

Rein, D. B., Wittenborn, J. S., Zhang, X., Honeycutt, A. A., Lesesne, S. B., Saaddine, J., & Vision Health
Cost-Effectiveness Study, G. (2009). Forecasting age-related macular degeneration through the
year 2050: the potential impact of new treatments. Arch Ophthalmol, 127(4), 533-540.
doi:10.1001/archophthalmol.2009.58

Ren, C., Liu, W., Yin, X., Zhang, B., & Lu, P. (2020). Renin-Angiotensin System Inhibitor Usage and Age-
Related Macular Degeneration among Hypertensive Patients: Results from the National Health
and Nutrition Examination Survey, 2005-2008. J Ophthalmol, 2020, 4252031.
doi:10.1155/2020/4252031

Richert, E., Koinzer, S., Tode, J., Schlott, K., Brinkmann, R., Hillenkamp, J., . . . Roider, J. (2018). Release of
Different Cell Mediators During Retinal Pigment Epithelium Regeneration Following Selective
Retina Therapy. Invest Ophthalmol Vis Sci, 59(3), 1323-1331. doi:10.1167/iovs.17-23163

Rodanant, N., Friberg, T. R., Cheng, L., Aurora, A., Bartsch, D., Toyoguchi, M., . . . Freeman, W. R. (2002).
Predictors of drusen reduction after subthreshold infrared (810 nm) diode laser macular grid
photocoagulation for nonexudative age-related macular degeneration. Am J Ophthalmol,
134(4), 577-585. doi:10.1016/s0002-9394(02)01691-4

Romdhoniyyah, D. F., Harding, S. P., Cheyne, C. P., & Beare, N. A. V. (2021). Metformin, A Potential Role
in Age-Related Macular Degeneration: A Systematic Review and Meta-Analysis. Ophthalmol
Ther, 10(2), 245-260. doi:10.1007/s40123-021-00344-3

Ruan, Y., Jiang, S., & Gericke, A. (2021). Age-Related Macular Degeneration: Role of Oxidative Stress and
Blood Vessels. Int J Mol Sci, 22(3). doi:10.3390/ijms22031296

Rubner, R., Li, K. V., & Canto-Soler, M. V. (2022). Progress of clinical therapies for dry age-related
macular degeneration. Int J Ophthalmol, 15(1), 157-166. doi:10.18240/ijo.2022.01.23

Salo, T., Makela, M., Kylmaniemi, M., Autio-Harmainen, H., & Larjava, H. (1994). Expression of matrix
metalloproteinase-2 and -9 during early human wound healing. Lab Invest, 70(2), 176-182.
Retrieved from https://www.ncbi.nlm.nih.gov/pubmed/8139259

Santos, R. A. S., Oudit, G. Y., Verano-Braga, T., Canta, G., Steckelings, U. M., & Bader, M. (2019). The
renin-angiotensin system: going beyond the classical paradigms. Am J Physiol Heart Circ Physiol,
316(5), H958-H970. doi:10.1152/ajpheart.00723.2018

Sharma, R., Bose, D., Maminishkis, A., & Bharti, K. (2020). Retinal Pigment Epithelium Replacement
Therapy for Age-Related Macular Degeneration: Are We There Yet? Annu Rev Pharmacol
Toxicol, 60, 553-572. doi:10.1146/annurev-pharmtox-010919-023245

Sivaprasad, S., & Chong, N. V. (2006). The complement system and age-related macular degeneration.
Eye (Lond), 20(8), 867-872. doi:10.1038/sj.eye.6702176

18
Suzuki, M., Kamei, M., Itabe, H., Yoneda, K., Bando, H., Kume, N., & Tano, Y. (2007). Oxidized
phospholipids in the macula increase with age and in eyes with age-related macular
degeneration. Mol Vis, 13, 772-778. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/17563727

Suzuki, Y., Ruiz-Ortega, M., & Egido, J. (2000). Angiotensin II: a double-edged sword in inflammation. J
Nephrol, 13 Suppl 3, S101-110. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/11132026

Takeda, H., Katagata, Y., Hozumi, Y., & Kondo, S. (2004). Effects of angiotensin II receptor signaling
during skin wound healing. Am J Pathol, 165(5), 1653-1662. doi:10.1016/S0002-9440(10)63422-
0

Van Haeringen, N. J. (1996). The renin-angiotensin system in the human eye. Br J Ophthalmol, 80(2), 99-
100. doi:10.1136/bjo.80.2.99

van Lookeren Campagne, M., LeCouter, J., Yaspan, B. L., & Ye, W. (2014). Mechanisms of age-related
macular degeneration and therapeutic opportunities. J Pathol, 232(2), 151-164.
doi:10.1002/path.4266

Vavvas, D. G., Daniels, A. B., Kapsala, Z. G., Goldfarb, J. W., Ganotakis, E., Loewenstein, J. I., . . . Miller, J.
W. (2016). Regression of Some High-risk Features of Age-related Macular Degeneration (AMD)
in Patients Receiving Intensive Statin Treatment. EBioMedicine, 5, 198-203.
doi:10.1016/j.ebiom.2016.01.033

Velez-Montoya, R., Oliver, S. C., Olson, J. L., Fine, S. L., Quiroz-Mercado, H., & Mandava, N. (2014).
Current knowledge and trends in age-related macular degeneration: genetics, epidemiology,
and prevention. Retina, 34(3), 423-441. doi:10.1097/IAE.0000000000000036

Virgili, G., Michelessi, M., Parodi, M. B., Bacherini, D., & Evans, J. R. (2015). Laser treatment of drusen to
prevent progression to advanced age-related macular degeneration. Cochrane Database Syst
Rev(10), CD006537. doi:10.1002/14651858.CD006537.pub3

Wang, Y., Zhong, Y., Zhang, L., Wu, Q., Tham, Y., Rim, T. H., . . . Liu, L. (2021). Global Incidence,
Progression, and Risk Factors of Age-Related Macular Degeneration and Projection of Disease
Statistics in 30 Years: A Modeling Study. Gerontology, 1-15. doi:10.1159/000518822

Wolter, J. R., & Falls, H. F. (1962). Bilateral confluent drusen. Arch Ophthalmol, 68, 219-226.
doi:10.1001/archopht.1962.00960030223013

Wright, C. B., & Ambati, J. (2017). Dry Age-Related Macular Degeneration Pharmacology. Handb Exp
Pharmacol, 242, 321-336. doi:10.1007/164_2016_36

Zarbin, M. A. (2004). Current concepts in the pathogenesis of age-related macular degeneration. Arch
Ophthalmol, 122(4), 598-614. doi:10.1001/archopht.122.4.598

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Chapter 2: Characterization of Selective Retinal Therapy and Wound Healing
Response in Hard Drusen Mouse Model of Age-Related Macular Degeneration
2.1 Introduction
Age-related macular degeneration (AMD) is a condition that is closely associated with drusen
formation. This has been a primary clinical biomarker of the disease progression, where ocular imaging
studies can be used to measure disease severity in humans. Additionally, AMD animal models have used
the development of drusen-like deposits between the Bruch’s membrane and the RPE layer in aged
animals (Edwards & Malek, 2007). Drusen development is commonly delineated as being either hard or
soft drusen based on the microscopic appearance or lack of a distinct border.
Genetic polymorphism studies in human studies have identified variations within the Cx3Cr1 gene
has greater risk for the development of AMD (Ma, Dang, Yang, Duan, & Zhang, 2015). The Cx3CL1/Cx3Cr1
axis has been extensively studied for neural diseases. Cx3Cr1 is highly expressed in microglia (Lee, Lee,
Song, Lee, & Chang, 2018). The high expression of Cx3Cr1 in monocytes is characteristic of non-classical
monocytes, which can induce the release of anti-inflammatory cytokines (Meghraoui-Kheddar,
Barthelemy, Boissonnas, & Combadiere, 2020). Landsman et al confirmed a 3-fold reduction in the
presence of non-classical monocytes in Cx3Cr1 -/- mice relative to wild-type mice (Landsman et al., 2009).
Accordingly, the gene knockout Cx3cr1 in mice would result in chronic activation of inflammation in retinal
tissue.
In relation to ocular disease, Jobling et al found the Cx3Cr1 pathway was instrumental in proper
retina development, where Cx3cr1 knockout (KO) led to PR dysfunction as early as 17 days after birth
(Jobling et al., 2018). One study showed microglia play a critical role in neuronal development
(Guttenplan, Blum, & Bennett, 2018). Furthermore, as expected, the microglia of Cx3Cr1-/- mice have
impaired migration due to the inability to utilize the Cx3CL1/Cx3Cr1 axis. As these impaired microglia
accumulate in the subretinal space, the mouse develops drusen-like deposits similar in characteristics as

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atherosclerotic plaques. Drusen accumulation occurs beneath the RPE as the microglia become bloated
with lipid droplets (Raoul et al., 2008). Cx3Cr1-/- can develop drusen-like deposits that is accompanied by
retinal lesions similar to those observed in humans with dAMD (Tuo et al., 2007).
Photocoagulation has previously been used to mitigate AMD progression by cauterizing sections
of the eye to prevent the spread of drusen (Figueroa, Regueras, & Bertrand, 1994). While this method
reduced drusen deposits, excessive amounts of energy commonly led to scar formation in treated eyes,
making photocoagulation a non-effective treatment method (Epstein, Aziz, Young, & Berrocal, 2013).
R:GEN is a selective retinal therapy (SRT) laser proposed to be a potential treatment for dAMD. Using a
532 nm wavelength, the laser can selectively target the melanosomes found in RPE, allowing it to
effectively avoid damage to the PR layers of the eye. Similar to photocoagulation, previous SRT studies
have also shown drusen regression with a much-improved safety profile (Guymer et al., 2019). Previous
exploration into the mechanisms of drusen regression following photocoagulation and SRT treatment
have both shown upregulation of MMPs (Jobling et al., 2015; Richert et al., 2018). Additionally, while there
is no existing literature with regards to SRT, microarray analysis of photocoagulation showed upregulation
of AT2R (Wilson et al., 2003). Because of the expansive literature highlighting the beneficial activity of
AT2R, we propose to address the gap in the knowledge and explore if and how R:GEN modulates the RAS.
It has been well established that AngII plays a critical role in the formation of atherosclerotic
plaque (Johnstone et al., 2004). Studies have shown modulation of RAS components may be an effective
strategy to prevent plaque formation. In particular, the use of ACE inhibitors (ACEis) or KO AT1R gene
(Allen & Jandeleit-Dahm, 2005; Aono et al., 2012). With respect to ocular disease, AngII/AT1R has been
proposed to contribute to retinal inflammation (Phipps et al., 2018). This biological activity has led to
coining of AngII/AT1R as components of the “Pathogenic Arm of RAS”. Accordingly, excess AngII can
sustain inflammation and potentially exacerbate ocular disease (Wilkinson-Berka, Suphapimol, Jerome,

21
Deliyanti, & Allingham, 2019). In contrast, the ability to activate the “Protective Arm of RAS” maybe able
to mitigate inflammation and pro-fibrotic healing associated with ocular disease (Abdallah et al., 2016).
In this chapter, the effect of R:GEN treatment in Cx3cr1-/- mice, a hard drusen mouse model, is
reported. Specifically, ocular images, histology of the retinal, and gene expression of MMPs were assessed
to confirm consistency with previously reported studies using SRT. The impact of R:GEN treatment on RAS
was also explored to determine if modulation of this pathway could determine a shift from pro-fibrotic or
non-fibrotic wound healing. Furthermore, the relationship between ocular and systemic RAS peptides
after R:GEN treatment was evaluated. In this context, the ability to determine a correlation will establish
an accurate and precise blood-based biomarker indicator of wound healing in an eye with hard drusen.
2.2 Materials and Methods
2.2.1 Animals
Experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of
University of Southern California (USC) that is in adherence to the National Institutes of Health (NIH)
Guidelines for the Care and Use of Laboratory Animals. Male and female Cx3Cr1 -/- mice were purchased
from Jackson Laboratories (Jackson Laboratory, Bar Harbor, ME). Mice colonies were aged to at least 12
months to emulate the humans AMD disease.
Animal Preparation and Anesthesia
Prior to R:GEN laser treatment, mice were anesthetized with a ketamine (80-100 mg/kg) and
xylazine (5-10 mg/kg) mixture given as an intraperitoneal (IP) injection. After sedation, each eye was
sterilely prepared and dilated using topical 2.5% phenylephrine HCL and 0.5% tropicamide, approximately
50 µL of each solution. In addition, approximately 50 µL of 0.5% tetracaine was also applied as a local
anesthetic. Pre-treatment images of the optic nerve and retina of each eye were acquired using the Envisu
R-Class Bioptigen System (Leica Microsystems Inc., Buffalo Grove, IL). Optical coherence tomography

22
(OCT) retinal sections were also acquired using HRA+OCT Spectralis (Heidelberg Engineering Inc., Franklin,
MA).
R:GEN Laser Application
Both eyes from each mouse were treated with the designated dosage of R:GEN laser on Day 0,
where a summary of the settings and parameters are reviewed in Table 2.1. Twenty laser (20) spots, 90µm
in diameter each, were applied around the optic nerve of each eye. The duration of each laser pulse used
was 1.7 µs, where energy auto-ramping delivered between 12.5 µJ and 17.5 µJ per lesion. The
optoacoustic (OA) and reflectometry (RFM) of the dual-dosimetry mechanism were established, such that
the RTF of each lesion would be 100% intensity.
Following laser application, mice were randomly stratified in to one of six different designated
timepoints at 0.04, 0.16, 3, 7, 30, and 90 days (N=7 per timepoint). The overall study design is summarized
in the Table 2.2.
End of Study Procedures
Prior to euthanasia, mice were anesthetized with ketamine (80-100 mg/kg) and xylazine (5-10
mg/kg) mixture given intraperitoneally (IP). Each eye was dilated and locally anaesthetized as described
above. Images of the optic nerve and retina of each eye were again acquired using the Envisu R-Class
Specifications R:GEN Laser System
Laser Wavelength (nm) 527nm
Aiming Beam 635 nm (<1mW)
Energy 30 µJ ~ 400 µJ [for mice : 5µJ ~ 100 µJ]
Pulse Duration 1.7μs
Pulse Rate ≤ 100Hz
Spot Size 200μm (for mice: ~90um)
Number of Pulses per Shot Up tp 15
Number of Spots 20
RTF sensors Optoacoustic (OA)
Reflectometry (RM)
Operation Mode Guidance Mode (AutoStop) SRT
Patterns Single
Table 2.1: R:GEN laser setting for mice retinal application

23
Bioptigen System (Leica Microsystems Inc., Buffalo Grove, IL). In addition, 0.01 mL of a 10% fluorescein
solution was administered as an IP injection and fluorescein angiography (FA) was performed to
determine the levels of leakage. OCT sections and FA were simultaneously acquired using HRA+OCT
Spectralis (Heidelberg Engineering Inc., Franklin, MA).
Upon completion of imaging, mice were immediately euthanized via isoflurane overdose, cardiac
puncture, and cervical dislocation. All animal procedures followed Institutional Animal Care and Use
Committee (IACUC) approved protocols.
2.2.2 Preparation of Blood Plasma for LC-MS/MS Analysis
A 26-gauge needle (Becton Dickinson, Franklin Lakes, IL) was heparinized (West-Ward,
Eatontown, NJ) for cardiac puncture. Approximately 1 mL of whole blood was collected and immediately
transferred to a clean 1.5 mL microcentrifuge tube containing 20 µL of protease inhibitor cocktail (PIC) as
detailed in the Table 2.3 below. The blood-filled tubes were gently inverted three times to ensure
thorough mixing of the PIC and immediately placed on ice. Blood samples were then centrifuged at 2,000
Animal
Model
R:Gen
Intensity
Time Points
(Days)
Group
Code
# of Animals Tissues
Collected
Analysis
Cx3Cr1 -/-
100% 0.04 A 7 Blood, Ocular
Fluids, Retina,
RPE
Retinal
Imaging, OCT,
FA, Flat
Mount, H&E,
RT-PCR, LC-
MS/MS
100% 0.16 B 7
100% 3 C 7
100% 7 D 7
100% 30 E 7
100% 90 F 7
NT 0 G 7
Table 2.2: Study Design for Cx3Cr1-/- hard drusen mice model

24
g at 4°C for 15 minutes. Plasma was aliquoted into a new, clean 1.5 mL microcentrifuge tube and
immediately frozen at -80°C until analysis.
2.2.3 Preparation of Eyes
Both eyes were enucleated using curved forceps (World Precision Instruments, Sarasota, FL) and
placed in separate petri dishes (VWR, Radnor, PA) containing balanced salt solution (Alcon, Geneva,
Switzerland) corresponding to each eye (OD, OS).
2.2.3.1 Preparation of Eyes for Flat Mount Staining
Three eyes from each time point were utilized for flat mount staining. For each eye, the
surrounding connective muscle and fat tissue were carefully excised using Vannas scissors (World
Precision Instruments, Sarasota, FL). The optic nerve was cut off, leaving a small stump to act as a
landmark and avoid damaging any of the posterior eye segment. After removing all excess tissue, the eye
was placed on a labeled, clean glass slide (VWR, Radnor, PA), encircled by a silicone isolator (Grace Bio-
Labs, Bend, OR). A couple drops, approximately 50 µL, of balanced salt solution (BSS) were applied to the
eye. A 31-gauge needle (Becton Dickinson, Franklin Lakes, NJ) was used to puncture the cornea, release
the aqueous humor (AQH), and deflate the eye. Four radial cuts were then made from the puncture
wound made by the 31-gauge needle: from the center of the cornea to the edge of the remaining optic
nerve. The tissue was then flattened while simultaneously rolling the lens to separate it from the retina
and discard.
Inhibitors 500X Amount of inhibitors (mg)
1,10 Phenanthroline (Sigma Aldrich, Cat# P9375 25g) 900
EDTA (Sigma Aldrich, Cat# E6758 100g) 1400
Pepstatin A (Sigma Aldrich, Cat# P4265 100 mg) 30
Enalapril maleate (Sigma Aldrich, Cat# E6888 1g) 75
Phenylmethylsulfonyl (PMSF) Sigma Aldrich Cat #78830-5G 34.8
Suspend in 4 mL of water.
Table 2.3: Protease Inhibitor Cocktail

25
Approximately 450 µL of BSS was then used to wash and lift the photoreceptors from the RPE-
sclera layer. The photoreceptors were subsequently removed and placed in a sterile 1.5 mL
microcentrifuge tube (VWR, Radnor, PA) containing 500 µL of TRIzol (Thermo Fisher, Waltham, MA). The
BSS solution, approximately 500 µL, was then collected into a labeled sterile 1.5 mL microcentrifuge tube
containing prepared PIC as summarized in the table above.
To the flattened RPE-sclera, 300 µL of 10% formalin (VWR, Radnor, PA) was applied. A glass cover
slip (VWR, Radnor, PA) was then placed on the silicone isolator and the slide was secured for staining.
2.2.3.2 Preparation of Eyes for Histopathology
Enucleated eyes were placed in their respectively labeled 10 mL scintillation vials (VWR, Radnor,
PA) containing Davidson’s fixative solution (VWR, Radnor, PA). After 72 hours of fixation, the Davidson’s
solution was replaced with ethanol until tissue was embedded in paraffin blocks for sectioning.
2.2.3.3 Preparation of Eyes for RT-PCR
For each eye, the surrounding connective muscle and fat tissue were carefully excised using
Vanass scissors. The optic nerve was cut off, leaving a small stump to act as a landmark and avoid
damaging any of the posterior eye segment. After removing all excess tissue, the eye was placed on a
labeled, clean glass slide, encircled by a silicone isolator. A couple drops, approximately 50 µL, of balanced
salt solution (BSS) were applied to the eye. A 31-gauge needle was used to puncture the cornea, release
the aqueous humor, and deflate the eye. Four radial cuts were then made from the puncture wound made
by the 31-gauge needle: from the center of the cornea to the edge of the remaining optic nerve. The tissue
was then flattened while simultaneously rolling the lens to separate it from the retina and discard.
Approximately 450 µL of BSS was then used to wash and lift the photoreceptors from the RPE-
sclera layer. The photoreceptors were subsequently removed and placed in a labeled clean 1.5 mL
microcentrifuge tube containing 500 µL of TRIzol (Invitrogen, Carlsbad, CA). The RPE-sclera was placed in
a separate, labeled clean microcentrifuge tube containing 500 µL of TRIzol. The BSS solution,

26
approximately 500 µL, was then collected into a sterile 1.5 mL microcentrifuge tube containing prepared
PIC as summarized in the table above. The retina and RPE tissues were then frozen at -80°C for storage.
2.2.4 Flat mount Staining
To stain the fixed RPE tissue for tight junctions, the following solutions were prepared as
summarized in the Table 2.4 below.
The glass coverslips were removed from the silicone isolators and the 10% formalin was aspirated.
The fixed RPE was washed three times by immersing the tissue in Hank’s Balanced Salt Solution (Corning,
Corning, NY) for 10 minutes per wash. Tissue was then permeabilized by incubating with prepared wash
buffer for 5 minutes. The wash buffer was then aspirated. The RPE was then incubated with 300μL of
prepared blocking buffer for 1 hour at room temperature in a humidified chamber (Sigma-Aldrich, St.
Louis, MO). The blocking buffer was then aspirated with a transfer pipet (VWR, Radnor, PA). The flat
mount was then incubated with 300 μL of prepared primary antibody solution for 16 hours at room
temperature in a humidified chamber. The primary antibody was then aspirated with a transfer pipet and
the tissue was subsequently washed with the prepared wash buffer 5 times, for 2 minutes per wash. The
flat mount was then incubated with the prepared secondary antibody for 1 hour. After aspirating the
secondary antibody, the flat mount was then rinsed with prepared propidium iodide solution 5 times, for

Component Manufacturer
Blocking Buffer
HBSS Corning
1.0% BSA (w/v) Calbiochem
0.1% Triton X-100 (v/v) Spectrum
Wash Buffer
HBSS Corning
0.1% Triton X-100 (v/v) Spectrum
Primary Antibody Buffer
Prepared Blocking Buffer
Rabbit Anti-ZO-1 Antibody (1:100) Invitrogen
Secondary Antibody Buffer
Prepared Blocking Buffer
Goat Anti-Rabbit Antibody (1:1000) Invitrogen
Propidium Iodide Solution
HBSS Corning
0.1% Triton X-100 (v/v) Spectrum
20 µg/mL Propidium Iodide Invitrogen
Table 2.4: Preparation of buffers for flat mount (Zo-1) staining

27
2 minutes per wash. The flat mount was then washed with the prepared wash buffer twice, for 2 min per
wash. 25 μL of Vectashield was then applied to the flat mount, and a cover slip was secured on top. The
finished slide was then cured overnight under refrigerated conditions. Images were taken on an Olympus
BX43 microscope (Olympus America Inc., PA)
2.2.5 Histopathology
The cornea and lens of each preserved eye was removed. The posterior pole of each eye was then
embedded on a paraffin block. Each eye was then cut in half through the optic nerve, with one half of the
block reserved and the remaining half sectioned. Without eliminating any segment, sections were cut
every 5-7 microns throughout the block. 4-5 sections were then mounted per slide. All slides were then
stained with H&E. Images were taken on the Aperio CS2 (Leica Biosystems, Deer Park, IL).
2.2.6 RT-PCR Analysis
2.2.6.1 Retina RNA Extraction
One scoop of stainless-steel beads was added to the retina tissue suspended in 500 µL of TRIzol.
Samples were placed in a TissueLyser II (QIAGEN, Germantown, MD) and homogenized for 2 minutes at
30 Hz. Samples were then centrifuged at 13,000g for 10 minutes at 4°C. RNA was extracted according to
the manufacturer’s protocol. RNA concentration and purity were measured using a NanoDrop
spectrophotometer (Thermo Fisher Scientific, Wilmington, DE). RNA concentration was adjusted to 10
ng/µL using nuclease-free water.
2.2.6.2 Retina RPE Extraction
One scoop of stainless-steel beads was added to the retina tissue suspended in 500 µL of TRIzol.
Samples were placed in a TissueLyser II (QIAGEN, Germantown, MD) and homogenized for 2 minutes at
30 Hz. Samples were checked for presence of pigment release after homogenization and homogenized
again if needed. Samples were then centrifuged at 13,000g for 10 minutes at 4°C. RNA was extracted
according to the manufacturer’s protocol. RNA concentration and purity were measured using a

28
NanoDrop spectrophotometer (Thermo Fisher Scientific, Wilmington, DE). RNA concentration was
adjusted to 10 ng/µL using nuclease-free water.
2.2.6.3 cDNA Synthesis
cDNA was synthesized using the extracted retina and RPE RNA using the RevertAid RT Reverse
Transcription kit (Thermo Fisher Scientific, Wilmington, DE) according to the manufacturer’s protocol.
Samples were placed in a Bio-Rad Thermal Cycler (Bio-Rad, Hercules, CA), where they were incubated
for 5 min at 25 °C followed by 60 min at 42 °C, and finally terminated by heating at 70 °C for 5 min.
2.2.6.4 cDNA Pre-amplification
The 20 µL of prepared cDNA was pre-amplified by adding 25 µL SsoAdvanced PreAmp Supermix
(Bio-Rad, Hercules, CA) and 5 µL of a prepared primer assay pool, for a final volume of 50 µL. The primer
assay pool included primer stocks of all genes that would be analyzed for RT-PCR, such that the final
preamplification reaction contains 50 nM of each primer, summarized in the table below. The cDNA
reaction mix was set for 12 cycles, for which the first stage was 3 minutes at 95°C for polymerase
activation, followed by 15 seconds for denaturation, and 4 minutes at 58°C for annealing/extension.
2.2.6.5 RT-PCR for Genes of Interest
Genes related to wound healing and extracellular matrix remodeling were probed alongside the
Renin-Angiotensin System. The RT-PCR master mix was prepared by mixing Power Up SYBR Green master-
mix (Applied Biosystems, Foster City, CA) and the forward and reverse primers for each gene. The primers
sequences used were selected from PrimerBank (Harvard, Boston, MA) and are listed in Table 2.5.

29
Prepared master mix and diluted cDNA were pipetted into a 384-well plate (VWR, Radnor, PA) by
the Integra Assist Plus (Integra Biosciences, Hudson, NH). Samples were loaded in triplicates, plates were
sealed with an adhesive plate sealer, and centrifuged at 1000 x g for 1 minute. The run method consisted
of an initial hold for 2 minutes at 50°C, followed by 95°C for 2 minutes. The amplification stage was set for
40 cycles, with a 1 second hold at 95°C and 30 seconds at 60°C for annealing/elongation. The final melt
curve stage was set for 15 seconds going from 1.6°C to 95°C, followed by 1 minute from 1.6°C to 60°C,
and finally 0.15°C to 95°C for 15 seconds. The data was collected and analyzed using Data Assist
(Invitrogen, Carlsbad, CA). Gene expression of MMP-2, MMP-3, MMP-9, TIMP-2, TGF- β1, ACE, ACE2,
AT1R, AT2R, and MasR were assessed using delta-delta CT, normalized to β-Actin as the reference gene
and compared to untreated animals. Fold changes were plotted in GraphPad Prism (GraphPad, San Diego,
CA)
2.2.7 LC-MS/MS Analysis
2.2.7.1 Solid Phase Extraction of Plasma and Ocular Fluid Samples
Frozen plasma and ocular fluid samples were thawed on ice, and from each sample, 150 µL was
aliquoted into a clean 1.5 mL microcentrifuge tube. Each sample was then spiked with 150 µL of a
prepared 40 ng/mL NorLeu
3
-Angiotensin(1-7) (NLE) internal standard solution. Samples were vortexed
thoroughly and placed on ice. While samples were incubating, Oasis MAX SPE columns (Waters, Milford,
Gene Forward (5’-3’) Reverse (5’-3’)
Mmp2 CAAGTTCCCCGGCGATGTC TTCTGGTCAAGGTCACCTGTC
Mmp3 ACATGGAGACTTTGTCCCTTTTG TTGGCTGAGTGGTAGAGTCCC
Mmp9 CTGGACAGCCAGACACTAAAG CTCGCGGCAAGTCTTCAGAG
Timp2 TCAGAGCCAAAGCAGTGAGC GCCGTGTAGATAAACTCGATGTC
AT1R AACAGCTTGGTGGTGATCGTC CATAGCGGTATAGACAGCCCA
AT2R AACTGGCACCAATGAGTCCG CCAAAAGGAGTAAGTCAGCCAAG
MasR AGAAATCCCTTCACGGTCTACA GTCACCGATAATGTCACGATTGT
ACE1 AGGTTGGGCTACTCCAGGAC GGTGAGTTGTTGTCTGGCTTC
ACE2 TCCAGACTCCGATCATCAAGC GCTCATGGTGTTCAGAATTGTGT
TGFB1 CTCCCGTGGCTTCTAGTGC GCCTTAGTTTGGACAGGATCTG
GAPDH AATGGATTTGGACGCATTGGT TTTGCACTGGTACGTGTTGAT
β-Actin GGCTCTATTCCCCTCCATCG CCAGTTGGTAACAATGCCATGT
Table 2.5: Primers for the specific gene targets

30
MA) were pre-conditioned using 1 mL of HPLC grade methanol (VWR, Radnor, PA), followed by 1 mL of
HPLC grade water (VWR, Radnor, PA).
Prepared samples were alkalinized by adding 700 µL of a 5% ammonium hydroxide solution and
vortexed thoroughly. Dissociated samples (1000 µL) were then layered onto the SPE columns. Columns
were then washed with 500 µL of 5% ammonium hydroxide, twice. Columns were then washed with 500
µL of methanol and allowed to dry for few minutes. Samples were eluted from the SPE using 500 µL
methanol with 2% formic acid, twice. Eluted samples were then dried under inert nitrogen gas. The
residue was then reconstituted using 50 µL of 10 mM Ammonium Formate buffer, pH 4.2 and transferred
to a clean HPLC vial (Sun-Sri, Rockwood, TN), where 40 µL were injected into an API4000 (AB Sciex,
Framingham, MA) for analysis.
2.2.7.2 Targeted LC-MS/MS Instrumentation and Conditions
RAS peptides were separated and quantified using an Agilent 1200 HPLC (Agilent, Santa Clara, CA)
connected to an API 4000 triple quad mass spectrometer (AB Sciex, Framingham, MA). Analyte data was
acquired using Analyst 1.6.2, where each analyte was quantified using multiple reaction monitoring
(MRM) identified during manual tuning of each compound. The MRM’s of the RAS peptides included in
the method are summarized in the Table 2.6 and 2.7 below. Acquired data was quantified using
MultiQuant 3.0 (AB Sciex, Framingham, MA).
Peptides were separated using an Agilent Poroshell 120 EC-C18 (4.6X100 mm, 2.7μm) column
(Agilent, Santa Clara, CA). Analytes were eluted using a gradient mobile phase consisting of two solvents:
1) Solvent A was HPLC grade water with 0.2% formic acid; and 2) Solvent B was HPLC grade methanol with
0.2% formic acid. The HPLC parameters and established gradient program are shown in the tables below.

31

Flow injection analysis (FIA) was performed to maximize the sensitivity of each MRM established
in the method. The optimized source temperature, gas pressures, and ion spray voltage are summarized
in Table 2.8 below.

2.2.8 Statistical Analysis
Results are expressed as mean ± SD unless specified otherwise. GraphPad Prism (GraphPad, San
Diego, CA), R (R Foundation for Statistical Computing, Vienna, Austria), and RStudio (Rstudio Inc, Boston,
MA) were used for data preparation, data cleaning, statistical analysis, and data visualization on
quantitative and qualitative data.
Ion Spray Settings Values
Collison Gas 10U*
Curtain Gas 20 U*
Ion Source Gas 1 40 U*
Ion Source Gas 2 45 U*
Ion Spray Voltage 5.5 kV
Temperature 450°C
Table 2.8: Ion Spray Settings

Parameters
Mobile Phase A: H 2O with 0.2% formic acid (FA)
Mobile Phase B: MeOH with 0.2%FA
Injection Volume: 40 µL
Run Time: 20 minutes
Flow Rate: 600µL/min
Upper Pressure Limit: 5000 psi
Autosampler Temp: 4
o
C
Table 2.6: Liquid chromatography parameters
Time (Min) Mobile Phase A (%) Mobile Phase B (%)
0 80 20
1.5 80 20
12.55 14 86
18 14 86
18.1 80 20
20 80 20
Table 2.7: HPLC Gradient Pump Program

32
2.3 Results
2.3.1 Applied Laser Settings
The total amount of energy (µJ) applied by R:GEN delivered to each eye is summarized in Figure
2.1 below. In Figure 2.1A, each point indicates the total amount of energy delivered to the right (OD) and
left (OS) eyes of each mouse (Figure 2.1A). The right eyes consistently received a total energy that ranged
between 1225 to 1253 µJ. In contrast, the energy delivered to left eyes had a wider range between 1175
and 1250 µJ. In this cohort, 69.1% of all treated eyes received 1253 µJ of energy, as seen in the distribution
of energy delivered shown in Figure 2.1B.
2.3.2 Fundus Images
Following R:GEN treatment, fundus imaging was assessed for each time interval. Representative
images are summarized in Figure 2.2. Visible laser induced lesions were seen after 0.04 and 0.16 days.
Figure 2.1: Total energy delivered to each treated eye indicate minimal difference in µJ in treated right eyes
(OD) but a wider spread in left eyes (OS) (A). Distribution of applied energies indicate most eyes were treated
with maximum energy of 1253 µJ (B).
A B

33
Lesions at these timepoint have evidence of edema. In contrast, residual lesion markings were observed
at 3, 7, and 30 days, with full lesion closure and resolution by Day 90.
2.3.3 Fluorescein Angiography
To confirm damage to the RPE, visible signs of fluorescein leakage from lesions were counted and
scored as following the Table 2.9. Dunn’s multiple comparison test was used to compare each time point
against Day 0.04 for wound closure and statistical significance.

Figure 2.2: Fundus images of Cx3Cr1 -/- hard drusen (HD) retina via Bioptigen after R:GEN treatment at 100%
intensity indicate presence of lesions until 30 days after treatment.
NT
7 Days 30 Days
0.16 Days
90 Days
3 Days 0.04 Days
Hard Drusen
100%

34
Over the course of the study, the number of visible laser lesions after the first day of treatment
were significantly reduced, indicating definitive wound closure if fluorescein was no longer able to
permeate at the site of injury. Lesion counts at 0.04 days were found to be significantly greater than
counts at 7, 30, and 90 days, but not at 0.16 or 3 days, as shown in Figure 2.4. Simultaneously, it was
observed that the severity of lesions had also decreased, as grade 3 and 4 lesions could no longer be
determined on day 3, with the majority of remaining lesions being grade 1, or the mildest case.

Figure 2.3: Representative fluorescein angiography of Cx3Cr1 -/- hard drusen (HD) retina via using HRA+OCT
Spectralis at end of study (EOS) time points indicate presence of open lesions. Clear signs of fluorescein leakage
were visible at 0.04 and 0.16 days, with minimal haze remaining by day 3, and zero leakage at 7, 30, and 90
days.
Score Criteria
1 Slight haze
2 Hazy but clear core, hazy perimeter
3 Bright core, hazy perimeter
4 Bright core, bright perimeter
Table 2.9: Lesion severity scoring and criteria for fluorescein
angiography images
NT
0.04 Days
7 Days
30 Days
0.16 Days
90 Days
3 Days

35

2.3.5 Flat Mount Staining for Tight Junctions
The RPE layer is known to normally exist in a cobble-stoned hexagonal pattern, forming tight
junctions that comprise the blood retinal barrier. Polymegathism, a variation in cell size within the
endothelial monolayer, has previously been reported to occur when RPE expand in size to replace dying
RPE (Nagai & Kalnins, 1996). To assess the extent of lesion damage to the RPE, flat mounts were fixed and
stained with anti-Zo-1 antibodies to visualize the tight junctions and examine cell morphology at the
designated time points. The wound repair response was assessed, which confirmed RPE polymegathism
in an effort to repair the laser-induced lesions, as shown in Figure 2.5 below. Cell membrane disruption
from R:GEN treatment is evident at 0.04 and 0.16 days, as a clear absence of tight junctions is visible, and

Dunn's multiple
comparisons test Mean rank diff. Summary Adjusted P Value
0.04 vs. 0.16 -6.375 ns >0.9999
0.04 vs. 3 12.35 ns 0.2417
0.04 vs. 7 21.40 ** 0.0066
0.04 vs. 30 24.19 ** 0.0014
0.04 vs. 90 24.19 *** 0.0009
Figure 2.4: Assessment and scoring of visible lesions in Cx3Cr1 -/- mouse at 100% R:GEN intensity (HD100).
Total lesion counts per eye (A), average count of lesion severity (B)

A B

36
signs of edema are present. Signs of elongation are evident by day 3, indicating the surrounding RPE are
extending to close the lesion.

2.3.5 H&E Assessment of R:GEN Lesions
To evaluate the specificity of the R:GEN laser in its ability to selectively target the RPE layer,
treated eyes were enucleated at the designated EOS time points, fixed with Davidson’s solution, and
stained with hematoxylin and eosin (H&E). Tissue sections were imaged at 40x magnification and analyzed
for lesions. Distinct breakages in the RPE layer are visible at 0.04 days, indicating disruption of the cell
membrane via R:GEN treatment. At day 7, the induced lesions are closed, and signs of previous disruption
can only be identified from occasional unevenness of the pigmented layer. Most notably, there are no
discernible changes to the inner nuclear layer (INL) nor outer nuclear layer (ONL). The unaffected
photoreceptors indicate the R:GEN laser is selective for the melanosome containing RPE while avoiding
bystander associated cellular damage as associated with traditional photocoagulation.

Figure 2.5: Zo-1 staining of isolated Cx3Cr1 -/- RPE layer treated with 100% intensity. Images are at 40x
magnification; scale bar = 20 µm. Signs of laser-induced lesions are observed at 0.04, 0.16, and 3 days.
Evidence of cell membrane elongation are obvious by day 3, indicating wound healing. Polymegathism is
observed at 7, 30, and 90 days as enlarged RPE fill in the damaged regions.
7 Day – 40X 30 Day – 40X 90 Day – 40X
3 Day – 40X 0.16 Day – 40X 0.04 Day – 40X

37

2.3.6 Fundus Autofluorescence as Measure of Drusen Regression
Because SRT was previously reported to reduce drusen deposits, drusen regression was a critical
metric for the R:GEN to meet in this study. While drusen was not detected in the H&E slides, the lipid
deposits were identifiable on Fundus autofluorescence (FAF) imaging used in pre-clinical and clinical
settings to determine the health of the retina and RPE layers. Distinct deposit borders could be easily
enumerated following R:GEN treatment. FAF signals come from lipofuscin, yellow-brown lipoprotein
deposits that accumulate in cell cytoplasm with age (Dolman & MacLeod, 1981). Commonly used in
clinic to determine drusen development and regression, FAF was also previously used to verify that a
significantly greater number of drusen-like deposits were present in the eyes of Ccl2-knockout mice,
another drusen model, as compared to age-matched healthy C57B/L mice (Luhmann et al., 2009; Ly,
Nivison-Smith, Assaad, & Kalloniatis, 2017). FAF counts showed a decreasing trend in the number of

Figure 2.6: Representative H&E images for Cx3Cr1 -/- mice treated with 100% intensity. Images are at 40x
magnification. Signs of laser-induced lesions are observed at 0.04 and 0.16 days and circled in red. Disruption
of the RPE is evident from the loss of pigment seen at 0.04 days. Pigment is filled in the induced lesion on day
7, with minimal evidence of prior disruption to the RPE layer.
CX3CR1 100% – 4 Hr – 40x CX3CR1 100% – 7 Day – 40x
CX3CR1 100% – 1 Hr – 40x
CX3CR1 – NT – 40x

38
visible deposits, especially at 30 and 90 days, although statistical significance was not found, as seen in
Figure 2.7 below.

2.3.7 RT-PCR
While the histology of laser-treated eyes indicated the R:GEN is selective for RPE, RT-PCR was
conducted to investigate and characterize the underlying mechanism of injury and wound repair in retinal
tissues. Tukey Fences for Outliers was applied to Delta CT (dCT) calculations to identify and remove
outliers prior to fold change analysis. Gene expression for MMPs were probed as an indicator of
extracellular matrix (ECM) remodeling and tissue repair. Gene expression for MMP-2 was upregulated at
0.04 and 7 days but downregulated at 30 days. MMP-3 was upregulated at 0.04, 3, and 90 days, while
MMP-9 was downregulated across all time points after laser treatment. Furthermore, there was a
significant upregulation of TIMP-2, the natural inhibitor of MMP-2, at 0.04, 7, and 90 Days of the study,
as shown in Figure 2.8.

Figure 2.7: Observed changes in fundus autofluorescence counts over time indicate a decreasing trend in sub-
retinal lipid deposits. Changes were not statistically significant due to high variability across animals.
Dunnett's multiple comparisons test Mean rank diff. Summary Adjusted P Value
NT vs. 0.04 Day 9.357 ns >0.9999
NT vs. 0.16 Day 2.857 ns >0.9999
NT vs. 3 Day 9.024 ns 0.9259
NT vs. 7 Day 2.057 ns >0.9999
NT vs. 30 Day 12.36 ns 0.3066
NT vs. 90 Day 11.02 ns 0.4910

39
Components of RAS (e.g., ACE1, ACE2, AT1R, AT2R, and MasR) were also probed to determine the
impact of R:GEN on RAS found in the RPE. ACE1 was found to be significantly downregulated at all time
points except on day 90, where gene expression was significantly upregulated. ACE2 was found to only be
significantly upregulated at 0.4 and 90 days, but significantly downregulated at 0.16, 3, and 30 days. AT1R
was found to be significantly upregulated at 7 and 90 days, but significantly downregulated at 0.04, 0.16,
3, and 30 days. Similarly, AT2R gene expression was also found to be significantly upregulated at 7 and 90
days, but significantly downregulated at 0.04, 0.16, 3, and 30 days. Findings are summarized in Figure 2.9.
Gene expression of MasR was significantly upregulated at 0.04, 7, and 90 days, but significantly
downregulated at 0.16, 3, and 30 days. TGF-β1 gene expression was also found to be significantly

Figure 2.8: Observed changes in laser-treated RPE gene expression of metalloproteinases relative to untreated.
MMP-2 (A), MMP-3 (B), MMP-9 (C), TIMP-2 (D)
A B
C D

40
upregulated at 0.04, 7, and 90 days, but significantly downregulated at day 3. Findings are summarized in
Figure 2.10.

Figure 2.10: Observed changes in laser-treated RPE gene expression of TGF- β1 and MasR relative to untreated.
TGF- β1 (A), MasR (B)
A B

Figure 2.9: Observed changes in laser-treated RPE gene expression of RAS components relative to untreated.
ACE1 (A), ACE2 (B), AT1R (C), AT2R (D)
A B
C D

41

2.3.8 LC-MS/MS
Renin Angiotensin Peptide Systemic Metabolism
Collected plasma was analyzed for systemic changes in RAS peptides over the time after laser
treatment. Systemic levels of Ang(1-10) or AngI, an inactive peptide precursor, were observed to stay low
during the first day of laser induction. Increased concentration of Ang(1-10) was seen starting on Days 3,
but dropped down to basal levels on Day 7. Interestingly, on day 90, systemic Ang(1-10) levels were
elevated. Levels of Ang(1-10) were not statistically significant for any time points when compared to
untreated eyes, as seen in Figure 2.11.
The byproduct metabolites of Ang(1-10) include AngII, Ang(1-9) and Ang(1-7); peptides known to
be active in RAS through their binding to their respective receptors. Systemic levels of Ang(1-9) were
elevated at 0.16 days after R:GEN treatment, where the changes were not statistically significant due to
the high variability. However on 3 and 90 days, the systemic levels of Ang(1-9) dropped significantly, as
shown in Figure 2.12 below.

Figure 2.11: Plasma levels of Ang(1-10) after R:GEN treatment over a 90 day time course

Dunnett's multiple comparisons test Mean rank diff. Summary Adjusted P Value
NT vs. 0.04 Day 8.667 ns >0.9999
NT vs. 0.16 Day 6.381 ns >0.9999
NT vs. 3 Day -10.62 ns >0.9999
NT vs. 7 Day 0.5000 ns >0.9999
NT vs. 30 Day -0.8333 ns >0.9999
NT vs. 90 Day -9.262 ns >0.9999

42
Systemic levels of AngII were found to be lowered across all time points after R:GEN treatment.
However, the reduced levels of AngII were only found to be statistically on days 3 and 30 when compared
to untreated animals, as indicated in Figure 2.13.
Interestingly, systemic Ang(1-7) also dropped at 0.04, 0.16, 3, 7, and 90 days after laser induction,
whereas Ang(1-7) was increased on day 30. These differences were only found to be statistically significant
on day 90, as seen in Figure 2.14.

Figure 2.12: Plasma levels of Ang(1-9) after R:GEN treatment over a 90 day time course

Figure 2.13: Plasma levels of AngII after R:GEN treatment over a 90 day time course

Dunnett's multiple comparisons test Mean rank diff. Summary Adjusted P Value
NT vs. 0.04 Day 2.143 ns >0.9999
NT vs. 0.16 Day 1.857 ns >0.9999
NT vs. 3 Day 24.07 * 0.0396
NT vs. 7 Day 14.33 ns 0.6866
NT vs. 30 Day 17.71 ns 0.2736
NT vs. 90 Day 26.79 * 0.0150
Dunnett's multiple comparisons test Mean rank diff. Summary Adjusted P Value
NT vs. 0.04 Day 13.19 ns 0.8066
NT vs. 0.16 Day 14.48 ns 0.6027
NT vs. 3 Day 32.83 ** 0.0012
NT vs. 7 Day 6.667 ns >0.9999
NT vs. 30 Day 17.69 ns 0.2682
NT vs. 90 Day 28.19 ** 0.0083

43
The levels of metabolites downstream of AngII and Ang(1-7) were also evaluated. AngII
metabolite includes Ang(2-8) and Ang(3-8), where the N-terminal cleavage of amino acid(s) will form these
metabolites. Ang(1-7) can be metabolized to Ang(1-5), which is also a ligand for Mas. Systemic Ang(2-8)
levels after R:GEN treatment were found to be downregulated relative to untreated animals, with
statistical significance found at day 90, as shown in Figure 2.15.
Similar to Ang(2-8), systemic levels of Ang(3-8) were also found to be decreased at all time points
after R:GEN treatment, but no time points were found to be statistically significant, likely due to high
variability, as seen in Figure 2.16.
For Ang(3-7), systemic levels of this peptide were decreased at 0.04, 0.16, 3, 30, and 90 days, whiles levels
were found to be increased at day 7. However, no statistical significance was found, as seen in Figure 2.17.

Figure 2.14: Plasma levels of Ang(1-7) after R:GEN treatment over a 90 day time course

Dunnett's multiple comparisons test Mean rank diff. Summary Adjusted P Value
NT vs. 0.04 Day 15.48 ns 0.4849
NT vs. 0.16 Day 9.190 ns >0.9999
NT vs. 3 Day 22.19 ns 0.0738
NT vs. 7 Day 4.500 ns >0.9999
NT vs. 30 Day 14.48 ns 0.6146
NT vs. 90 Day 28.05 ** 0.0093

Figure 2.15: Plasma levels of Ang(2-8) after R:GEN treatment over a 90 day time course

Dunnett's multiple comparisons test Mean rank diff. Summary Adjusted P Value
NT vs. 0.04 Day 17.00 ns 0.3290
NT vs. 0.16 Day 11.14 ns >0.9999
NT vs. 3 Day 22.07 ns 0.0760
NT vs. 7 Day 5.000 ns >0.9999
NT vs. 30 Day 17.00 ns 0.3290
NT vs. 90 Day 32.21 ** 0.0016

44

In contrast, systemic Ang(1-5) was found to initially decrease at 0.04 days, but then elevated at
all other time points, as seen in Figure 2.18. No statistical significance was found for systemic Ang(1-5)
between treated and untreated mice.

Renin Angiotensin Peptide Ocular Metabolism

Figure 2.18: Plasma levels of Ang(1-10) after R:GEN treatment over a 90 day time course

Dunnett's multiple comparisons test Mean rank diff. Summary Adjusted P Value
NT vs. 0.04 Day 5.810 ns >0.9999
NT vs. 0.16 Day 6.381 ns >0.9999
NT vs. 3 Day 1.238 ns >0.9999
NT vs. 7 Day 0.3333 ns >0.9999
NT vs. 30 Day -0.9048 ns >0.9999
NT vs. 90 Day -5.476 ns >0.9999

Figure 2.16: Plasma levels of Ang(3-8) after R:GEN treatment over a 90 day time course

Figure 2.17: Plasma levels of Ang(1-10) after R:GEN treatment over a 90 day time course

Dunnett's multiple comparisons test Mean rank diff. Summary Adjusted P Value
NT vs. 0.04 Day 6.857 ns >0.9999
NT vs. 0.16 Day 10.57 ns >0.9999
NT vs. 3 Day 18.79 ns 0.2043
NT vs. 7 Day -0.6667 ns >0.9999
NT vs. 30 Day 9.357 ns >0.9999
NT vs. 90 Day 21.00 ns 0.1069
Dunnett's multiple comparisons test Mean rank diff. Summary Adjusted P Value
NT vs. 0.04 Day 19.69 ns 0.1826
NT vs. 0.16 Day 14.75 ns 0.6294
NT vs. 3 Day -2.625 ns >0.9999
NT vs. 7 Day 3.357 ns >0.9999
NT vs. 30 Day -4.875 ns >0.9999
NT vs. 90 Day 2.000 ns >0.9999

45
Collected ocular fluids were analyzed for RAS peptides levels over the time after laser treatment.
To evaluate the role that angiotensin peptides may have in the ocular healing that coincides with the
observed histological findings. A nonlinear fit was constructed using the concentrations measured from
the ocular samples, and fluctuations within the RAS peptide metabolic pathway were examined.
After R:GEN treatment, the ocular Ang(1-10) or AngI levels were projected to increase up until
day 30 and then declined through day 90. Gene expression of ACE1 was also compared against Ang(1-10),
as Ang(1-10) is a substrate for ACE1, which subsequently gets metabolized to AngII. Comparing ocular
levels of AngII against ocular Ang(1-10) and ACE1, AngII was found to increase slightly but remain relatively
flat across all time points after R:GEN treatment, as seen in Figure 2.19. To determine whether the lack of
AngII formation in the eye could be explained with gene expression of ACE1, we have confirmed
downregulation of ACE1 gene expression in RPE, suggesting the observations in AngII are correct.

Figure 2.19: Ocular Metabolism of Ang(1-10) to AngII in relations to ACE1 gene expression
A(1-10)
Ang I
A(1-9)
A(1-7)
A(1-8)
AngII
ACE2
ACE
ACE2
AT
1
AT
2 Mas
ACE

46
Although Ang(1-10) is known to primarily get metabolized by ACE1, it has also known to be a
substrate for ACE2, converting Ang(1-10) to Ang(1-9). To determine if R:GEN treatment will have an effect
how Ang(1-10) gets metabolized, ocular levels of Ang(1-9) in relations to Ang(1-10) were also evaluated,
as seen in Figure 2.20. Ocular Ang(1-9) steadily decreased over the course of the study. Meanwhile, the
fluctuations in ACE2 gene expression within the RPE did not coincide with the changes observed in Ang(1-
10) nor Ang(1-9).

Figure 2.20: Ocular Metabolism of Ang(1-10) to Ang(1-9) in relations to ACE2 gene expression
A(1-10)
Ang I
A(1-9)
A(1-7)
A(1-8)
AngII
ACE
ACE2
ACE
AT
1
AT
2 Mas
ACE2

47
Since ACE2 also catalyzes the conversion of AngII to Ang(1-7), we evaluated changes of ocular
Ang(1-7) in relations to these two parameters, as shown in Figure 2.21. As mentioned above, ACE2 gene
expression was immediately upregulated after R:GEN treatment, but subsequently downregulated until
day 90. When ocular levels of Ang(1-7) were evaluated, peptide levels consistently decreased over the
study.

Another source for Ang(1-7) is the metabolism of Ang(1-9) to Ang(1-7) via ACE1 metabolism, as shown in
Figure 2.22. As previously mentioned, ocular levels of Ang(1-9) consistently decreased across the study

Figure 2.21: Ocular Metabolism of AngII to Ang(1-7) in relations to ACE2 gene expression
A(1-10)
Ang I
A(1-9)
A(1-7)
A(1-8)
AngII
ACE
ACE2
ACE
AT 1
AT
2 Mas
ACE2

48
with gene expression of ACE1 remaining significantly downregulated until day 90. Similarly, Ang(1-7)
decreased as well over the same period, although at a rate that is more drastic than the rate of Ang(1-9).
In addition to RAS peptide metabolism, the potential binding of each active peptide to its
respective receptor was explored in relationship to the peptide fluctuations observed in the ocular fluids
of treated animals. Gene expression of both AT1R and AT2R were immediately downregulated at 0.04,
0.16, 3, and 30 days after laser treatment, as seen in Figure 2.23. When examining the relationship
between ocular AngII levels with AT1R and AT2R expression, the findings would suggest that the general
increase in ocular levels of AngII are related to the decrease in both of its receptor subtypes.

Figure 2.22: Ocular Metabolism of Ang(1-9) to Ang(1-7) in relations to ACE1 gene expression
A(1-10)
Ang I
A(1-9)
A(1-7)
A(1-8)
AngII
ACE2
ACE
ACE2
AT
1
AT
2 Mas
ACE

49
Ang(1-7) is the endogenous ligand for MasR, for which the binding to this receptor would also
activate the pro-regenerative arm of RAS. While gene expression of MasR was significantly upregulated
at 0.04, 7, and 90 days, expression was significantly downregulated at 0.016 and 3, and 30 days. When
comparing the relationship between Ang(1-7) and MasR gene expression, it appears that rate of decrease
in ocular Ang(1-7) accelerates when MasR gene expression is significantly upregulated, as seen in Figure

Figure 2.23: Ocular Binding of AngII in relations to gene expression of AT1R and AT2R

Figure 2.24: Ocular Binding of Ang(1-7) in relation to MasR gene expression
A(1-10)
Ang I
A(1-9)
A(1-7)
A(1-8)
AngII
ACE
ACE2
ACE
ACE2
AT 1
AT 2 Mas
A(1-10)
Ang I
A(1-9)
A(1-7)
A(1-8)
AngII
ACE
ACE2
ACE
ACE2
AT
1
AT
2 Mas

50
2.24. Given the binding of Ang(1-7) to MasR initiates non-fibrotic wound healing, the data would suggest
non-fibrotic wound healing spikes at 0.04, 7, and 90 day time points.
Ang(1-9) was recently shown to be a substrate that bind onto AT2R, and gene expression for the
receptor was also compared in relations to its ocular concentrations, as shown in Figure 2.25. AT2R gene
expression was significantly downregulated early at 0.04, 0.16, and 3 days, with significant increases at 7
and 90 days. While ocular Ang(1-9) was shown to decrease steadily over the course of the study, these
levels do not appear to be as affected by upregulation of AT2R.
Correlations and Statistical Analyses of Gene Expression and RAS Metabolomics
The quantified RAS concentrations were then converted to fold change relative to untreated
control mice such that the observed changes in plasma and ocular fluid could be correlated as well as
compared the RT-PCR results, as shown in Figure 2.26. A two-tailed Spearman correlation analysis was

Figure 2.25: Ocular Binding of Ang(1-9) in relations to AT2R gene expression
A(1-10)
Ang I
A(1-9)
A(1-7)
A(1-8)
AngII
ACE
ACE2
ACE
ACE2
AT
1
AT
2 Mas

51
conducted with a 95% confidence level between the plasma and ocular levels to determine whether the
changes within the ocular space can be tracked by the levels in the eye.
It was found that a negative correlation of ocular and plasma fold changes of Ang(1-10) and Ang
(1-5) were statistically significant in Cx3Cr1 -/- mice, r = -0.7857 (p = 0.0480) and r = -0.8214 (p = 0.0341),
respectively. The full analysis of correlations is shown below in Table 2.12 below.

Figure 2.26 Aggregate fold change in RAS peptides of 100% intensity treatment between plasma and ocular
matrices for correlation analysis.
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.001
0.01
0.1
1
10
Time (Days)
Fold Change from Untreated
Plasma_Ocular_AngII
Ocular
Plasma
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.1
1
10
Plasma_Ocular_Ang(2-8)
Time (Days)
Fold Change from Untreated
Ocular
Plasma
3
7
7
7
6
7
7
5
10
11
11
9
10
9
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.1
1
10
Time (Days)
Fold Change from Untreated
Plasma_Ocular_Ang(1-10)
Plasma
Ocular
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.1
1
10
100
Plasma_Ocular_Ang(1-7)
Time (Days)
Fold Change from Untreated
Ocular
Plasma
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.1
1
10
Plasma_Ocular_Ang(3-8)
Time (Days)
Fold Change from Untreated
Ocular
Plasma
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.1
1
10
Plasma_Ocular_Ang(1-9)
Time (Days)
Fold Change from Untreated
Ocular
Plasma
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.01
0.1
1
10
100
1000
Plasma_Ocular_Ang(1-5)
Time (Days)
Fold Change from Untreated
Ocular
Plasma
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.01
0.1
1
10
100
Plasma_Ocular_Ang(3-7)
Time (Days)
Fold Change from Untreated
Ocular
Plasma
Correlation of Plasma and Ocular Fluid
Peptide A(1-10) A(1-9) Ang II A(1-7) A(1-5) A(2-8) A(3-8) A(3-7)
Spearman r
-0.786 0.357 -0.321 0.214 -0.8214 -0.5 -0.393 -0.703
P (two-tailed)
0.048 0.444 0.498 0.662 0.034 0.267 0.3956 0.0897
Number of XY Pairs 7 7 7 7 7 7 7 7
Table 2.12: Correlation analysis between plasma and ocular RAS peptides

52
Given the observed sustained increases in the relative gene expression of AT2R and MasR in the
RPE, it would be expected that a shift towards the regenerative arm occurred with R:GEN treatment using
both 50% and 100% intensities. The correlation of gene expression of the RAS enzymes, receptors, and
ocular peptide levels are summarized in Figure 2.27 and Figure 2.28. It appears that the peptide ratios,
which would be indicative of enzymatic cleavage, did not have a positive correlation with gene expression
of ACE1 and ACE2 in RPE. However, the ratio of AngII/AngI and gene expression of ACE1 were found to be
negatively correlated and statistically significant (p = 0.03).

Correlation of Ocular Fluid & RPE Enzyme
Enzyme RPE - ACE1 RPE - ACE2
Peptide A(1-8)/A(1-10) A(1-7)/A(1-9) A(1-9)/A(1-10) A(1-7)/A(1-8)
Spearman r -0.07143 -0.6429 -0.5714 -0.3571
P (two-tailed) 0.9063 0.1389 0.2000 0.4444
Number of XY
Pairs 7 7 7 7

Figure 2.27: Correlation between Ocular RAS peptide ratios and gene expression their respective enzymes in
RPE

53
None of the correlations between ocular RAS peptides and their respective receptors were found
to be statistically significant, as shown in Figure 2.28. However, it appears that ocular peptide levels are
negatively correlated with the expression of their respective receptors.

Correlation of Ocular Fluid & RPE Receptor
RPE - AT1R RPE - AT2R RPE - MasR
AngII AngII A(1-9) A(1-7)
Spearman r -0.5357 -0.3929 -0.4286 -0.3243
P (two-tailed) 0.2357 0.3956 0.3536 0.4746
Number of XY Pairs 7 7 7 7
Figure 2.28: Correlation between Ocular RAS peptides and gene expression their respective receptors in RPE

54
Correlation and Statistical Analyses of Fundus Autofluorescence and RAS Components
To probe the hypothesis that activation of the RAS pathway could lead to drusen regression, a
two-tailed Spearman correlation analysis was conducted between the FAF counts and RAS components.
Desirable negative relationships were interpreted as molecular signals that would increase as drusen
counts decrease. While the signals in this analysis were all identified as weakly negative correlations, they
include MasR, Ang(1-10), Ang II, Ang(1-7), the ratios Ang II/Ang(1-10), and Ang(1-9)/Ang(1-10). The results
are shown in Table 2.11 below, ranked in order of strongest to weakest correlation.
2.4 Discussion
In this chapter, we explored the safety and efficacy of the Lutronic R:GEN laser in a hard drusen
mouse model of AMD with a time course design. Images, histology, gene expression, and metabolomics
techniques were utilized to provide insight into this potential therapy. While previously the safety of
R:GEN in healthy C57B/L mice, we have not yet explored the safety and efficacy of SRT in animal models
of AMD. Furthermore, there have been no prior studies of SRT and its effect on ocular RAS, an
underexplored pathway in AMD which overlaps with multiple pathways known to be dysregulated in the
disease. Characterizing the R:GEN-induced changes in RAS may provide new insights into treating dry
AMD, a disease for which no FDA approved treatments exist.
In this hard drusen mouse study, through a combination of RT-PCR, histology, and fundus
imaging, we showed that R:GEN is safe and capable of reducing drusen deposits. H&E cross-sections of
collected eyes indicated that laser-induced lesions to the RPE had healed by day 7. Furthermore, there
Correlation of FAF Counts and RAS Components
Relationship r P (two-tailed) P value Summary
Counts vs. Ang II -0.3571 0.4444 ns
Counts vs. Ang(1-10) -0.2143 0.6615 ns
Counts vs. MasR -0.1429 0.7825 ns
Counts vs. AngII/Ang(1-10) -0.1429 0.7825 ns
Counts vs. Ang(1-9)/Ang(1-10) -0.07143 0.9063 ns
Counts vs. Ang(1-7) -0.03571 0.9635 ns
Table 2.11: Correlation analysis between FAF drusen counts and ocular RAS components

55
was no indication of damage to the photoreceptor layer in the H&E cross-sections, a known side effect
of previous photocoagulation studies, proving R:GEN is significantly safer than conventional
photocoagulation. We showed that hard drusen deposits, as measured by Fundus Autofluorescence
(FAF), had trended lower over time after R:GEN treatment, although statistical significance was not met.
It was previously reported that selective retinal therapy (SRT) could potentially be a treatment
for dAMD. While previous work in our lab tested the safety within healthy, normal C57/Bl mice, a hard
drusen mouse model (Cx3Cr1 -/-), would provide more insights into the efficacy of the R:GEN laser and
be more translatable to a clinical setting. Drusen deposits are known to be a clinical marker of AMD
progression, although the presence of hard drusen is a lower risk factor when compared to soft drusen
(Klein et al., 2015). However, because hard drusen has previously been reported to merge into larger
soft drusen deposits via the “oil spill” phenomenon, it could be valuable to treat hard drusen early to
prevent further disease progression (Curcio, 2018). Therefore, to explore this option, it is critical to
explore the safety and efficacy of the R:GEN laser in an animal model that develops hard drusen.
2.4.1 R:GEN upregulates ECM turnover in hard drusen (Cx3Cr1 -/-) mouse model
Previously reported findings proposed the drusen regression observed in SRT-treated AMD
patients is a result of the extracellular matrix remodeling that occurs during the tissue healing response
(Jobling et al., 2015). We confirmed that R:GEN was capable of upregulating metalloproteinase
expression in our Cx3Cr1 -/- mouse model, with gene expression for MMP-2 being upregulated at 0.04
and 7 days; MMP-3 being upregulated at 0.04, 3, and 90 days. However, MMP-9 was downregulated,
suggesting that wound repair in the Cx3Cr1 -/- model prioritizes wound contraction and not cell
migration (Caley, Martins, & O'Toole, 2015).
2.4.2 R:GEN shifts ocular RAS towards pro-regeneration in hard drusen mouse model
To further explore the molecular mechanism of wound healing after SRT, components of the
Renin Angiotensin System (RAS) were also measured, as the modulation of this system is known to

56
promote either fibrotic or non-fibrotic wound healing. To date, no other studies have explored how SRT
modulates RAS in the eye of animals with hard drusen, nor have they been proposed as biomarkers
capable of predicting safety and efficacy with treatment.
While the RAS pathway is traditionally known to regulate hemostasis, it has more recently been
shown to play a vital role in tissue repair (K. Rodgers et al., 1997; K. E. Rodgers, Espinoza, et al., 2003).
Prolonged elevation of Angiotensin II (AngII) is known to activate the pathogenic arm of RAS through its
binding to AT1R, which subsequently activates TGF- β, the master regulator of fibrosis (Daniel, 2008; Meng
et al., 2016). Meanwhile the regenerative arm of RAS counterbalances the pathogenic arm when AngII
binds to AT2R or when Ang(1-7) binds to Mas receptor(K. E. Rodgers, Roda, et al., 2003).
To characterize how SRT would modulate RAS and whether plasma levels of angiotensin peptides
could be representative of ocular activity, a spearman correlation was conducted between plasma levels
and ocular levels of each peptide. It was observed that the relationship between plasma and ocular levels
of Ang(1-10) and Ang(1-5) were identified as statistically significant in Cx3Cr1 -/- mice. Surprisingly, the
relationship between plasma and ocular levels were found to have negative correlations.
From this data, it could be surmised that the decrease in plasma Ang(1-10) immediately after
R:GEN treatment (0.04 and 0.16 days) may be indicative of successfully creating lesions in the RPE.
Furthermore, the observed increase in ocular Ang(1-10) would provide more substrate for the angiotensin
converting enzymes (ACE1 and ACE2) to generate the active downstream metabolites. Similarly, the
plasma/ocular relationship for the active peptide AngII was also found to be negatively correlated,
although not statistically significant. These findings highlight the possibility that decreased levels of these
two peptides in plasma may suggest transport across into the ocular microenvironment.
Because downstream peptides of RAS were found to be upregulated at various time points in this
study, the effect that R:GEN had on the expression and activity of RAS enzymes were explored. The ratios
of RAS peptides, calculated as product/substrate, suggested negligible conversion of AngII from Ang(1-

57
10), but significant conversion of Ang(1-9) to Ang(1-7) although ACE1 expression in RPE was actually found
to be downregulated until day 90. Similarly, our metabolomic approach indicated the conversion of Ang(1-
10) to Ang(1-9) was negligible, while conversion of AngII to Ang(1-7) was elevated, even though ACE2
expression in RPE was again downregulated. As such, the correlation analysis between gene expression
and peptide metabolomics were all found to have negative relationships, although none were found to
be statistically significant. Unexpectedly, the gene expression which runs counter to what was determined
in the metabolomics approach, as well as what is known regarding peptide metabolism within the
pathway. The conflicting results from suggest additional gaps that may be explored in future iterations of
this study. For example, one possible explanation for this discrepancy might be the release of soluble ACE1
and ACE2 into the vitreous humor after laser induction, which might continue to convert RAS peptides
even though gene expression was found to be downregulated. However, due to the limitations regarding
volume of a mouse eye and sample size, the presence and quantitation of soluble enzymes could not be
performed.
While the metabolomic analysis of RAS peptides suggested a shift towards the production of
pro-regenerative peptides, activation of the pro-regenerative arm of RAS is dependent on the peptides
binding to their respective receptors. Correlation analysis was again conducted to determine if any
relationships between the angiotensin peptides and their respective receptors exist, for which changes
in peptide levels were correlated with changes in gene expression of their respective receptors. While
no correlations were found to be statistically significant, it was observed that expression of RAS
receptors AT1R and AT2R were downregulated until day 7, or the final day of the acute healing phase as
determined by histology. The downregulation in RPE AT1R and AT2R highlight a stark difference in the
RPE of this Cx3Cr1 -/- model compared to the gene expression of C57B/L RPE shown in previous studies
(not shown). However, MasR expression in Cx3Cr1 knockout RPE was significantly upregulated at 0.04

58
days, matching the early increase in ocular Ang(1-7) levels, although the subsequent divergence
between ocular Ang(1-7) and RPE MasR levels match the gene expression pattern of AT1R and AT2R.
Furthermore, the correlation analysis conducted between the FAF results and the RAS
components probed within this study indicate a weakly negative relationship between counted deposits
and various peptides, peptide ratios, and receptor gene expression. These results provide additional
supporting evidence that RAS activation may be valuable in promoting drusen regression and slowing
the progression of AMD. Future efforts in probing and building on these correlations should expand the
number of samples to increase statistical power, as well as aim to sample match FAF counts to RAS
measures to validate these findings.
Advantages of this hard drusen mouse model includes the development of drusen-like deposits
and other AMD-like characteristics, easy colony expansion, and relatively short disease pathogenesis
compared to non-rodent models. However, the size of a mouse eye created physical challenges
regarding studying ocular diseases. Surgical precision was significantly more challenging when compared
to the dissection of non-rodent eyes. Furthermore, harvesting photoreceptors and RPE for molecular
studies generated low yield per animal, limiting the types and number of analyses that could be
performed. Typical mouse eyes yield an estimated 50,000 cells, which should be taken into
consideration for the design of future studies (Fernandez-Godino, Garland, & Pierce, 2016). While the
results of this study support the claims that R:GEN, a proprietary SRT laser, is safe and has evidence of
drusen regression, further study should be conducted in a model of soft drusen, which would be more
representative of AMD risk.

2.5 Conclusion
In this chapter, it was reported that R:GEN was well-tolerated in Cx3Cr1 -/- mice and
upregulated mechanisms related to extra cellular matrix remodeling and wound healing post-treatment.

59
Analyses of all collected images for each time point and laser intensity show that eyes with hard drusen
are capable of recovering from treatment. R:GEN treatment was also shown to modulate ocular RAS,
which along with the observed drusen regression, suggest that potential therapeutic targets may also
exist within this pathway. In addition, the negative correlation between plasma and ocular levels of
Ang(1-10) and the negative correlation between ocular Ang(1-10) with FAF counts suggest systemic
decrease in Ang(1-10) after R:GEN treatment might be indicative of laser efficacy.

60
2.6 Chapter 2 References
Abdallah, W. F., Louie, S. G., Zhang, Y., Rodgers, K. E., Sivok, E., G, S. d., & Humayun, M. S. (2016).
NorLeu3A(1-7) Accelerates Clear Corneal Full Thickness Wound Healing. Invest Ophthalmol Vis
Sci, 57(4), 2187-2194. doi:10.1167/iovs.15-18515

Allen, T. J., & Jandeleit-Dahm, K. A. (2005). Preventing atherosclerosis with angiotensin-converting
enzyme inhibitors: emphasis on diabetic atherosclerosis. Curr Drug Targets Cardiovasc Haematol
Disord, 5(6), 503-512. doi:10.2174/156800605774962068

Aono, J., Suzuki, J., Iwai, M., Horiuchi, M., Nagai, T., Nishimura, K., . . . Higaki, J. (2012). Deletion of the
angiotensin II type 1a receptor prevents atherosclerotic plaque rupture in apolipoprotein E-/-
mice. Arterioscler Thromb Vasc Biol, 32(6), 1453-1459. doi:10.1161/ATVBAHA.112.249516

Caley, M. P., Martins, V. L., & O'Toole, E. A. (2015). Metalloproteinases and Wound Healing. Adv Wound
Care (New Rochelle), 4(4), 225-234. doi:10.1089/wound.2014.0581

Curcio, C. A. (2018). Soft Drusen in Age-Related Macular Degeneration: Biology and Targeting Via the Oil
Spill Strategies. Invest Ophthalmol Vis Sci, 59(4), AMD160-AMD181. doi:10.1167/iovs.18-24882

Daniel, C. (2008). Blocking of angiotensin II is more than blocking of transforming growth factor-beta.
Kidney Int, 74(5), 551-553. doi:10.1038/ki.2008.290

Dolman, C. L., & MacLeod, P. M. (1981). Lipofuscin and its Relation to Aging. In S. Fedoroff & L. Hertz
(Eds.), Advances in Cellular Neurobiology (Vol. 2, pp. 205-247): Elsevier.

Edwards, A. O., & Malek, G. (2007). Molecular genetics of AMD and current animal models.
Angiogenesis, 10(2), 119-132. doi:10.1007/s10456-007-9064-2

Epstein, I. J., Aziz, H. A., Young, R. C., & Berrocal, A. M. (2013). Macular hard exudates and scar
formation after laser photocoagulation in retinopathy of prematurity. J Pediatr Ophthalmol
Strabismus, 50 Online, e30-32. doi:10.3928/01913913-20130625-01

Fernandez-Godino, R., Garland, D. L., & Pierce, E. A. (2016). Isolation, culture and characterization of
primary mouse RPE cells. Nat Protoc, 11(7), 1206-1218. doi:10.1038/nprot.2016.065

Figueroa, M. S., Regueras, A., & Bertrand, J. (1994). Laser photocoagulation to treat macular soft drusen
in age-related macular degeneration. Retina, 14(5), 391-396. doi:10.1097/00006982-
199414050-00001

Guttenplan, K., Blum, J., & Bennett, M. (2018). A Role for Microglia in Retinal Development. J Neurosci,
38(43), 9126-9128. doi:10.1523/JNEUROSCI.1813-18.2018

Guymer, R. H., Wu, Z., Hodgson, L. A. B., Caruso, E., Brassington, K. H., Tindill, N., . . . Laser Intervention
in Early Stages of Age-Related Macular Degeneration Study, G. (2019). Subthreshold
Nanosecond Laser Intervention in Age-Related Macular Degeneration: The LEAD Randomized
Controlled Clinical Trial. Ophthalmology, 126(6), 829-838. doi:10.1016/j.ophtha.2018.09.015

61
Jobling, A. I., Guymer, R. H., Vessey, K. A., Greferath, U., Mills, S. A., Brassington, K. H., . . . Fletcher, E. L.
(2015). Nanosecond laser therapy reverses pathologic and molecular changes in age-related
macular degeneration without retinal damage. FASEB J, 29(2), 696-710. doi:10.1096/fj.14-
262444

Jobling, A. I., Waugh, M., Vessey, K. A., Phipps, J. A., Trogrlic, L., Greferath, U., . . . Fletcher, E. L. (2018).
The Role of the Microglial Cx3cr1 Pathway in the Postnatal Maturation of Retinal
Photoreceptors. J Neurosci, 38(20), 4708-4723. doi:10.1523/JNEUROSCI.2368-17.2018

Johnstone, M. T., Perez, A. S., Nasser, I., Stewart, R., Vaidya, A., Al Ammary, F., . . . Quist, W. C. (2004).
Angiotensin receptor blockade with candesartan attenuates atherosclerosis, plaque disruption,
and macrophage accumulation within the plaque in a rabbit model. Circulation, 110(14), 2060-
2065. doi:10.1161/01.CIR.0000143627.55926.4C

Klein, R., Myers, C. E., Lee, K. E., Gangnon, R. E., Sivakumaran, T. A., Iyengar, S. K., & Klein, B. E. (2015).
Small Drusen and Age-Related Macular Degeneration: The Beaver Dam Eye Study. J Clin Med,
4(3), 424-440. doi:10.3390/jcm4030425

Landsman, L., Bar-On, L., Zernecke, A., Kim, K. W., Krauthgamer, R., Shagdarsuren, E., . . . Jung, S. (2009).
CX3CR1 is required for monocyte homeostasis and atherogenesis by promoting cell survival.
Blood, 113(4), 963-972. doi:10.1182/blood-2008-07-170787

Lee, M., Lee, Y., Song, J., Lee, J., & Chang, S. Y. (2018). Tissue-specific Role of CX3CR1 Expressing Immune
Cells and Their Relationships with Human Disease. Immune Netw, 18(1), e5.
doi:10.4110/in.2018.18.e5

Luhmann, U. F., Robbie, S., Munro, P. M., Barker, S. E., Duran, Y., Luong, V., . . . MacLaren, R. E. (2009).
The drusenlike phenotype in aging Ccl2-knockout mice is caused by an accelerated accumulation
of swollen autofluorescent subretinal macrophages. Invest Ophthalmol Vis Sci, 50(12), 5934-
5943. doi:10.1167/iovs.09-3462

Ly, A., Nivison-Smith, L., Assaad, N., & Kalloniatis, M. (2017). Fundus Autofluorescence in Age-related
Macular Degeneration. Optometry and vision science : official publication of the American
Academy of Optometry, 94(2), 246-259. doi:10.1097/OPX.0000000000000997

Ma, B., Dang, G., Yang, S., Duan, L., & Zhang, Y. (2015). CX3CR1 polymorphisms and the risk of age-
related macular degeneration. Int J Clin Exp Pathol, 8(8), 9592-9596. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/26464724

Meghraoui-Kheddar, A., Barthelemy, S., Boissonnas, A., & Combadiere, C. (2020). Revising CX3CR1
Expression on Murine Classical and Non-classical Monocytes. Front Immunol, 11, 1117.
doi:10.3389/fimmu.2020.01117

Meng, X. M., Nikolic-Paterson, D. J., & Lan, H. Y. (2016). TGF-beta: the master regulator of fibrosis. Nat
Rev Nephrol, 12(6), 325-338. doi:10.1038/nrneph.2016.48

Nagai, H., & Kalnins, V. I. (1996). Normally occurring loss of single cells and repair of resulting defects in
retinal pigment epithelium in situ. Exp Eye Res, 62(1), 55-61. doi:10.1006/exer.1996.0007

62
Phipps, J. A., Vessey, K. A., Brandli, A., Nag, N., Tran, M. X., Jobling, A. I., & Fletcher, E. L. (2018). The Role
of Angiotensin II/AT1 Receptor Signaling in Regulating Retinal Microglial Activation. Invest
Ophthalmol Vis Sci, 59(1), 487-498. doi:10.1167/iovs.17-22416

Raoul, W., Feumi, C., Keller, N., Lavalette, S., Houssier, M., Behar-Cohen, F., . . . Sennlaub, F. (2008).
Lipid-bloated subretinal microglial cells are at the origin of drusen appearance in CX3CR1-
deficient mice. Ophthalmic Res, 40(3-4), 115-119. doi:10.1159/000119860

Richert, E., Koinzer, S., Tode, J., Schlott, K., Brinkmann, R., Hillenkamp, J., . . . Roider, J. (2018). Release of
Different Cell Mediators During Retinal Pigment Epithelium Regeneration Following Selective
Retina Therapy. Invest Ophthalmol Vis Sci, 59(3), 1323-1331. doi:10.1167/iovs.17-23163

Rodgers, K., Abiko, M., Girgis, W., St Amand, K., Campeau, J., & diZerega, G. (1997). Acceleration of
dermal tissue repair by angiotensin II. Wound Repair Regen, 5(2), 175-183. doi:10.1046/j.1524-
475X.1997.50210.x

Rodgers, K. E., Espinoza, T., Felix, J., Roda, N., Maldonado, S., & diZerega, G. (2003). Acceleration of
healing, reduction of fibrotic scar, and normalization of tissue architecture by an angiotensin
analogue, NorLeu3-A(1-7). Plast Reconstr Surg, 111(3), 1195-1206.
doi:10.1097/01.PRS.0000047403.23105.66

Rodgers, K. E., Roda, N., Felix, J. E., Espinoza, T., Maldonado, S., & diZerega, G. (2003). Histological
evaluation of the effects of angiotensin peptides on wound repair in diabetic mice. Exp
Dermatol, 12(6), 784-790. doi:10.1111/j.0906-6705.2003.00087.x

Tuo, J., Bojanowski, C. M., Zhou, M., Shen, D., Ross, R. J., Rosenberg, K. I., . . . Chan, C. C. (2007). Murine
ccl2/cx3cr1 deficiency results in retinal lesions mimicking human age-related macular
degeneration. Invest Ophthalmol Vis Sci, 48(8), 3827-3836. doi:10.1167/iovs.07-0051

Wilkinson-Berka, J. L., Suphapimol, V., Jerome, J. R., Deliyanti, D., & Allingham, M. J. (2019). Angiotensin
II and aldosterone in retinal vasculopathy and inflammation. Exp Eye Res, 187, 107766.
doi:10.1016/j.exer.2019.107766

Wilson, A. S., Hobbs, B. G., Shen, W. Y., Speed, T. P., Schmidt, U., Begley, C. G., & Rakoczy, P. E. (2003).
Argon laser photocoagulation-induced modification of gene expression in the retina. Invest
Ophthalmol Vis Sci, 44(4), 1426-1434. doi:10.1167/iovs.02-0622

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Chapter 3: Characterization of Selective Retinal Therapy and Wound Healing
Response in Soft Drusen Mouse Model of Age-Related Macular Degeneration
3.1 Introduction
In the previous chapter, the effects of R:GEN were characterized using a hard drusen mouse
model. Images and histology indicated that R:GEN was able to selectively target RPE and treated eyes
were capable of recovering from the induced lesions. Gene expression changes, and LC-MS/MS
metabolomics showed modulation of the RAS, along with confirming the activation of extracellular matrix
turnover as reported in literature. However, the presence of soft drusen is a greater risk factor for the
development and progression of dAMD, for which therapeutic interventions are more critical. Therefore,
it is critical to explore the safety and efficacy of the R:GEN laser in an animal model that develops soft
drusen to better represent a a patient who is developing dAMD. In this chapter, I present our findings on
the effect of R:GEN on CxCr5 -/- mice, a soft drusen animal model.
CxCr5 is a chemokine receptor expressed on B cells and essential to the trafficking of these
immune cells to lymphoid follicles (Crotty, 2011; Gunn et al., 1998). B-lymphocytes have previously been
found to be protective in limiting the progression of atherosclerosis, while deficiency of B-cells have
increased atherosclerotic lesions (Caligiuri, Nicoletti, Poirier, & Hansson, 2002; Major, Fazio, & Linton,
2002). Due to the similarity between atherosclerotic plaques and drusen, it could be surmised from the
literature that B-lymphocytes may play a role in the formation of drusen or AMD pathogenesis.
Although there has been increasing evidence that the immune system plays a pivotal part in the
pathogenesis of AMD, the role of B-lymphocytes in AMD is still relatively unclear (Frederick & Kleinman,
2014; Hampe, 2012; Küppers, 2005). Previously, Patel et al reported the presence of circulating
autoantibodies against retinal antigens and found both patients with drusen deposits and patients with
choroidal neovascularization to have significantly higher titers of these antibodies than control subjects
(Patel et al., 2005). This would suggest the involvement of B-lymphocytes in AMD as they can give rise to

64
autoantibodies. However, because Hector et al found no statistically significant differences between the
circulating B-lymphocytes of patients with wet AMD and controls, this might suggest the involvement of
B-lymphocytes to be closer related to dry AMD (Hector & Sørensen, 2017).
While the role that B-lymphocytes may play in the development of AMD has not yet been fully
characterized, CxCr5-kockout mice have previously been shown to develop characteristics representative
of early and late state AMD, including the formation of drusen-like subretinal deposits, RPE atrophy, and
photoreceptor death (Huang, Liu, Wang, & Li, 2017). Specifically, the subretinal deposits are described as
soft drusen-like in appearance, lacking a distinct border, making CxCr5 -/- mice a reasonable soft drusen
mouse model for testing the safety and efficacy of R:GEN. Huang et al also found CxCr5 to be upregulated
in the retinal tissues of aged healthy C57/Bl mice in comparison to the retinal tissue of young C57/Bl mice,
which led to their conclusion that CxCr5 may protect the RPE and retina from degeneration with age
(Huang et al., 2017). More recently, it has been shown that CxCr5 deficiencies cause dysfunction of RPE
cells, resulting in abnormal cell shape and pigmentation, as well as compromised barrier function,
underscoring the importance of CxCr5 in maintaining RPE health (Lennikov, Mukwaya, Saddala, & Huang,
2021).
In this chapter, a comprehensive evaluation was under taken with R:GEN treatment in CxCr5-/-
mice, our chosen soft drusen mouse model. I sought to identify if and how gene expression and
metabolomics may vary with a dose escalation design. Similar to the previous chapter, the impact of
R:GEN treatment on RAS was also explored, expecting a similar shift towards non-fibrotic wound healing.
Furthermore, while the relationship between ocular and systemic RAS peptides after R:GEN treatment
was evaluated in the previous chapter, evaluating correlations between systemic and ocular levels of
peptides for potential blood-based markers, I take one step further in identifying a dose-response in
ocular levels of peptides from increased laser energy.

65
3.2 Materials and Methods
3.2.1 Animals
Experiments were approved by the Institutional Animal Care and Use Committee of University of
Southern California and followed the National Institutes of Health Guidelines for the Care and Use of
Laboratory Animals. Male and female CxCr5 -/- (B6.129S2(Cg)-Cxcr
5tm1Lipp
/J) mice were purchased from
Jackson Laboratories (Jackson Laboratory, Bar Harbor, ME), which were aged to at least 12 months to
emulate the humans AMD disease.
Anesthesia and Animal Preparation
Prior to R:GEN laser treatment, mice were anesthetized with a ketamine (80-100 mg/kg) and
xylazine (5-10 mg/kg) mixture given as an intraperitoneal (IP) injection. After sedation, each eye was
sterilely prepared and dilated using topical 2.5% phenylephrine HCL and 0.5% tropicamide, approximately
50 µL of each solution. In addition, approximately 50 µL of 0.5% tetracaine was also applied as a local
anesthetic. Pre-treatment images of the optic nerve and retina of each eye were acquired using the Envisu
R-Class Bioptigen System (Leica Microsystems Inc., Buffalo Grove, IL). Optical coherence tomography
(OCT) retinal sections were also acquired using HRA+OCT Spectralis (Heidelberg Engineering Inc., Franklin,
MA).

66
R:GEN Laser Application
Each mouse eye was administered the designated dosage of R:GEN laser on Day 0, where a
summary of the settings and parameters are summarized in Table 3.1. To each eye, 20 laser spots were
applied in a circular pattern around the optical nerve. The duration of each laser pulse used was 1.7 µs,
where energy auto-ramping delivered between 12.5 µJ and 17.5 µJ per lesion. The optoacoustic (OA) and
reflectometry (RFM) of the dual-dosimetry mechanism were established, such that the RTF of each lesion
would be either 50% or 100% intensity. The laser induced a resultant RPE lesion with a diameter of
approximately 90µm.
Following laser application, mice were allowed to recover from treatment until pre-determined
designated timepoints at 0.04, 0.16, 3, 7, 30, and 90 days, where N=8 were euthanized. The overall study
design is summarized in the Table 3.2.
End of Study Procedures
Prior to euthanasia, mice were anesthetized with ketamine (80-100 mg/kg) and xylazine (5-10
mg/kg) mixture given IP. Each eye was dilated and locally anaesthetized as described above. Images of the
optic nerve and retina of each eye were again acquired using the Envisu R-Class Bioptigen System (Leica
Microsystems Inc., Buffalo Grove, IL). In addition, 0.01 mL of a 10% fluorescein solution was administered
as an IP injection and fluorescein angiography (FA) was perform to determine the levels of leakage. OCT
Specifications R:GEN Laser System
Laser Wavelength (nm) 527nm
Aiming Beam 635 nm (<1mW)
Energy 30 µJ ~ 400 µJ [for mice : 5µJ ~ 100 µJ]
Pulse Duration 1.7μs
Pulse Rate ≤ 100Hz
Spot Size 200μm (for mice: ~90um)
Number of Pulses per Shot Up tp 15
Number of Spots 20
RTF sensors Optoacoustic (OA)
Reflectometry (RM)
Operation Mode Guidance Mode (AutoStop) SRT
Patterns Single
Table 3.1: R:GEN laser setting for mice retinal application

67
sections and FA were simultaneously acquired using HRA+OCT Spectralis (Heidelberg Engineering Inc.,
Franklin, MA).
Upon completion of imaging, mice were immediately euthanized via isoflurane overdose,
cardiac puncture, and cervical dislocation. All animal procedures followed Institutional Animal Care and
Use Committee (IACUC) approved protocols.
3.2.2 Preparation of Blood Plasma for LC-MS/MS Analysis
A 26-gauge needle (Becton Dickinson, Franklin Lakes, IL) was heparinized (West-Ward,
Eatontown, NJ) for cardiac puncture. Approximately 1 mL of whole blood was collected and immediately
transferred to a clean 1.5 mL microcentrifuge tube containing 20 µL of protease inhibitor cocktail (PIC) as
detailed in chapter 2. The blood-filled tubes were gently inverted three times to ensure thorough mixing
of the PIC and immediately placed on ice. Blood samples were then centrifuged at 2,000 g at 4°C for 15
minutes. Plasma was aliquoted into a new, clean 1.5 mL microcentrifuge tube and immediately frozen at
-80°C until analysis.
Type of
Model
R:GEN
Intensity
Timepoint
(Days)
Designation Number
of
Animals
EOS Sample Collection
CxCr5 -/-
100% 0.04 H 8 In Life:
Baseline OCT, FA and FAF at baseline and
end of study analyses

End of Study:
• Blood and Ocular Fluids for RAS
metabolomics.
• Designated number of eyes for
flatmount and H&E analyses
• RT-PCR for retina and RPE for MMPs
and RAS components
100% 0.16 I 8
100% 3 J 8
100% 7 K 8
100% 30 L 8
100% 90 M 8
50% 0.04 N 8
50% 0.16 O 8
50% 3 P 8
50% 7 Q 8
50% 30 R 8
50% 90 S 8
NT 0 T 8
Table 3.2: Study Design for CxCr5-/- soft drusen mice model

68
3.2.3 Preparation of Eyes
Both eyes were enucleated using curved forceps (World Precision Instruments, Sarasota, FL) and
placed in separate in sterile petri dishes (VWR, Radnor, PA) containing balanced salt solution (BSS) (Alcon,
Geneva, Switzerland) corresponding to each eye (OD, OS).
3.2.3.1 Preparation of Eyes for Flat Mount Staining
Three eyes from each timepoint were utilized for flat mount to visualize R:GEN induced lesion
closure over time. For each eye, the surrounding connective muscle and fat tissue were carefully excised
using Vannas scissors (World Precision Instruments, Sarasota, FL). The optic nerve was cut off, leaving a
small stump to act as a landmark and avoid damaging any of the posterior eye segment. After removing
all excess tissue, the eye was placed on a labeled, clean glass slide (VWR, Radnor, PA), encircled by a
silicone isolator (Grace Bio-Labs, Bend, OR). A couple drops, approximately 50 µL, of balanced salt solution
(BSS) were applied to the eye. A 31-gauge needle (Becton Dickinson, Franklin Lakes, NJ) was used to
puncture the cornea, release the aqueous humor (AQH), and deflate the eye. Four radial cuts were then
made from the puncture wound made by the 31-gauge needle: from the center of the cornea to the edge
of the remaining optic nerve. The tissue was then flattened while simultaneously rolling the lens to
separate it from the retina. To the flattened RPE-sclera, 300 µL of 10% formalin (VWR, Radnor, PA) was
applied. A glass cover slip (VWR, Radnor, PA) was then placed on the silicone isolator and the slide was
secured for staining.
The glass coverslips were removed from the silicone isolators and the 10% formalin was aspirated.
The fixed RPE was washed three times by immersing the tissue in Hank’s Balanced Salt Solution (Corning,
Corning, NY) for 10 minutes per wash. Tissue was then permeabilized by incubating with prepared wash
buffer for 5 minutes. The wash buffer was then aspirated. The RPE was then incubated with 300μL of
prepared blocking buffer for 1 hour at room temperature in a humidified chamber (Sigma-Aldrich, St.
Louis, MO). The blocking buffer was then aspirated with a transfer pipet (VWR, Radnor, PA). The flat

69
mount was then incubated with 300 μL of prepared primary antibody solution for 16 hours at room
temperature in a humidified chamber. The primary antibody was then aspirated with a transfer pipet and
the tissue was subsequently washed with the prepared wash buffer 5 times, for 2 minutes per wash. The
flat mount was then incubated with the prepared secondary antibody for 1 hour. After aspirating the
secondary antibody, the flat mount was then rinsed with prepared propidium iodide solution 5 times, for
2 minutes per wash. The flat mount was then washed with the prepared wash buffer twice, for 2 min per
wash. 25 μL of Vectashield was then applied to the flat mount, and a cover slip was secured on top. The
finished slide was then cured overnight under refrigerated conditions. Images were taken on an Olympus
BX43 microscope (Olympus America Inc., PA)
To stain the fixed RPE tissue for tight junctions, solutions were prepared as reported in Chapter
2.
3.2.3.2 Preparation of Eyes for Histopathology
Enucleated eyes were placed in their respectively labeled 10 mL scintillation vials (VWR, Radnor,
PA) containing Davidson’s fixative solution (VWR, Radnor, PA). After 72 hours of fixation, the Davidson’s
solution was replaced with ethanol until tissue was embedded in paraffin blocks for sectioning.
The cornea and lens of each preserved eye was removed. The posterior pole of each eye was then
embedded on a paraffin block. Each eye was then cut in half through the optic nerve, with one half of the
block reserved and the remaining half sectioned. Without eliminating any segment, sections were cut
every 5-7 microns throughout the block. 4-5 sections were then mounted per slide. All slides were then
stained with H&E. Images were taken on the Aperio CS2 (Leica Biosystems, Deer Park, IL).
3.2.3.3 Preparation of Eyes for RT-PCR
For each eye, the surrounding connective muscle and fat tissue were carefully excised using
Vanass scissors. The optic nerve was cut off, leaving a small stump to act as a landmark and avoid
damaging any of the posterior eye segment. After removing all excess tissue, the eye was placed on a

70
labeled, clean glass slide, encircled by a silicone isolator. A couple drops, approximately 50 µL, of balanced
salt solution (BSS) were applied to the eye. A 31-gauge needle was used to puncture the cornea, release
the aqueous humor (AQH). Four radial cuts were then made from the puncture wound made by the 31-
gauge needle: from the center of the cornea to the edge of the remaining optic nerve. The tissue was then
flattened while simultaneously rolling the lens to separate it from the retina and discard.
Approximately 450 µL of BSS was then used to wash and lift the photoreceptors from the RPE-
sclera layer. The photoreceptors were subsequently removed and placed in a labeled clean 1.5 mL
microcentrifuge tube containing 500 µL of TRIzol (Invitrogen, Carlsbad, CA). The RPE-sclera was placed in
a separate, labeled clean microcentrifuge tube containing 500 µL of TRIzol. The BSS solution,
approximately 500 µL, was then collected into a sterile 1.5 mL microcentrifuge tube containing prepared
PIC as summarized in the table above. The retina and RPE tissues were then frozen at -80°C for storage.
3.2.6 RT-PCR Analysis
3.2.6.1 Retina RNA Extraction
One scoop of stainless-steel beads was added to the retina tissue suspended in 500 µL of TRIzol.
Samples were placed in a TissueLyser II (QIAGEN, Germantown, MD) and homogenized for 2 minutes at
30 Hz. Samples were then centrifuged at 13,000g for 10 minutes at 4°C. RNA was extracted according to
the manufacturer’s protocol. RNA concentration and purity were measured using a NanoDrop
spectrophotometer (Thermo Fisher Scientific, Wilmington, DE). RNA concentration was adjusted to 10
ng/µL using nuclease-free water.
3.2.6.2 Retina RPE Extraction
One scoop of stainless-steel beads was added to the retina tissue suspended in 500 µL of TRIzol.
Samples were placed in a TissueLyser II (QIAGEN, Germantown, MD) and homogenized for 2 minutes at
30 Hz. Samples were checked for presence of pigment release after homogenization and homogenized
again if needed. Samples were then centrifuged at 13,000g for 10 minutes at 4°C. RNA was extracted

71
according to the manufacturer’s protocol. RNA concentration and purity were measured using a
NanoDrop spectrophotometer (Thermo Fisher Scientific, Wilmington, DE). RNA concentration was
adjusted to 10 ng/µL using nuclease-free water.
3.2.6.3 cDNA Synthesis
cDNA was synthesized using the extracted retina and RPE RNA using the RevertAid RT Reverse
Transcription kit (Thermo Fisher Scientific, Wilmington, DE) according to the manufacturer’s protocol.
Samples were placed in a Bio-Rad Thermal Cycler (Bio-Rad, Hercules, CA), where they were incubated for
5 min at 25 °C followed by 60 min at 42 °C, and finally terminated by heating at 70 °C for 5 min. 3.2.6.4
cDNA Pre-amplification
The 20 µL of prepared cDNA was pre-amplified by adding 25 µL SsoAdvanced PreAmp Supermix
(Bio-Rad, Hercules, CA) and 5 µL of a prepared primer assay pool, for a final volume of 50 µL. The primer
assay pool included primer stocks of all genes that would be analyzed for RT-PCR, such that the final
preamplification reaction contains 50 nM of each primer, summarized in the table below. The cDNA
reaction mix was set for 12 cycles, for which the first stage was 3 minutes at 95°C for polymerase
activation, followed by 15 seconds for denaturation, and 4 minutes at 58°C for annealing/extension.
3.2.6.5 RT-PCR for Targeted Genes
Genes related to wound healing and extracellular matrix remodeling were probed alongside the
Renin-Angiotensin System. The RT-PCR master mix was prepared by mixing Power Up SYBR Green master-
mix (Applied Biosystems, Foster City, CA) and the forward and reverse primers for each gene. The primers
sequences used were as listed in chapter 2.
Prepared master mix and diluted cDNA were pipetted into a 384-well plate (VWR, Radnor, PA) by
the Integra Assist Plus (Integra Biosciences, Hudson, NH). Samples were loaded in triplicates, plates were
sealed with an adhesive plate sealer, and centrifuged at 1000 g for 1 minute. The run method consisted
of an initial hold for 2 minutes at 50°C, followed by 95°C for 2 minutes. The amplification stage was set for

72
40 cycles, with a 1 second hold at 95°C and 30 seconds at 60°C for annealing/elongation. The final melt
curve stage was set for 15 seconds going from 1.6°C to 95°C, followed by 1 minute from 1.6°C to 60°C,
and finally 0.15°C to 95°C for 15 seconds. The data was collected and analyzed using Data Assist
(Invitrogen, Carlsbad, CA). Gene expression of MMP-2, MMP-3, MMP-9, TIMP-2, TGF- β1, ACE, ACE2,
AT1R, AT2R, and MasR were assessed using delta-delta CT, normalized to β-Actin as the reference gene
and compared to untreated animals. Fold changes were plotted in GraphPad Prism (GraphPad, San Diego,
CA)
3.2.7 RAS Metabolomics Using Quantitive LC-MS/MS Analysis
3.2.7.1 Solid Phase Extraction of Plasma and Ocular Fluid Samples
Frozen plasma and ocular fluid samples were thawed on ice, and from each sample, 150 µL was
aliquoted into a clean 1.5 mL microcentrifuge tube. Each sample was then spiked with 150 µL of a
prepared 40 ng/mL NorLeu
3
-Angiotensin(1-7) (NLE) internal standard solution. Samples were then
vortexed thoroughly and placed back on ice. While samples were incubating, Oasis MAX solid phase
extraction columns (Waters, Milford, MA) which were pre-conditioned using 1 mL of HPLC grade methanol
(VWR, Radnor, PA), followed by 1 mL of HPLC grade water (VWR, Radnor, PA).
Prepared samples were alkalinized by adding 700 µL of a 5% ammonium hydroxide solution and
vortexed thoroughly. Dissociated samples (1000 µL) were then layered onto the primed solid phase
extraction (SPE) columns. Columns were then washed with 500 µL of 5% ammonium hydroxide, twice.
Columns were then washed with 500 µL of HPLC grade methanol and then dried for a couple minutes.
Samples were then eluted from the SPE columns with 500 µL of methanol with 2% formic acid, twice.
Eluted samples were then dried under inert nitrogen gas. The residue was then reconstituted using 50 µL
of 10 mM Ammonium Formate buffer, pH 4.2 and transferred to a clean HPLC vial (Sun-Sri, Rockwood,
TN), where 40 µL were injected into an API4000 (AB Sciex, Framingham, MA) for analysis.

73
3.2.7.2 Targeted LC-MS/MS Instrumentation and Conditions
RAS peptides were separated and quantified using an Agilent 1200 HPLC (Agilent, Santa Clara, CA)
connected to an API 4000 triple quad mass spectrometer (AB Sciex, Framingham, MA). Analyte data was
acquired using Analyst 1.6.2, where each analyte was quantified using multiple reaction monitoring
(MRM) identified during manual tuning of each compound set in the positive mode. The MRM’s of the
peptides included in the method are as summarized in chapter 2.
Peptides were separated using an Agilent Poroshell 120 EC-C18 (4.6X100 mm, 2.7μm) column
(Agilent, Santa Clara, CA). Analytes were eluted using a gradient mobile phase consisting of two solvents:
1) Solvent A was HPLC grade water with 0.2% formic acid; and 2) Solvent B was HPLC grade methanol with
0.2% formic acid. The HPLC parameters and established gradient program are summarized in chapter 2.
Flow injection analysis (FIA) was performed to maximize the sensitivity of each MRM established
in the method. The optimized source temperature, gas pressures, and ion spray voltage are summarized
in chapter 2.
3.2.8 Statistical Analysis
Results are expressed as mean ± SD unless specified otherwise. GraphPad Prism (GraphPad, San
Diego, CA), R (R Foundation for Statistical Computing, Vienna, Austria), and RStudio (RStudio Inc, Boston,
MA) were used for data preparation, data cleaning, statistical analysis, and data visualization on
quantitative and qualitative data.

74
3.3 Results
3.3.1 Applied Laser Settings
The energies (µJ) for R:GEN applied laser spots delivered to each eye are summarized in Figure
3.1 below. In Figure 3.1A, Blue and Red points indicate the total amount of energy given to mice that were
assigned to either 50% or 100% intensity group (Figure 1A). Mice assigned to 100% intensity group
received a total energy that ranged between 1200 to 1253 µJ. In contrast, mice assigned to 50% intensity
had a wider range of energy between 1100 and 1253 µJ. For a large number of animals in the 50% intensity
treatment group, the energy applied to each eye did not drop below 1100 µJ. In this cohort, 66.3%
received 1253 µJ of energy regardless which group they were assigned, as seen in the distribution of
energy delivered shown in Figure 3.1B.

Figure 3.1: Total energy delivered to each treated eye indicate minimal difference in µJ between 50% and 100%
intensity (A). Distribution of applied energies indicate most eyes were treated with maximum energy of 1253
µJ (B).
A B

75
3.3.2 Fundus Images
Following R:GEN treatment at either 50% or 100% intensity, fundus imaging was assessed for
each time interval. Representative images are summarized in Figure 3.2. Visible laser induced lesions
were seen after 0.04 and 0.16 days. Lesions at these timepoint have evidence of edema. In contrast,
residual lesion markings were observed at 3, 7, and 30 days showing full lesion closure and resolution by
Day 90. Unsurprisingly, given the total energy delivered per eye were not much different between the
two treatment groups, FA difference between the 50% and 100% intensities were not detectable.

Figure 3.2: Fundus images of CxCr5 -/- soft drusen (SD) retina via Bioptigen after R:GEN treatment at 100% and
50% intensity indicate presence of lesions until 90 days after treatment.
NT
7 Days 30 Days
0.16 Days
90 Days
3 Days 0.04 Days
So0 Drusen
100%
NT
0.04 Days
7 Days 30 Days
0.16 Days
90 Days
3 Days
So0 Drusen
50%

76
3.3.3 Fluorescein Angiography
To confirm damage to the RPE, visible signs of fluorescein leakage from lesions were counted and
scored as following the table below. Dunn’s multiple comparison test was used to compare each time
point against Day 0.04 for wound closure and statistical significance.

Figure 3.3: Representative fluorescein angiography of CxCr5 -/- soft drusen (SD) retina via using HRA+OCT
Spectralis at end of study (EOS) time points indicate presence of open lesions. Clear signs of fluorescein leakage
were visible at 0.04 and 0.16 days, with minimal haze remaining at 3 and 7 days, and zero leakage at 30 and 90
days.
NT
0.04 Days
7 Days 30 Days
0.16 Days
90 Days
3 Days
Score Criteria
1 Slight haze
2 Hazy but clear core, hazy perimeter
3 Bright core, hazy perimeter
4 Bright core, bright perimeter
Table 3.3: Lesion severity scoring and criteria for fluorescein
angiography images

77
In general, the number of visible lesions after the first day of treatment were significantly reduced,
indicating definitive wound closure if fluorescein was no longer able to permeate at the site of injury. In
the 100% intensity treatment arm, lesion counts at 0.04 days were found to be significantly greater than
counts at 7, 30, and 90 days, but not at 0.16 or 3 days, as shown in Figure 3.4. Simultaneously, it was
observed that the severity of lesions had also decreased, as grade 3 and 4 lesions could no longer be
determined on day 3, with the majority of remaining lesions being grade 1, or the mildest case.

In the 50% intensity treatment arm, the lesion counts at 0.04 days were found to be significantly
greater than counts at 3, 30, and 90 days, but not at 0.16 or 7 days as shown in Figure 3.5. Simultaneously,

Dunn's multiple
comparisons test Mean rank diff. Summary Adjusted P Value
0.04 vs. 0.16 5.794 ns >0.9999
0.04 vs. 3 18.64 ns 0.2468
0.04 vs. 7 26.24 ** 0.0023
0.04 vs. 30 36.94 **** <0.0001
0.04 vs. 90 41.44 **** <0.0001

Figure 3.4: Assessment and scoring of visible lesions in CxCr5 -/- mouse at 100% R:GEN intensity (SD100). Total
lesion counts per eye (A), average count of lesion severity (B)

A B

78
it was observed that the severity of lesions had also decreased, as grade 3 and 4 lesions could no longer
be determined on day 3, with the majority of remaining lesions being grade 1, or the mildest case.

3.3.4 Flat Mount Staining for Tight Junctions
The RPE layer is known to normally exist in a cobble-stoned hexagonal pattern, forming tight
junctions that comprise the blood retinal barrier. Polymegathism, a variation in cell size within the
endothelial monolayer, has previously been reported to occur when cells of the RPE layer grow larger to
replace dying RPE (Nagai & Kalnins, 1996). To assess the extent of lesion damage to the RPE, flat mounts
were fixed and stained with anti-Zo-1 antibodies to visualize the tight junctions and examine cell
morphology at the designated time points. The wound repair response was assessed, and it was confirmed
in this study that polymegathism occurred to repair the laser-induced lesions, as shown in Figure 3.6
below. Cell membrane disruption from R:GEN treatment is evident at 0.04 and 0.16 days, as a clear

Dunn's multiple
comparisons test Mean rank diff. Summary Adjusted P Value
0.04 vs. 0.16 -5.242 ns >0.9999
0.04 vs. 3 26.98 ** 0.0012
0.04 vs. 7 14.39 ns 0.2224
0.04 vs. 30 29.76 **** <0.0001
0.04 vs. 90 36.59 **** <0.0001

Figure 3.5: Assessment and scoring of visible lesions in CxCr5 -/- mouse at 50% R:GEN intensity (SD50). Total
lesion counts per eye (A), average count of lesion severity (B)
A B

79
absence of tight junctions is visible, and signs of edema are present. Signs of elongation begin to appear
at 0.16 days and are more obvious by 3 days, indicating the surrounding RPE are extending to close the
lesion.

3.3.5 H&E Assessment of R:GEN Lesions
To evaluate the specificity of the R:GEN laser in its ability to selectively target the RPE layer,
treated eyes were enucleated at the designated EOS time points, fixed with Davidson’s solution, and
stained with hematoxylin and eosin (H&E). Tissue sections were imaged at 40x magnification and analyzed
for lesions. Distinct breakages in the RPE layer are visible at 0.16 Days, indicating disruption of the cell
membrane via R:GEN treatment. At Day 3, signs of pigmentation re-emerge as the RPE surrounding the
induced lesions begin to elongate and fill in the gap. The induced lesions are mostly closed by Day 7,
although signs of disruption, such as unevenness of the pigmented layer still exist. Most notably, there
are no discernible changes to the inner nuclear layer (INL) nor outer nuclear layer (ONL). The unaffected

Figure 3.6: Zo-1 staining of isolated CxCr5 -/- RPE layer treated with 100% intensity. Images are at 40x
magnification; scale bar = 20 µm. Signs of laser-induced lesions are observed at 0.04, 0.16, and 3 days.
Evidence of cell membrane elongation begins to appear at 0.16 day to close lesions. Polymegathism is seen at
7, 30, and 90 days as abnormally large RPE fill in the damaged regions.
7 Day – 40X 30 Day – 40X 90 Day – 40X
3 Day – 40X 0.16 Day – 40X 0.04 Day – 40X

80
photoreceptors indicate the R:GEN laser is selective for the melanosomes of the RPE and avoids the
trauma associated with traditional photocoagulation.

3.3.6 RT-PCR
While the histology of laser-treated eyes indicated the R:GEN is selective for RPE, RT-PCR was
conducted to investigate and characterize the underlying mechanism of injury and wound repair in retinal
tissues. Tukey Fences for Outliers was applied to Delta CT (dCT) calculations to identify and remove
outliers prior to fold change analysis. Metalloproteinases (MMP) are enzymes that play an integral role in
ECM remodeling and tissue repair. Gene expression for MMP-2, MMP-3, and MMP-9 of RPE were seen to
be upregulated across all time points in both treatment arms of 50% and 100% intensity. Furthermore,

Figure 3.7: Representative H&E images for CxCr5 -/- mice treated with 100% intensity. Images are at 40x
magnification. Signs of laser-induced lesions are observed at 0.16 and 3 days and circled in red. Disruption of
the RPE is evident from the loss of pigment seen at 0.16 days. Pigment appears to fill in the induced lesion on
Day 3, with majority of lesion closure by Day 7, although uneven pigment indicates previous damage.
CXCR5 100% – 0.16 Days – 40x CXCR5 100% – 7 Day – 40x
CXCR5 100% – 3 Day – 40x
CXCR5 – NT – 40x

81
there was a significant downregulation of TIMP-2, the natural inhibitor of MMP-2, at 0.16 and 30 Days of
the 100% intensity arm, as shown in Figure 3.8.

Figure 3.8: Observed changes in laser-treated RPE gene expression of metalloproteinases relative to untreated.
MMP-2 (A), MMP-3 (B), MMP-9 (C), TIMP-2 (D)
NT 0.04 0.16 3 7 30 90
0.1
1
10
100
1000
10000
Time (Days)
Fold Change from Untreated
MMP2
100%
50%
NT 0.04 0.16 3 7 30 90
0.1
1
10
100
1000
10000
Time (Days)
Fold Change from Untreated
MMP9
100%
50%
NT 0.04 0.16 3 7 30 90
0.1
1
10
100
1000
10000
Time (Days)
Fold Change from Untreated
MMP3
100%
50%
NT 0.04 0.16 3 7 30 90
0.1
1
10
100
1000
10000
Time (Days)
Fold Change from Untreated
TIMP2
100%
50%
A
D
B
C

82
The gene expression of RAS components (e.g., ACE1, ACE2, AT1R, AT2R, and MasR) were also
probed to determine the impact of R:GEN on RAS metabolism of the RPE. ACE1 was found to be
significantly upregulated in all time points at the 50% intensity. In contrast, gene expression of ACE1 at
the 100% intensity was significantly upregulated at 0.04, 30, and 90 days but downregulated at 0.16, 3,
and 7 days. ACE2 was found to only be significantly upregulated at 90 days, where its expression was
either significantly downregulated or unchanged. AT1R was found to be significantly upregulated for both
50% and 100% intensity at 0.04, 0.16, and 90 days, with no change at 3 and 7 days. Gene expression of
AT2R was found to be significantly upregulated for both 50% and 100% intensity at all time points. Findings
are summarized in Figure 3.9.
Gene expression of MasR was significantly upregulated for the 100% intensity treatment arm at
0.04, 30, and 90 days, but significantly downregulated at 0.16, 3, and 7 days. For the 50% intensity
treatment arm, MasR expression was significantly upregulated at 0.16, 3, and 90 days but significantly

Figure 3.9: Observed changes in laser-treated RPE gene expression of metalloproteinases relative to untreated.
ACE1 (A), ACE2 (B), AT1R (C), AT2R (D)
NT 0.04 0.16 3 7 30 90
0.01
0.1
1
10
100
1000
10000
100000
Time (Days)
Fold Change from Untreated
ACE1
100%
50%
NT 0.04 0.16 3 7 30 90
0.01
0.1
1
10
100
1000
10000
100000
Time (Days)
Fold Change from Untreated
AT1R
100%
50%
NT 0.04 0.16 3 7 30 90
0.01
0.1
1
10
100
1000
10000
100000
Time (Days)
Fold Change from Untreated
ACE2
100%
50%
NT 0.04 0.16 3 7 30 90
0.01
0.1
1
10
100
1000
10000
100000
Time (Days)
Fold Change from Untreated
AT2R
100%
50%
A
D
B
C

83
downregulated at 30 days. TGF- β1 gene expression for the 100% intensity arm was found to be
significantly upregulated at 0.04 and 90 days, but significantly downregulated at 0.16 and 30 days.
Findings are summarized in Figure 3.10 below.

3.3.7 LC-MS/MS
Renin Angiotensin Peptide Systemic Metabolism
Collected plasma was analyzed for systemic changes in RAS peptides over the time after laser
treatment. Minimal fluctuations in systemic levels were observed for Ang(1-12) across the time period

Figure 3.10: Observed changes in laser-treated RPE gene expression of MasR relative to untreated. TGF- β1 (A),
MasR (B)
NT 0.04 0.16 3 7 30 90
0.001
0.01
0.1
1
10
100
1000
Time (Days)
Fold Change from Untreated
TGFB1
100%
50%
A
NT 0.04 0.16 3 7 30 90
0.1
1
10
100
1000
Time (Days)
Fold Change from Untreated
MasR
100%
50%
B

Figure 3.11: Plasma levels of Ang(1-12) after R:GEN treatment over a 90 day time course
Dunnett's multiple comparisons test
Predicted (LS)
mean diff. 95.00% CI of diff. Significant? Summary
Adjusted P
Value
50%
NT vs. 0.04 Day -0.2565 -0.8801 to 0.3671 No ns 0.7597
NT vs. 0.16 Day -0.06179 -0.6854 to 0.5618 No ns 0.9996
NT vs. 3 Day -0.3013 -0.9249 to 0.3223 No ns 0.6257
NT vs. 7 Day -0.2383 -0.8619 to 0.3853 No ns 0.8098
NT vs. 30 Day -0.008387 -0.6320 to 0.6152 No ns >0.9999
NT vs. 90 Day -0.03101 -0.6546 to 0.5926 No ns 0.9998
100%
NT vs. 0.04 Day 0.2553 -0.3697 to 0.8804 No ns 0.7701
NT vs. 0.16 Day 0.2687 -0.4070 to 0.9444 No ns 0.7890
NT vs. 3 Day 0.02021 -0.6049 to 0.6453 No ns 0.9999
NT vs. 7 Day -0.1422 -0.7673 to 0.4828 No ns 0.9785
NT vs. 30 Day 0.08171 -0.5434 to 0.7068 No ns 0.9982
NT vs. 90 Day 0.1077 -0.5437 to 0.7590 No ns 0.9953

84
where RAS peptides were measured. No statistical significance was found between when a Two-Way
ANOVA was conducted, as shown in Figure 3.11.
Systemic levels of Ang(1-10) or AngI, an inactive peptide precursor, were observed to stay low
during the first day of laser induction. Increased concentration of Ang(1-10) was seen starting on Days 3,
but dropped down to basal levels on Day 7. Interestingly, on Day 30, and 90 systemic Ang(1-10) levels
were elevated, and these time points were the only ones found to be statistically significant when
compared to untreated eyes, as seen in Figure 3.12.

The byproduct metabolites of Ang(1-10) include AngII, Ang(1-9) and Ang(1-7); peptides that
exhibit functionality in RAS through their binding of their respective receptors. Systemic AngII did not
fluctuate much, where changes were not significantly different. This may be due to the role of AngII in
maintaining hemodynamic blood pressure. Systemic levels of Ang(1-9) were elevated at 0.04 and 0.16
days after R:GEN treatment, where the changes were not statistically significant due to the high variability.
However on Day 3, the systemic levels of Ang(1-9) dropped significantly, and then returned to baseline

Figure 3.12: Plasma levels of Ang(1-10) after R:GEN treatment over a 90 day time course
Dunnett's multiple comparisons test
Predicted (LS)
mean diff. 95.00% CI of diff. Significant? Summary
Adjusted P
Value
50%
NT vs. 0.04 Day -0.1777 -0.5501 to 0.1948 No ns 0.4387
NT vs. 0.16 Day -0.1333 -0.4180 to 0.1514 No ns 0.4536
NT vs. 3 Day -4.077 -12.13 to 3.973 No ns 0.3911
NT vs. 7 Day -0.2794 -0.8573 to 0.2985 No ns 0.4280
NT vs. 30 Day -3.472 -6.156 to -0.7885 Yes * 0.0163
NT vs. 90 Day -4.584 -8.202 to -0.9658 Yes * 0.0180
100%
NT vs. 0.04 Day 0.009906 -0.2735 to 0.2933 No ns 0.9998
NT vs. 0.16 Day -0.07817 -0.6972 to 0.5409 No ns 0.9814
NT vs. 3 Day -3.494 -7.752 to 0.7641 No ns 0.1052
NT vs. 7 Day -0.1221 -0.4170 to 0.1728 No ns 0.5560
NT vs. 30 Day -6.505 -11.15 to -1.865 Yes * 0.0111
NT vs. 90 Day -3.335 -5.694 to -0.9765 Yes * 0.0153

85
on Day 7. On day 30 and 90, the circulating levels of Ang(1-9) was significantly lower than baseline levels,
shown in Figure 3.13 below.

Systemic levels of AngII were found to be relatively stable across all time points, except for 30
Days after R:GEN treatment using the 100% intensity. AngII on Day 30 was found to be lower than levels
of untreated animals, a statistically significant difference as indicated in Figure 3.14 .

Figure 3.13: Plasma levels of Ang(1-9) after R:GEN treatment over a 90 day time course

Figure 3.14 : Plasma levels of AngII after R:GEN treatment over a 90 day time course

Dunnett's multiple comparisons test
Predicted (LS)
mean diff. 95.00% CI of diff. Significant? Summary
Adjusted P
Value
50%
NT vs. 0.04 Day -0.3483 -0.7467 to 0.05005 No ns 0.0838
NT vs. 0.16 Day -0.5046 -1.063 to 0.05388 No ns 0.0742
NT vs. 3 Day 0.4475 0.1352 to 0.7598 Yes * 0.0100
NT vs. 7 Day -0.5445 -1.341 to 0.2522 No ns 0.1881
NT vs. 30 Day 0.4769 0.09898 to 0.8548 Yes * 0.0183
NT vs. 90 Day 0.4140 0.05408 to 0.7740 Yes * 0.0278
100%
NT vs. 0.04 Day -0.1961 -0.6706 to 0.2783 No ns 0.5572
NT vs. 0.16 Day -0.1817 -0.6638 to 0.3005 No ns 0.5214
NT vs. 3 Day 0.4542 0.004048 to 0.9043 Yes * 0.0482
NT vs. 7 Day -0.1276 -0.6964 to 0.4411 No ns 0.9142
NT vs. 30 Day 0.4584 0.09730 to 0.8195 Yes * 0.0178
NT vs. 90 Day 0.5018 0.2750 to 0.7287 Yes ** 0.0029
Dunnett's multiple comparisons test
Predicted (LS)
mean diff. 95.00% CI of diff. Significant? Summary
Adjusted P
Value
50%
NT vs. 0.04 Day -0.06523 -0.2459 to 0.1154 No ns 0.6589
NT vs. 0.16 Day -0.01075 -0.1268 to 0.1053 No ns 0.9982
NT vs. 3 Day 0.04594 -0.2962 to 0.3880 No ns 0.9900
NT vs. 7 Day -0.01547 -0.2293 to 0.1984 No ns 0.9996
NT vs. 30 Day -0.01769 -0.5589 to 0.5235 No ns 0.9999
NT vs. 90 Day 0.08161 -0.1477 to 0.3109 No ns 0.6693
100%
NT vs. 0.04 Day -0.03010 -0.1715 to 0.1114 No ns 0.9294
NT vs. 0.16 Day -0.02371 -0.2120 to 0.1646 No ns 0.9816
NT vs. 3 Day 0.08387 -0.06959 to 0.2373 No ns 0.3350
NT vs. 7 Day -0.06697 -0.2263 to 0.09235 No ns 0.5437
NT vs. 30 Day 0.1466 0.01553 to 0.2776 Yes * 0.0313
NT vs. 90 Day 0.1320 -0.04011 to 0.3041 No ns 0.1124

86
Interestingly, systemic Ang(1-7) dropped slightly (not statistically significant) at 0.04 and 0.16 days
after laser induction, where by days 3 and 7 the levels were at or above baseline levels. These differences
were not found to be statistically significant. Interestingly, Ang(1-7) levels dropped below baseline levels
on Day 30 and returned to baseline on Day 90, which was statistically significant for the 50% intensity
treatment arm, Figure 3.15.
The levels of metabolites downstream of AngII and Ang(1-7) were also evaluated. AngII
metabolite includes Ang(2-8) and Ang(3-8), where the N-terminal cleavage of amino acid(s) will form these
metabolites. Ang(1-7) can be metabolized to alamandine or Ang(1-5), which are ligands for MrG and Mas,
respectively. Systemic Ang(2-8) levels did not change between untreated and days after R:GEN treatment,
except for Day 90 of the 100% intensity treatment arm, as shown in Figure 3.16. While not statistically
significant overall, it is important to note that systemic levels were trending downwards on Day 30 and
90.

Figure 3.15: Plasma levels of Ang(1-7) after R:GEN treatment over a 90 day time course

Dunnett's multiple comparisons test
Predicted (LS)
mean diff. 95.00% CI of diff. Significant? Summary
Adjusted P
Value
50%
NT vs. 0.04 Day 0.3106 -0.1881 to 0.8094 No ns 0.2430
NT vs. 0.16 Day 0.2393 -0.1605 to 0.6392 No ns 0.2690
NT vs. 3 Day -0.06557 -1.576 to 1.445 No ns 0.9998
NT vs. 7 Day -0.2217 -0.9217 to 0.4783 No ns 0.7492
NT vs. 30 Day 0.5501 0.1271 to 0.9731 Yes * 0.0159
NT vs. 90 Day -0.3259 -1.517 to 0.8650 No ns 0.8323
100%
NT vs. 0.04 Day 0.1620 -0.1726 to 0.4967 No ns 0.4290
NT vs. 0.16 Day 0.2533 -0.05810 to 0.5648 No ns 0.0948
NT vs. 3 Day 0.1735 -0.8383 to 1.185 No ns 0.9702
NT vs. 7 Day 0.1913 -0.2905 to 0.6732 No ns 0.5892
NT vs. 30 Day 0.3828 -0.09063 to 0.8561 No ns 0.1107
NT vs. 90 Day 0.05815 -1.152 to 1.268 No ns 0.9997

87

Ang(3-8) increased significantly at 0.04 days after R:GEN treatment in the 50% intensity arm, but
no other time points were found to be statistically significant due to high variability, as seen in Figure
3.17. However levels appear to be elevated on Day 7, which was also seen with Ang(3-7).

Figure 3.16: Plasma levels of Ang(2-8) after R:GEN treatment over a 90 day time course

Dunnett's multiple comparisons test
Predicted (LS)
mean diff. 95.00% CI of diff. Significant? Summary
Adjusted P
Value
50%
NT vs. 0.04 Day -0.01924 -0.1440 to 0.1055 No ns 0.9806
NT vs. 0.16 Day 0.05348 -0.06926 to 0.1762 No ns 0.5117
NT vs. 3 Day 0.04641 -0.3855 to 0.4783 No ns 0.9962
NT vs. 7 Day -0.03774 -0.4009 to 0.3255 No ns 0.9966
NT vs. 30 Day 0.1884 -0.02418 to 0.4010 No ns 0.0797
NT vs. 90 Day 0.1857 -0.03029 to 0.4016 No ns 0.0891
100%
NT vs. 0.04 Day 0.03855 -0.06218 to 0.1393 No ns 0.6181
NT vs. 0.16 Day 0.02786 -0.05609 to 0.1118 No ns 0.6142
NT vs. 3 Day 0.1122 -0.08773 to 0.3120 No ns 0.3156
NT vs. 7 Day -0.03907 -0.1200 to 0.04186 No ns 0.4314
NT vs. 30 Day 0.1345 -0.06280 to 0.3319 No ns 0.1901
NT vs. 90 Day 0.2357 0.02071 to 0.4506 Yes * 0.0369

Figure 3.17: Plasma levels of Ang(3-8) after R:GEN treatment over a 90 day time course

Dunnett's multiple comparisons test
Predicted (LS)
mean diff. 95.00% CI of diff. Significant? Summary
Adjusted P
Value
50%
NT vs. 0.04 Day -0.3024 -0.5847 to -0.02016 Yes * 0.0377
NT vs. 0.16 Day -0.1250 -0.3106 to 0.06063 No ns 0.1962
NT vs. 3 Day -0.02901 -0.2523 to 0.1943 No ns 0.9916
NT vs. 7 Day -0.5530 -2.152 to 1.046 No ns 0.6899
NT vs. 30 Day 0.01949 -0.08580 to 0.1248 No ns 0.9578
NT vs. 90 Day -0.0007640 -0.07235 to 0.07082 No ns >0.9999
100%
NT vs. 0.04 Day -0.04055 -0.3133 to 0.2322 No ns 0.9842
NT vs. 0.16 Day -0.1463 -0.3802 to 0.08765 No ns 0.1939
NT vs. 3 Day -0.006602 -0.1467 to 0.1335 No ns 0.9997
NT vs. 7 Day -0.1426 -0.3901 to 0.1048 No ns 0.2962
NT vs. 30 Day -0.05923 -0.2247 to 0.1063 No ns 0.6688
NT vs. 90 Day 0.002595 -0.09257 to 0.09776 No ns 0.9999

88
For systemic levels of Ang(3-7), the 50% intensity treatment had increased levels across all time
points, but statistical significance was only found at 0.16, 3, and 30 days. With the 100% intensity
treatment groups, statistical significance was found at 0.04 and 7 days after laser induction, as seen in
Figure 3.18.

In contrast, Ang(1-5) was highly elevated with high variability on day 3 for both treatment groups,
with statistical significance in the 50% treatment arm at day 90, as seen in Figure 3.19. These findings
suggest a trend for which AngII and Ang(1-7) are metabolized into their down stream peptides starting
from Day 3 and continuing through to Day 7. Of the nine peptides measured, Ang(1-10), Ang(1-9), AngII,
and Ang(1-7) are of particular interest due to their roles in balancing the pathogenic and regenerative
activities associated with the RAS. Interestingly, it appears that systemic Ang(1-9) also fluctuate in
opposition to A(1-10).

Figure 3.18: Plasma levels of Ang(3-7) after R:GEN treatment over a 90 day time course

Dunnett's multiple comparisons test
Predicted (LS)
mean diff. 95.00% CI of diff. Significant? Summary
Adjusted P
Value
50%
NT vs. 0.04 Day -0.4377 -0.9225 to 0.04703 No ns 0.0744
NT vs. 0.16 Day -0.3630 -0.7013 to -0.02479 Yes * 0.0374
NT vs. 3 Day -0.4787 -0.8579 to -0.09949 Yes * 0.0183
NT vs. 7 Day -1.202 -4.196 to 1.792 No ns 0.5774
NT vs. 30 Day -0.7863 -1.542 to -0.03035 Yes * 0.0426
NT vs. 90 Day -0.5900 -1.203 to 0.02313 No ns 0.0583
100%
NT vs. 0.04 Day -1.113 -1.952 to -0.2741 Yes * 0.0145
NT vs. 0.16 Day -0.3868 -1.091 to 0.3177 No ns 0.2631
NT vs. 3 Day -0.1870 -0.4860 to 0.1119 No ns 0.2413
NT vs. 7 Day -0.7593 -1.159 to -0.3599 Yes ** 0.0024
NT vs. 30 Day -0.4235 -0.9905 to 0.1436 No ns 0.1440
NT vs. 90 Day -0.6131 -1.300 to 0.07407 No ns 0.0717

89

Renin Angiotensin Peptide Ocular Metabolism
Collected ocular fluids were analyzed for RAS peptides levels over the time after laser treatment.
To evaluate the acute phase of ocular healing that coincides with the observed histological findings, the
data analysis conducted was subset up to day 7.
A nonlinear fit of a local regression (LOWESS) was constructed using the concentrations measured
from the ocular samples. RAS peptide metabolic pathway is overviewed. After 50% R:GEN treatment, the
ocular Ang(1-10) or AngI levels increased immediately after treatment but began to decline after the first
day. In contrast, mice receiving 100% R:GEN treatment had a reduction of Ang(1-10). In the LOWESS
model, mice receiving 50% intensity had an inverse relationship when compare to animals receiving 100%
intensity as seen in Figure 3.20.
Since Ang(1-10) is a substrate for ACE1 to form AngII or Ang(1-8), we evaluate the ocular levels of
AngII over the same time period to determine whether changes in Ang(1-10) corresponds to formation of

Figure 3.19: Plasma levels of Ang(1-5) after R:GEN treatment over a 90 day time course

Dunnett's multiple comparisons test
Predicted (LS)
mean diff. 95.00% CI of diff. Significant? Summary
Adjusted P
Value
50%
NT vs. 0.04 Day -0.02580 -0.05727 to 0.005673 No ns 0.1054
NT vs. 0.16 Day -0.02238 -0.04712 to 0.002353 No ns 0.0738
NT vs. 3 Day -1.575 -4.178 to 1.028 No ns 0.2616
NT vs. 7 Day -0.02244 -0.07783 to 0.03295 No ns 0.5703
NT vs. 30 Day -0.3725 -0.7514 to 0.006390 No ns 0.0535
NT vs. 90 Day -0.3368 -0.5762 to -0.09732 Yes * 0.0110
100%
NT vs. 0.04 Day 0.005946 -0.01589 to 0.02778 No ns 0.8378
NT vs. 0.16 Day -0.004502 -0.04775 to 0.03875 No ns 0.9926
NT vs. 3 Day -1.208 -4.206 to 1.790 No ns 0.5779
NT vs. 7 Day -0.04466 -0.1052 to 0.01587 No ns 0.1496
NT vs. 30 Day -0.5141 -1.068 to 0.03926 No ns 0.0668
NT vs. 90 Day -0.3132 -0.6571 to 0.03077 No ns 0.0673

90
AngII. Ocular AngII levels for 50% and 100% intensity was proportionally increased which peaked on day
3 and declined thereafter.
To determine whether the formation of AngII correlated with expression of ACE1, which is an
important enzyme catalyzing this metabolism. We have confirmed that only minimal changes in gene
expression of ACE1 were found in retinal cells (data not shown). The expression of RPE ACE1 is
summarized in Figure 3.20B. ACE1 expression increased in animals receiving 50% intensity and remained
elevated. While animals receiving 100% intensity showed significant upregulation of ACE1 at 0.04 days
after R:GEN treatment, subsequent time points indicate downregulation. The gene expression findings do
not appear to correspond with the increased ocular concentration of AngII beyond 0.04 days.

Figure 3.20: Ocular metabolism of Ang(1-10) to AngII via ACE1
A(1-10)
Ang I
A(1-9)
A(1-7)
A(1-8)
AngII
↑ ACE
ACE2
ACE
ACE2
AT
1
AT
2 Mas

91
Although Ang(1-10) is a primary substrate for ACE1, it has also been found to be a substrate for
ACE2, metabolizing Ang(1-10) to Ang(1-9). To determine if R:GEN treatment will have an effect how
Ang(1-10) gets metabolized, ocular levels of Ang(1-9) in relations to Ang(1-10) were also evaluated, as
seen in Figure 3.21. Ocular Ang(1-9) increased proportionally with the level of R:GEN treatment intensity,
where maximal concentration was estimated to occur at approximately day 5. ACE2 gene expression in
the RPE was evaluated over the same time horizon. Overall, expression of ACE2 was seen to be
downregulated after R:GEN treatment during the acute phase.

Figure 3.21: Metabolism of Ang(1-10) to Ang(1-9) via ACE2
A(1-10)
Ang I
A(1-9)
A(1-7)
A(1-8)
AngII
ACE
ACE2
ACE
↓ACE2
AT
1
AT
2 Mas

92
Since ACE2 also catalyzes the conversion of AngII to Ang(1-7), we evaluated changes of ocular
Ang(1-7) in relations to these two parameters, as shown in Figure 3.22. As mentioned above, ACE2 gene
expression was found to be downregulated after R:GEN treatment and remained lower through the entire
7-day time horizon. When ocular levels of Ang(1-7) were evaluated, mice receiving 50% intensity show
an initial decrease where levels increased, peaking around Day 5. In contrast, mice receiving R:GEN laser
treatment at 100% intensity increased Ang(1-7) at 0.04 and 0.16 days, but declined thereafter.

Figure 3.22: Metabolism of AngII to Ang(1-7) via ACE2
A(1-10)
Ang I
A(1-9)
A(1-7)
A(1-8)
AngII
ACE
ACE2
ACE
↓ACE2
AT
1
AT
2 Mas

93
Another source for Ang(1-7) is the metabolism of Ang(1-9) to Ang(1-7) via ACE1 metabolism, as
shown in Figure 3.23. Ocular levels of Ang(1-9) are elevated above untreated levels even on Day 7. Gene
expression of ACE1 remained significantly elevated for mice treated with 50% intensity, whereas mice
receiving 100% intensity had elevated expression of ACE1 at 0.04 days but decreased expression until day
7. In contrast, the ocular levels of Ang(1-7) increased immediately and dropped by Day 3 for animals
receiving 100% intensity. In contrast, ocular levels of Ang(1-7) for mice receiving 50% R:GEN dropped
initially, and then began to increase by Day 3. Ocular Ang(1-7) increased initially corresponding to a drop
in Ang(1-9), suggesting that Ang(1-7) levels may come through this pathway when treating with R:GEN at
50% intensity.

Figure 3.23: Metabolism of Ang(1-9) to Ang(1-7) via ACE1
A(1-10)
Ang I
A(1-9)
A(1-7)
A(1-8)
AngII
↑ ACE
ACE2
ACE
ACE2
AT
1
AT
2 Mas

94
In addition to RAS peptide metabolism, the potential binding of each active peptide to its
respective receptor was explored in relationship to the peptide fluctuations observed in the ocular fluids
of treated animals. AT1R expression was elevated after laser treatment at 50% and 100% intensity. At
0.04 and 0.16 days after laser treatment, the expression for AT1R was above a 2-fold increase from
baseline for both 50% and 100% intensity, a clinically relevant escalation as compared to untreated
animals. Animals treated with 50% intensity had a sustained upregulation of AT1R up to day 7, while
animals in the 100% intensity arm had AT1R expression return to baseline by day 3.
Too probe the counter-balancing of the pathologic arm of RAS, gene expression of AT2R was also
evaluated for both animals treated with 50% and 100% intensity. It was found that gene expression of
AT2R was significantly upregulated in both 50% and 100% treatment groups and sustained through day 7,
as seen in Figure 3.24. When examining the relationship between ocular AngII levels with AT1R and AT2R
expression, the findings would suggest that AngII will likely bind to AT2R given the significant and
sustained gene expression. In fact, the expression of AT2R would indicate a greater potential for activating
the pro-regenerative arm of RAS.

Figure 3.24: Binding of AngII to AT1R and AT2R
A(1-10)
Ang I
A(1-9)
A(1-7)
A(1-8)
AngII
↑ ACE
↑ ACE2
↑ ACE
↑ ACE2
AT 1
AT 2 Mas

95
Ang(1-7) is the endogenous ligand for MasR, for which the binding to this receptor would also
activate the pro-regenerative arm of RAS. While gene expression of MasR was significantly upregulated
at 0.04 days using the 100% intensity treatment, expression was quickly downregulated for the remained
of the acute wound repair phase. However, in the 50% treatment group, gene expression of MasR was
significantly increased and remained elevated up to day 7, as seen in Figure 3.25.
Upregulation for MasR appear to correspond with time points where ocular Ang(1-7) levels
increase, suggesting a positively correlated relationship between the peptide and its receptor.
Ang(1-9) was recently shown to be a substrate that bind onto AT2R, and gene expression for the
receptor was also compared in relations to its ocular concentrations, as shown in Figure 3.26. AT2R gene

Figure 3.25: Binding of Ang(1-7) to MasR
A(1-10)
Ang I
A(1-9)
A(1-7)
A(1-8)
AngII
↑ ACE
↑ ACE2
↑ ACE
↑ ACE2
AT 1
AT 2 Mas

Figure 3.26: Binding of Ang(1-9) to AT2R
A(1-10)
Ang I
A(1-9)
A(1-7)
A(1-8)
AngII
↑ ACE
↑ ACE2
↑ ACE
↑ ACE2
AT 1
AT 2 Mas

96
expression was significantly upregulated when treated with both 50% and 100% intensities, where levels
remained elevated up to day 7. Similarly, ocular levels of Ang(1-9) were found to increase with both 50%
and 100% intensities, and also remained elevated until day 7.
Correlations and Statistical Analyses of Gene Expression and RAS Metabolomics
The quantified RAS concentrations were then converted to fold change relative to untreated
control mice such that the observed changes in plasma and ocular fluid could be correlated as well as
compared the RT-PCR results, as shown in Figure 3.27. A two-tailed Spearman correlation analysis was
conducted with a 95% confidence level between the plasma and ocular levels to determine whether the
changes within the ocular space can be tracked by the levels in the eye.

97
In contrast to the CXCR5 -/- mice treated at 100% intensity, it was found that a negative
correlation of ocular and plasma fold changes of A(1-12) and A(1-10) were statistically significant in mice
treated with 50% intensity, r = -0.928 (p = 0.0067) and r = 0.8929 (p = 0.0123), respectively. The full
analysis of correlations is shown below in Table 3.4 below.

Figure 3.27 Aggregate fold change in RAS peptides of 100% intensity treatment between plasma and ocular
matrices for correlation analysis.
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.1
1
10
100
Plasma_OcularA(1-12)
Time (Days)
Fold Change from Untreated
Ocular
Plasma
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.001
0.01
0.1
1
10
Time (Days)
Fold Change from Untreated
Plasma_OcularAngII
Ocular
Plasma
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.1
1
10
Plasma_OcularA(2-8)
Time (Days)
Fold Change from Untreated
Ocular
Plasma
7
8
6
8
8
8
7
13
12
13
11
13
12
13
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.1
1
10
100
Time (Days)
Fold Change from Untreated
Plasma_OcularA(1-10)
Ocular
Plasma
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.1
1
10
100
1000
10000
Plasma_OcularA(1-7)
Time (Days)
Fold Change from Untreated
Ocular
Plasma
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.1
1
10
Plasma_OcularA(3-8)
Time (Days)
Fold Change from Untreated
Ocular
Plasma
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.1
1
10
Plasma_OcularA(1-9)
Time (Days)
Fold Change from Untreated
Ocular
Plasma
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.1
1
10
100
Plasma_OcularA(1-5)
Time (Days)
Fold Change from Untreated
Ocular
Plasma
0.00 0.04 0.16 3.00 7.00 30.00 90.00
0.1
1
10
100
Plasma_OcularA(3-7)
Time (Days)
Fold Change from Untreated
Ocular
Plasma
Peptide A(1-12) A(1-10) A(1-9) Ang II A(1-7) A(1-5)' A(2-8) A(3-8) A(3-7)'
Spearman r -0.9286 0.8929 -0.2857 -0.5 -0.197 -0.4286 0.5 0.4286 -0.1786
P (2-tailed) 0.0067 0.0123 0.556 0.2667 0.6857 0.3536 0.2667 0.3536 0.7131
Number 7 7 7 7 7 7 7 7 7
Table 3.4: Correlation analysis between plasma and ocular RAS peptides using 50% R:GEN intensity

98
Given the observed sustained increases in the relative gene expression of AT2R and MasR in the
RPE, it would be expected that a shift towards the regenerative arm occurred with R:GEN treatment using
both 50% and 100% intensities. The correlation of gene expression of the RAS enzymes, receptors, and
ocular peptide levels are summarized in Figure 3.28 and Figure 3.29 below. It appears that the peptide
ratios, which would be indicative of enzymatic cleavage, did not have a positive correlation with gene
expression of ACE1 and ACE2 in RPE. However, the ratio of AngII/AngI and gene expression of ACE1 were
found to be negatively correlated and statistically significant (p = 0.03).

Correlation of Ocular Fluid & RPE Enzyme
Enzyme RPE - ACE1 RPE - ACE2
Peptide A(1-8)/A(1-10) A(1-7)/A(1-9) A(1-9)/A(1-10) A(1-7)/A(1-8)
Spearman r -0.8214 -0.6786 -0.3571 -0.5714
P (two-tailed) 0.0341 0.1095 0.4444 0.2000
Number of XY Pairs 7 7 7 7
Figure 3.28: Correlation between Ocular RAS peptide ratios and gene expression their respective enzymes in
RPE

99
None of the correlations between ocular RAS peptides and their respective receptors were found
to be statistically significant. However, it appears that AT1R and AT2R expression is induced in the RPE
post-laser treatment, while MasR is immediately upregulated at 0.04 days but downregulated until day
30.

Dose Response and Area Under the Curve of Ocular RAS
To determine if there was a dose-dependent response from increased R:GEN laser intensity, area
under the curve (AUC) was calculated for each peptide measured within the ocular space. Concentrations
were plotted across each timepoint, and AUC was calculated using the trapezoid rule (H. J. Motulsky). All
peptides, except for Ang(1-7) and Ang(1-5) were found to increase at 100% intensity over 50% intensity
in the acute phase of healing, although the standard error for the peptides were found to overlap. Peptide

Correlation of Ocular Fluid & RPE Receptor
RPE - AT1R RPE - AT2R RPE - MasR
AngII AngII A(1-9) A(1-7)
Spearman r 0.07143 -0.2143 -0.03571 -0.4865
P (two-tailed) 0.9063 0.6615 0.9635 0.2730
Number of XY Pairs 7 7 7 7
Figure 3.29: Correlation between ocular RAS peptides and RPE gene expression their respective receptors in
RPE

100
ratios were assessed using AUCs, where 50% intensity appears to have greater enzymatic activity than
100%, except for the Ang(1-9)/Ang(1-10) ratio.

3.4 Discussion
In this chapter, we verified the safety and efficacy of the Lutronic R:GEN laser in a soft drusen
mouse model of AMD with a dose-escalation and time course design. Like the hard drusen study,
images, histology, gene expression, and metabolomics techniques were utilized to provide insight into
the effect of SRT on eyes afflicted with soft drusen. In the previous chapter, we proved that R:GEN was
safe and regressed drusen in a hard drusen animal model. We continue to characterize the effects of
SRT on ocular RAS and build upon our previous work by looking to identify a dose-dependent response
with R:GEN treatment.
Area Under the Curve (ng*hr/mL ± Std Error)
Laser Intensity

50% 100%
Ang(1-10)
23.65 ± 6.567 45.18 ± 44.13
Ang(1-9)
13.43 ± 2.129 33.43 ± 22.18
Ang II
20.62 ± 19.26 36.38 ± 24.83
Ang(1-7)
51.10 ± 116.2 25.42 ± 32.47
Ang(2-8)
15.38 ± 3.455 38.58 ± 31.45
Ang(3-8)
20.70 ± 3.579 31.68 ± 16.45
Ang(3-7)
0.9689 ± 1.176 5.603 ± 8.938
Ang(1-5)
60.93 ± 24.70 53.91 ± 26.81

RAS Peptide Ratios
50% 100%
ACE
Ang(1-8)/Ang(1-10) 0.871 0.805
Ang(1-7)/Ang(1-9) 3.804 0.760
ACE2
Ang(1-7)/Ang(1-8) 2.478 0.698
Ang(1-9)/Ang(1-10) 0.567 0.739
Table 3.5: Area Under the Curve for ocular RAS peptides. Peptide ratios were assessed to anticipate enzymatic
activity of ACE and ACE2

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In this study, through a combination of RT-PCR, histology, and fundus imaging, we showed that
R:GEN is also safe in our CxCr5 -/- soft drusen animal model. Similar to the study reported in chapter 2,
H&E cross-sections of collected eyes showed that R:GEN selectively targeted the RPE while leaving the
photoreceptors undamaged. However, due to the lack of a distinct border, we were unable to use
Fundus Autofluorescence (FAF) to measure drusen deposits, a limitation not anticipated given the
findings from the hard drusen mouse model.
While the previous chapter discussed the application of R:GEN in a hard drusen mouse model
(Cx3Cr1 -/-), the presence of soft drusen is a greater risk factor for the development and progression of
dAMD (Klein et al., 2015). Therefore, the results of this study in in the CxCr5 -/- model may be more
instrumental in translating the benefit of R:GEN laser from rodent to humans.
3.4.1 R:GEN upregulates ECM turnover in soft drusen (CxCr5 -/-) mouse model
Like in chapter 2, we confirm the upregulation of metalloproteinases (MMPs) after R:GEN
treatment, as previously reported in an SRT study by Jobling et al (Jobling et al., 2015). However, unlike
the gene expression of MMPs in the Cx3Cr1-knockout model, the gene expression of MMPs are
upregulated significantly and sustained for the entire 90 days of the study post-R:GEN treatment. These
findings would suggest that the wound healing response in RPE of CxCr5-knockouts might initiate both
wound contraction and cell migration (Caley et al., 2015; Martins, Caley, & O'Toole, 2013). Because it
was previously reported that MMP activity was decreased in aging eyes, leading to the conclusion that
impaired ECM turnover was linked to the development of AMD, our results indicate R:GEN might be
beneficial in reactivating this pathway (Guo, Hussain, Limb, & Marshall, 1999). While not conducted for
this study, it may be beneficial to compare the gene expression levels of our treated CxCr5-knockout
mice against those of healthy C57/Bl mice to verify if R:GEN treatment could induce MMP changes to
emulate that of healthy normal mice.

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3.4.2 R:GEN shifts ocular RAS towards pro-regeneration in soft drusen (CxCr5 -/-) mouse model
As discussed in previous chapters, the Renin Angiotensin System (RAS) plays a role in promoting
either fibrotic or non-fibrotic wound healing (Jadhav et al., 2013). In chapter 2, R:GEN was reported to
stimulate RAS, immediately increasing ACE2and MasR after treatment, while increasing the ocular
production of Ang(1-9) and Ang(1-7); two prominent peptides of the protective arm of RAS. As previously
mentioned, interest in the RAS pathway has increased as the body of evidence regarding its role in fibrosis
and wound healing continues to increase (AlQudah, Hale, & Czubryt, 2020). Homeostasis is typically
maintained by balancing the pathogenic AngII/AT1R arm with the protective/regenerative AngII/AT2R and
Ang(1-7)/MasR arms of RAS (Xie, Hu, Cheng, Liu, & Wei, 2022). The data in this study suggests a shunting
towards the protective arm after R:GEN treatment, given the increased expression of peptides and
receptors in this axis as opposed to those of the pathogenic arm.
In this study, we found additional evidence that the protective arm of RAS was activated with
R:GEN, where ocular Ang(1-9) and Ang(1-7) were again upregulated from treatment. AT1R and AT2R were
also found to have persistent upregulation, although relative gene expression of AT2R was greater,
suggesting a preference for the protective arm. Additionally, gene expression of ACE at 50% intensity was
upregulated during the acute phase while ACE was downregulated at 100% for the same period.
Surprisingly, gene expression of ACE2 was downregulated during the acute phase, counter to what was
expected given the increased production of Ang(1-7) and the literature regarding RAS metabolism (R. A.
S. Santos et al., 2019).
To characterize how SRT would modulate RAS and whether plasma levels of angiotensin peptides
could be representative of ocular activity, a spearman correlation was conducted between each peptide
in plasma, ocular fluid. It was observed that the relationship between plasma and ocular levels of Ang(1-
12) and Ang(1-10) were identified as statistically significant. Plasma and ocular Ang(1-12) levels were
found to have a negative relationship. Surprisingly, plasma and ocular Ang(1-10) levels were positively

103
correlated in contrast to the negative relationship found in chapter 2, although ocular levels were found
to increase in both animal models. From this data, it is believed that the increase in ocular Ang(1-10) is to
provide more substrate for the angiotensin converting enzymes, primarily ACE according to the RT-PCR
results, to generate the active metabolites downstream. However, because plasma Ang(1-10) levels
between hard and soft drusen models appear to be contradictory, additional research may be required to
identify the reasoning behind this discrepancy, especially when translating into human trials. Additional
confounders or patient attributes may exist in determining the feasibility of using systemic Ang(1-10) to
determine wound repair after R:GEN treatment. However, while the plasma/ocular relationships for AngII
was not found to be statistically significant, it was found to be negatively correlated across both animal,
suggesting that the observed decrease in plasma levels would indicate an increase in ocular levels.
Again using our metabolomic approach, we explored the effect that R:GEN had on the expression
and activity of RAS enzymes in comparison to our findings with regards to gene expression. The ratios of
RAS peptides, calculated as product/substrate, indicated negligible changes in ocular ACE1 but
significantly upregulated activity of ACE2. This increased ACE2 activity would suggest greater production
of Ang(1-9) and Ang(1-7), both of which are prominent peptides in the protective arm of RAS. This
approach runs counter to what was determined in RT-PCR, although is more likely to be accurate when
compared to gene expression. Again, one possible explanation for this discrepancy might be the release
of soluble ACE1 and ACE2 into the vitreous humor after laser induction, which might continue to convert
RAS peptides even though gene expression was found to be downregulated. However, due to the
limitations regarding volume of a mouse eye, the presence and quantitation of soluble enzymes could not
be performed. The second explanation for the data is a temporal lag between a significant upregulation
in ACE1 expression at 0.04 days, followed by a significant upregulation in AT1R and AT2R, such that
generated peptides are quickly bound to their receptors. As such, the correlation analysis between gene

104
expression and peptide metabolomics were all found to have negative relationships, with ACE1 and
AngII/AngI being statistically significant (p = 0.03).
While the metabolomics of RAS peptides suggested a shift towards the production of peptides
known to be pro-regenerative, the actual activity is dependent on binding to their respective receptors.
Correlation analysis was again conducted to determine if any relationships between the angiotensin
peptides and their respective receptors exist, for which changes in peptide levels were correlated with
changes in gene expression of their respective receptors. While no correlations were found to be
statistically significant, it was observed AT2R expression was much greater than that of AT1R expression,
once again suggesting a potential shift towards the pro-regenerative arm of RAS if peptides bind to both
receptor subtypes equally.
Comparing the systemic and ocular levels of each RAS peptide, we observe that the level of each
peptide varies in its response to tissue injury and repair. However, it appears that both Ang(1-10) and
Ang(1-9) remain relatively stable in the ocular fluid, except for a drastic decrease in Ang(1-10) in the first
hour.
In this study, mice were treated with 50% and 100% laser intensities to assess if any response to
a “dose increase” would exist. As expected, the higher intensity treatment group stimulated greater
production of RAS peptides. Unexpectedly, area under the curve (AUC) for Ang(1-7) and Ang(1-5) were
higher in 50% treatment groups and lower in 100% treatment groups. Furthermore, while peptide
production was greater at 100%, the peptide ratios were found to be greater at 50%, except for Ang(1-
7)/Ang(1-9), suggesting greater metabolism at the lower treatment intensity. These findings suggest that
activation of the protective arm of RAS occurs at the 50% intensity, where ACE2 and Ang(1-7) levels were
higher than that of 100%. These results suggest that a dose-dependent response to laser treatment does
exist in an ocular context, despite the relatively small difference in treatment energies as evidenced from

105
Figure 3.1. Furthermore, future iterations of R:GEN-related studies may require multiple doses to optimize
for specific peptide production, such as the tradeoff between Ang(1-7) and Ang(1-9).
3.5 Conclusion
In this chapter, it was reported that dose escalation of R:GEN from 50% intensity to 100%
intensity was well-tolerated in CxCr5 -/- mice and upregulated mechanisms related to extra cellular
matrix remodeling and wound healing. Analyses of all collected images for each time point and laser
intensity indicate a diseased eye is capable of recovering from treatment. Gene expression for RAS
enzymes were unexpectedly downregulated in RPE, given the increase in downstream RAS peptides in
collected ocular fluids. The negative correlation between plasma and ocular levels of Ang(1-12) and
Ang(1-10) during the acute phase of wound healing highlight the potential for a blood-based biomarker
capable of elucidating reparative activity in the eye. Additionally, the AUC of ocular angiotensin peptides
suggest that a dose-dependent response from laser treatment does exist, creating an opportunity to
further refine this therapeutic modality for specific sub-populations of AMD patients.

106
3.6 Chapter 3 References
AlQudah, M., Hale, T. M., & Czubryt, M. P. (2020). Targeting the renin-angiotensin-aldosterone system in
fibrosis. Matrix Biol, 91-92, 92-108. doi:10.1016/j.matbio.2020.04.005

Caley, M. P., Martins, V. L., & O'Toole, E. A. (2015). Metalloproteinases and Wound Healing. Adv Wound
Care (New Rochelle), 4(4), 225-234. doi:10.1089/wound.2014.0581

Caligiuri, G., Nicoletti, A., Poirier, B., & Hansson, G. K. (2002). Protective immunity against
atherosclerosis carried by B cells of hypercholesterolemic mice. The Journal of clinical
investigation, 109(6), 745-753. doi:10.1172/JCI7272

Crotty, S. (2011). Follicular Helper CD4 T Cells (TFH). Annual Review of Immunology, 29(1), 621-663.
doi:10.1146/annurev-immunol-031210-101400

Frederick, P. A., & Kleinman, M. E. (2014). The Immune System and AMD. Curr Ophthalmol Rep, 2(1), 14-
19. doi:10.1007/s40135-013-0037-x

Gunn, M. D., Ngo, V. N., Ansel, K. M., Ekland, E. H., Cyster, J. G., & Williams, L. T. (1998). A B-cell-homing
chemokine made in lymphoid follicles activates Burkitt's lymphoma receptor-1. Nature,
391(6669), 799-803. doi:10.1038/35876

Guo, L., Hussain, A. A., Limb, G. A., & Marshall, J. (1999). Age-dependent variation in metalloproteinase
activity of isolated human Bruch's membrane and choroid. Invest Ophthalmol Vis Sci, 40(11),
2676-2682. Retrieved from https://www.ncbi.nlm.nih.gov/pubmed/10509665

H. J. Motulsky. GraphPad Statistics Guide. Retrieved from
http://www.graphpad.com/guides/prism/7/statistics/index.htm

Hampe, C. S. (2012). B Cell in Autoimmune Diseases. Scientifica, 2012, 215308.
doi:10.6064/2012/215308

Hector, S. M., & Sørensen, T. L. (2017). Circulating monocytes and B-lymphocytes in neovascular age-
related macular degeneration. Clinical ophthalmology (Auckland, N.Z.), 11, 179-184.
doi:10.2147/OPTH.S121332

Huang, H., Liu, Y., Wang, L., & Li, W. (2017). Age-related macular degeneration phenotypes are
associated with increased tumor necrosis-alpha and subretinal immune cells in aged Cxcr5
knockout mice. PLoS One, 12(3), e0173716. doi:10.1371/journal.pone.0173716

Jadhav, S. S., Sharma, N., Meeks, C. J., Mordwinkin, N. M., Espinoza, T. B., Roda, N. R., . . . Rodgers, K. E.
(2013). Effects of combined radiation and burn injury on the renin-angiotensin system. Wound
Repair Regen, 21(1), 131-140. doi:10.1111/j.1524-475X.2012.00867.x

Jobling, A. I., Guymer, R. H., Vessey, K. A., Greferath, U., Mills, S. A., Brassington, K. H., . . . Fletcher, E. L.
(2015). Nanosecond laser therapy reverses pathologic and molecular changes in age-related
macular degeneration without retinal damage. FASEB J, 29(2), 696-710. doi:10.1096/fj.14-
262444

107
Klein, R., Myers, C. E., Lee, K. E., Gangnon, R. E., Sivakumaran, T. A., Iyengar, S. K., & Klein, B. E. (2015).
Small Drusen and Age-Related Macular Degeneration: The Beaver Dam Eye Study. J Clin Med,
4(3), 424-440. doi:10.3390/jcm4030425

Küppers, R. (2005). Mechanisms of B-cell lymphoma pathogenesis. Nature Reviews Cancer, 5(4), 251-
262. doi:10.1038/nrc1589

Lennikov, A., Mukwaya, A., Saddala, M. S., & Huang, H. (2021). Deficiency of C-X-C chemokine receptor
type 5 (CXCR5) gene causes dysfunction of retinal pigment epithelium cells. Lab Invest, 101(2),
228-244. doi:10.1038/s41374-020-00491-4

Major, A. S., Fazio, S., & Linton, M. F. (2002). B-Lymphocyte Deficiency Increases Atherosclerosis in LDL
Receptor–Null Mice. Arteriosclerosis, Thrombosis, and Vascular Biology, 22(11), 1892-1898.
doi:10.1161/01.ATV.0000039169.47943.EE

Martins, V. L., Caley, M., & O'Toole, E. A. (2013). Matrix metalloproteinases and epidermal wound
repair. Cell Tissue Res, 351(2), 255-268. doi:10.1007/s00441-012-1410-z

Nagai, H., & Kalnins, V. I. (1996). Normally occurring loss of single cells and repair of resulting defects in
retinal pigment epithelium in situ. Exp Eye Res, 62(1), 55-61. doi:10.1006/exer.1996.0007

Patel, N., Ohbayashi, M., Nugent, A. K., Ramchand, K., Toda, M., Chau, K.-Y., . . . Chong, V. (2005).
Circulating anti-retinal antibodies as immune markers in age-related macular degeneration.
Immunology, 115(3), 422-430. doi:10.1111/j.1365-2567.2005.02173.x

Santos, R. A. S., Oudit, G. Y., Verano-Braga, T., Canta, G., Steckelings, U. M., & Bader, M. (2019). The
renin-angiotensin system: going beyond the classical paradigms. Am J Physiol Heart Circ Physiol,
316(5), H958-H970. doi:10.1152/ajpheart.00723.2018

Xie, J. X., Hu, J., Cheng, J., Liu, C., & Wei, X. (2022). The function of the ACE2/Ang(1-7)/Mas receptor axis
of the renin-angiotensin system in myocardial ischemia reperfusion injury. Eur Rev Med
Pharmacol Sci, 26(6), 1852-1859. doi:10.26355/eurrev_202203_28330

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Chapter 4: Cell-Based Assays for Characterization of Ocular Damage Model
4.1 Introduction
In previous chapters, it was shown that laser induced retinal pigmented epithelial (RPE) injury
leads to changes in the Renin-Angiotensin System (RAS). Our in vivo data suggest that there is an increase
in the expression of RAS receptors in response to laser-induced RPE injuries. To further probe the role of
RAS in retinal wound healing, we evaluated the impact of AngII and Ang(1-7) biosynthesis with respective
expression of receptors in human RPE cell line, ARPE-19. ARPE-19 is a commonly used immortalized RPE
cell line isolated from a 19-year-old male donor (Dunn, Aotaki-Keen Ae Fau - Putkey, Putkey Fr Fau -
Hjelmeland, & Hjelmeland). Like normal RPE, ARPE-19 can pigment and form cobblestone structures in
vitro.
However, limitations with the accessible laser equipment exist with translating the prior in vivo
design for in vitro cells assays. To bridge this gap, it was critical to identify appropriate stimuli that could
induce similar biochemical responses. Photobiomodulation (PBM), a field of research that uses low energy
light to stimulate biological systems, has previously demonstrated upregulated mitochondrial activity and
increased production of reactive oxygen species (Amaroli et al., 2021). Previous studies in the effects of
Q-switched Nd:YAG nanosecond pulse lasers have also shown higher levels of oxidative damage and
localized inflammation, leading to increased lipid peroxidation (Bergandi et al., 2018). Well-established as
part of the innate immune response, acute rapid production of hydrogen peroxide (H 2O 2) is known to
occur after damage to epithelium (van der Vliet & Janssen-Heininger, 2014). Therefore, it was surmised
that induction of H 2O 2 would emulate the response to laser treatment as observed by the increase in
reactive oxygen species. Additionally other groups have previously studied the effects of oxidative stress
in the eye by using chronic H 2O 2 induced damage on ARPE-19, demonstrating the utility of this particular
in vitro model (Kaczara, Sarna, & Burke, 2010).

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In this chapter, we evaluated the biological response after H 2O 2 induction to understand the
overall RPE response to tissue injury. Using various enzymatic inhibitors (e.g., enalapril, MLN4760) and
receptor inhibitors (e.g., Losartan, PD123319, and A779) of RAS, we probe the molecular events that are
associated with RAS-mediated responses in RPE. Enalapril is a potent ACE inhibitor, while MLN4760 is
specific inhibitor of ACE2. In this study, losartan, an inhibitor for AT1R, was used to determine the role of
AT1R activation after H 2O 2-mediated injury. We used PD123319, an inhibitor for AT2R, to probe the role
of AT2R after H 2O 2-mediated injury and whether its inhibition may alter the molecular response. Key to
all cellular response to tissue injury is Mas receptor activation. To further understand the impact of MasR,
this chapter will use A779, a Mas receptor antagonist, that is capable of blocking Ang(1-7) mediated
effects. To ensure that compensatory upregulation is blocked, the concentration of these inhibitors used
will be at least 10-fold IC 50, or at concentrations capable of exhibiting complete inhibition of enzymes and
receptors (Swinney, 2011). We will probe the effect that RAS inhibition has on extracellular levels of RAS
peptides, mimicking the immediate release into the vitreous post-injury. The RAS peptide components
will be quantified using a multiplexed RAS LC-MS metabolomic assay, that will be correlated with gene
expression of the targeted RAS components, setting the stage for future exploration RAS peptide
dynamics in the ocular space using in vitro models.
4.2 Materials and Methods
4.2.1 Cells
ARPE-19 cells were purchased from American Type Culture Collection (ATCC, Manassas, VA and
were maintained in culture media comprised of DMEM/F-12 (Corning Life Sciences, Corning, NY), 1% fetal
bovine serum, and 1% Antibiotic-Antimycotic (Gibco, Carlsbad, CA). Cultured cells were maintained in 24-
well plates (VWR, Radnor, PA) using a humidified incubator set to 37
o
C and 5% CO 2.

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4.2.2 Angiotensin II Dose Escalation
Angiotensin II (AngII) induction has been implicated in causing cellular damage, confluent ARPE-
19 cells were treated with AngII in a concentration escalation design from 1 nM to 1 µM. Cells were
treated for 24 and 48 hours, at which time conditioned media were collected. The cells were then
treated with 500 µL of RNAzol RT was added to each well. Cells were pipetted repeatedly to create a
homogenous lysate, which was then transferred to a clean 1.5 mL microcentrifuge tube and stored at -
20
o
C.
4.2.3 Renin-Angiotensin Inhibition and Stress Induction
Cells were incubated with RAS inhibitors, shown in Table 4.1, for 30 minutes followed by hydrogen
peroxide (H 2O 2) treatment in a dose escalation manner where the concentration was 0.5 to 2.0 mM. At
the end of each time point, cell media was collected for LC-MS/MS analysis and stored 1.5 mL
microcentrifuge tubes containing 20 µL of prepared protease inhibitor as previously described in Chapter
2. After aspiration, 500 µL of RNAzol RT (Sigma-Aldrich, St. Louis, MO) was added to each well and rocked
on a plate rocker for 5 minutes. Cells were then scraped using a Biotium Mini Cell Scraper (Fisher Scientific,
Hampton, NH). Cell lysate was collected and transferred to a clean 1.5 mL microcentrifuge tube. Tubes
Inhibitor
Name
Target IC 50 Source
Enalapril ACE1 1.2 nM (Gonzalez Amaya, Cabrera, Matallana, Arevalo, & Guevara-Pulido, 2020;
Gross et al., 1981)
MLN-4760 ACE2 0.5 µM (Joshi, Balasubramanian, Vasam, & Jarajapu, 2016)
Losartan AT1R 20 nM (Burnier, 2001)
PD123319 AT2R 10 nM Boulay, Servant G Fau - Luong, Luong Tt Fau - Escher, Escher E Fau -
Guillemette, and Guillemette
A779 MasR 0.3 nM (R. A. Santos et al., 2003)
Table 4.1: List of RAS inhibitors and respective IC 50’s
Cell Line Inhibitor Concentration Time Points (Days) H 2O 2 (mM) Analysis
ARPE-19
Enalapril 1 µM 0.04 0, 0.5, 1, 2 RT-PCR, LC-MS/MS
Losartan 1 µM
PD123319 1 µM
A779 1 µM
MLN4760 10 µM
Table 4.2: Study Design for mechanistic dissection of RAS pathway and impact of tissue injury

111
containing RNAzol RT were frozen immediately at -20
o
C. The overall study design is summarized in the
Table 4.2.
4.2.4 RT-PCR Analysis
4.2.4.1 ARPE-19 RNA Extraction and cDNA Synthesis
RNA was extracted according to manufacturer’s protocol and concentrations were adjusted to 10
ng/µL using nuclease-free water. cDNA was synthesized using the extracted ARPE-19 RNA using the
RevertAid RT Reverse Transcription kit (Thermo Fisher Scientific, Wilmington, DE) according to the
manufacturer’s protocol. Samples were placed in a Bio-Rad Thermal Cycler (Bio-Rad, Hercules, CA), where
they were incubated for 5 min at 25 °C followed by 60 min at 42 °C, and finally terminated by heating at
70 °C for 5 min.
4.2.4.2 cDNA Pre-amplification
The 20 µL of prepared cDNA was pre-amplified by adding 25 µL SsoAdvanced PreAmp Supermix
(Bio-Rad, Hercules, CA) and 5 µL of a prepared primer assay pool, for a final volume of 50 µL. The primer
assay pool included primer stocks of all genes that would be analyzed for RT-PCR, such that the final
preamplification reaction contains 50 nM of each primer, summarized in the table below. The cDNA
reaction mix was set for 12 cycles, for which the first stage was 3 minutes at 95°C for polymerase
activation, followed by 15 seconds for denaturation, and 4 minutes at 58°C for annealing/extension.
4.2.4.3 RT-PCR for Targeted Genes
Genes of the Renin-Angiotensin System were probed along with genes downstream of peptide-
receptor binding. The RT-PCR master mix was prepared by mixing Power Up SYBR Green master-mix
(Applied Biosystems, Foster City, CA) and the forward and reverse primers for each gene. The primers
sequences used were selected from PrimerBank (Harvard, Boston, MA) and are listed in Table 4.3.

112
Prepared master mix and diluted cDNA were pipetted into a 384-well plate (VWR, Radnor, PA) by
the Integra Assist Plus (Integra Biosciences, Hudson, NH). Samples were loaded in triplicates, plates were
sealed with an adhesive plate sealer, and centrifuged at 1000 g for 1 minute. The run method consisted
of an initial hold for 2 minutes at 50°C, followed by 95°C for 2 minutes. The amplification stage was set for
40 cycles, with a 1 second hold at 95°C and 30 seconds at 60°C for annealing/elongation. The final melt
curve stage was set for 15 seconds going from 1.6°C to 95°C, followed by 1 minute from 1.6°C to 60°C,
and finally 0.15°C to 95°C for 15 seconds. The data was collected and analyzed using Data Assist
(Invitrogen, Carlsbad, CA). Gene expression of MMP2, MMP3, MMP9, and TIMP2, TGF- β1 were assessed
to verify if H 2O 2 would initiate extracellular matrix remodeling, similar to R:GEN. ACE, ACE2, AT1R, AT2R,
and MasR were probed to verify the effect of H 2O 2 damage, as well as effect of combined inhibitor
treatment. NOX-2, SHP-1, and MCP-1 were probed as signals downstream of RAS receptors to verify
activation. Gene expression was assessed using delta-delta CT, normalized to β-Actin as the reference
gene and compared to untreated cells. Multiple comparisons tests were corrected using Dunnett‘s test,
and fold changes were plotted in GraphPad Prism (GraphPad, San Diego, CA).
Gene Forward (5’-3’) Reverse (5’-3’)
MMP2 ATACCATCGAGACCATGCG CCAATGATCCTGTATGTGATCTG
MMP3 GACTCCACTCACATTCTCC AAGTCTCCATGTTCTCTAACTG
MMP9 TGCAACGTGAACATCTTCG GAATCGCCAGTACTTCCCA
TIMP2 ATCTCATTGCAGGAAAGGC TCTGGTACCTGTGGTTCAG
ACE1 GGAGGAATATGACCGGACATCC TGGTTGGCTATTTGCATGTTCTT
ACE2 CGAAGCCGAAGACCTGTTCTA GGGCAAGTGTGGACTGTTCC
AT1R CAGCGTCAGTTTCAACCTGTACG GCAGGTGACTTTGGCTACAAGC
AT2R CCATGTTCTGACCTTCCTGGATG CGGATTAACGCAGCTGTTGGTG
MasR ATGGATGGGTCAAACGTGACA CGATGTGCATTCCCGACTG
ACTB CATGTACGTTGCTATCCAGGC CTCCTTAATGTCACGCACGAT
GAPDH GGAGCGAGATCCCTCCAAAAT GGCTGTTGTCATACTTCTCATGG
TGF- β1 GGCCAGATCCTGTCCAAGC GTGGGTTTCCACCATTAGCAC
NOX-2 AACGAATTGTACGTGGGCAGA GAGGGTTTCCAGCAAACTGAG
SHP-1 GGTGTCCACGGTAGCTTCC ACAGGTCATAGAAATCCCCTGAG
MCP-1 CAGCCAGATGCAATCAATGCC CAGATCCAGTCTTGCGGACAAG
Table 4.3: Primers for the specific gene targets

113
4.2.5 LC-MS/MS Analysis
4.2.5.1 Solid Phase Extraction of Collected Cell Media Samples
Frozen cell media containing protease inhibitors was thawed on ice, and from each sample, 150
µL was aliquoted into a clean 1.5 mL microcentrifuge tube. Each sample was then spiked with 150 µL of a
prepared 40 ng/mL NorLeu
3
-Angiotensin(1-7) (NLE) internal standard solution. Samples were then
vortexed thoroughly and placed back on ice. While samples were incubating, Oasis MAX solid phase
extraction columns (Waters, Milford, MA) which were pre-conditioned using 1 mL of HPLC grade methanol
(VWR, Radnor, PA), followed by 1 mL of HPLC grade water (VWR, Radnor, PA).
Prepared samples were alkalinized by adding 700 µL of a 5% ammonium hydroxide solution and
vortexed thoroughly. Dissociated samples (1000 µL) were then layered onto the primed solid phase
extraction (SPE) columns. Columns were then washed with 500 µL of 5% ammonium hydroxide, twice.
Columns were then washed with 500 µL of HPLC grade methanol and then dried for a couple minutes.
Samples were then eluted from the SPE columns with 500 µL of methanol with 2% formic acid, twice.
Eluted samples were then dried under inert nitrogen gas. The residue was then reconstituted using 50 µL
of 10 mM Ammonium Formate buffer, pH 4.2 and transferred to a clean HPLC vial (Sun-Sri, Rockwood,
TN), where 40 µL were injected into an API4000 (AB Sciex, Framingham, MA) for analysis.
4.2.5.2 Targeted LC-MS/MS Instrumentation and Conditions
RAS peptides were separated and quantified using an Agilent 1200 HPLC (Agilent, Santa Clara, CA)
connected to an API 4000 triple quad mass spectrometer (AB Sciex, Framingham, MA). Analyte data was
acquired using Analyst 1.6.2, where each analyte was quantified using scheduled multiple reaction
monitoring (MRM) identified during manual tuning of each compound. The MRM’s of the peptides
included in the method as previously summarized in Chapter 2. Acquired data was quantified using
MultiQuant 3.0 (AB Sciex, Framingham, MA).

114
Peptides were separated using an Agilent Poroshell 120 EC-C18 (4.6X100 mm, 2.7μm) column
(Agilent, Santa Clara, CA). Analytes were eluted using a gradient mobile phase consisting of two solvents:
1) Solvent A was HPLC grade water with 0.2% formic acid; and 2) Solvent B was HPLC grade methanol with
0.2% formic acid. The HPLC parameters and gradient program is described in Chapter 2 (Method Section:
HPLC parameters and gradient program [Table 2.6 – Table 2.8]).
4.2.6 Statistical Analysis
Results are expressed as mean ± SD unless specified otherwise. GraphPad Prism (GraphPad, San
Diego, CA), R (R Foundation for Statistical Computing, Vienna, Austria), and RStudio (Rstudio Inc, Boston,
MA) were used for data preparation, data cleaning, statistical analysis, and data visualization on
quantitative and qualitative data. Two-sided p-values less than 0.05 were considered statistically
significant. Statistically significant results are bolded and denoted with an asterisk (*).
4.3 Results
4.3.1 Angiotensin II Dose Escalation
4.3.1.1 Dose Dependent Response of RAS Enzymes and Receptors
ARPE-19 incubated in AngII in a dose escalation manner ranging from 1 nM to 1000 nM, where
the impact of RAS components was evaluated at 24 hours. The effect of AngII on ARPE-19 was used to
explore and confirm the gene expression found in the prior animal studies. RAS gene component
expression in response to AngII are summarized in Figure 4.1. ACE1 gene expression was upregulated at
100 nM and 1 µM after 24 hours of treatment.

115
AngII treatment of ARPE-19 showed that ACE2 expression was significantly increased at 1 nM
AngII after 24 hours of treatment. The expression of ACE2 expression was continued to be at similar
levels even with AngII concentration. However, ACE2 expression remained significantly upregulated
across all concentrations of AngII, except for 100nM and 1µM.
Gene expression of AT1R was found to be upregulated at 24 hours, even at 1 nM AngII. Gene
expression of AT2R was also found to be upregulated at 24 hours starting from 1 nM of Ang II treatment.
Similar to AT2R, MasR was found to increase at 24 hours, appearing to increase its expression up until
100 nM. Findings are summarized in Figure 4.2.

4.3.1.2 Ang II Induces Changes in Metalloproteinases
AngII is capable of inducing MMP activation by binding to AT1R, and gene expression of MMP’s
were probed in this cell model to compare against the results from the mouse models in Chapters 2 and

Figure 4.1: Gene expression of RAS enzymes 24 hours after AngII treatment.
0 1 10 100 1000
0.01
0.1
1
10
100
Ang II Concentration (nM)
Fold Change from Untreated
ACE1
✱
✱
NT 1 10 100 1000
0.1
1
10
100
1000
Ang II Concentration (nM)
Fold Change from Untreated
ACE2
✱
✱

Figure 4.2: Gene expression of RAS receptors 24 hours after AngII treatment.
0 1 10 100 1000
0.01
0.1
1
10
100
1000
Ang II Concentration (nM)
Fold Change from Untreated
AT1R
✱
✱
0 1 10 100 1000
0.1
1
10
100
1000
10000
Ang II Concentration (nM)
Fold Change from Untreated
AT2R
✱
✱ ✱✱
0 1 10 100 1000
0.1
1
10
100
1000
Ang II Concentration (nM)
Fold Change from Untreated
MasR
✱✱
✱

116
3 (Browatzki et al., 2005). As shown in Figure 4.3 below, escalating Ang II concentrations upregulated
MMP-2 across 24 from 1 nM to 1µM AngII. Similar to AT2R and MasR, gene expression of MMP-2
increased continuously, while gene expression during the 48-hour period was found to peak at 10 nM
and subsided at higher concentrations. Gene expression of MMP-3 did not show any clear trends, with
downregulation at 24 hours using 1 nM and 100 nM. At 48 hours, MMP-3 was downregulated using 1
nM and upregulated using 100 nM, again signaling no clear trend. MMP-9 expression was found to
increase at 24 hours. At 48 hours, gene expression of MMP-9 appears bell-shaped, with upregulation
occurring at 1 nM and 10 nM, with no changes at 100 nM and 1 µM. TIMP-2 expression at 24 hours was
found to be upregulated with 10 nM, 100 nM, and 1 µM concentrations. At 48 hours of AngII incubation,
gene expression for TIMP-2 was found to be upregulated at 10 and 100 nM but downregulated at 1 µM,
while gene expression of MMPs were shown to be modulated with Ang II incubation. Findings are
summarized in Figure 4.3.

Figure 4.3: Gene expression of metalloproteinases 24 hours after AngII treatment.

NT 1 10 100 1000
0.1
1
10
100
Ang II Concentration (nM)
Fold Change from Untreated
MMP2
✱
NT 1 10 100 1000
0.1
1
10
100
1000
10000
100000
Ang II Concentration (nM)
Fold Change from Untreated
MMP9
NT 1 10 100 1000
0.1
1
10
Ang II Concentration (nM)
Fold Change from Untreated
MMP3
NT 1 10 100 1000
0.1
1
10
Ang II Concentration (nM)
Fold Change from Untreated
TIMP2

117
4.3.2 Cell Injury and RAS Inhibition
Hydrogen peroxide (H 2O 2) treatment is a method employed to investigate cellular response to
oxidative stress induced tissue damage. Due to the variety of cell types and their sensitivity to H 2O 2,
treatment concentrations can typically range from 100 µM to 2 mM (Park, 2014; Shentu et al., 2018).
This in vitro model was used to dissect the molecular effects of laser-induced tissue damage to further
understand the in vivo data found in Chapters 2 and 3. H 2O 2 was used to induce oxidative stress in ARPE-
19 cells, for which single pulses of H 2O 2 were shown to be more tolerable but its effects are rapidly
eliminated making it a viable stimulus for immediate damage (Kaczara et al., 2010). Previously, we
showed extracellular matrix remodeling of MMPs and RAS component changes in this tissue injury
model. To mimic the in vivo models, ARPE-19 cells were incubated with H 2O 2 using concentration
escalation from 0.5 to 2mM, to determine whether oxidative stress levels correlate with gene
expression and metabolomic changes in these pathways. To determine the impact of H 2O 2 on RAS and
MMP systems, ARPE-19 were treated for 1 hour with H 2O 2 treatment to investigate the immediate
effects on ARPE-19 over this time horizon in parallel with the dramatic changes seen in the animal
studies. The same MMP and RAS genes analyzed in the animal models were probed to determine the
similarities between R:GEN and H 2O 2 treatments on RPE.

118
4.3.2.1 RT-PCR for Genes of RAS Components
Control (No RAS Inhibition)
Confluent ARPE-19 cells were treated with H 2O 2 in a concentration escalation. Gene expression
for MMP’s and RAS components were assessed using fold change in comparisons with untreated ARPE-
19. After H 2O 2 treatment, ACE1 was shown to be shown to be significantly upregulated in ARPE-19
treated with 2 mM H 2O 2. Similarly, ACE2 gene expression was also significantly upregulated at 1 hour
using the 2 mM concentration of H 2O 2 (p<0.05), as shown in Figure 4.4.
The expression of RAS receptors such as AT1R was also found to be significantly upregulated
after 1 hour of 2 mM H 2O 2 treatment while 1 mM exhibited downregulation. Gene expression of AT2R
was found to increase with H 2O 2 treatment where significant upregulation was observed at 1 mM and 2
mM. Similar to the other RAS components, MasR expression was also significantly upregulated at 1
hours with 2 mM H 2O 2 treatment. The findings are summarized in Figure 4.5. From the results, where 1

Figure 4.5: Gene expression changes in RAS receptors after H 2O 2treatment. Trends in expression suggest dose-
response with cellular damage.

0 mM 0.5 mM 1 mM 2 mM
0.01
0.1
1
10
100
1000
H2O2 Concentration
Fold Change from Untreated
AT1R
***
0 mM 0.5 mM 1 mM 2 mM
0.1
1
10
100
1000
H2O2 Concentration
Fold Change from Untreated
MasR
****
0 mM 0.5 mM 1 mM 2 mM
0.1
1
10
100
H2O2 Concentration
Fold Change from Untreated
AT2R
*
****

Figure 4.4: Gene expression changes in angiotensin converting enzymes after H 2O 2 treatment. Trends in
expression suggest dose-response with cellular damage.

0 mM 0.5 mM 1 mM 2 mM
0.1
1
10
100
H2O2 Concentration
Fold Change from Untreated
ACE1
*
0 mM 0.5 mM 1 mM 2 mM
0.1
1
10
100
H2O2 Concentration
Fold Change from Untreated
ACE2
**

119
mM H 2O 2 was selected as the concentration for all downstream inhibition experiments as it was capable
of inducing enough damage to illicit a strong cellular response without excessive cytotoxicity.
Enalapril
To determine the molecular mechanism driving the tissue injury response in RAS and MMPs,
ARPE-19 cells were incubated with 1 µM enalapril for 30 minutes to completely inhibit ACE1 (Gross et
al., 1981). Cells were then treated with increasing concentrations of H 2O 2 (0 to 2 mM) and RAS
component changes in relations to cellular damage. In the presence of enalapril alone, ACE1 was found
to be upregulated at 1 hour, relative to untreated controls although not statistically significant (p =
0.1421), as seen in Figure 4.6. After H 2O 2 treatment, ACE1 was also upregulated, although to a lesser
degree than H 2O 2 alone. At 1 hour, Enalapril also induced upregulation of ACE2, although not found to
be significant (p = 0.0589). However, after H 2O 2 treatment, Enalapril treated cells downregulated ACE2.
Treatment Group ACE (Mean ± StDev) ACE2 (Mean ± StDev)
NT 1.000 ± 0.075 1.000 ± 0.490
Enalapril 2.138 ± 0.075 2.714 ± 0.184
1 mM H 2O 2 1.096 ± 0.678 1.221 ± 0.452
Enalapril + 1 mM H 2O 2 1.481 ± 0.682 0.146 ± 0.017

Gene Comparisons Summary Adjusted P Value
ACE
NT vs. Enalapril ns 0.142
1 mM H 2O 2 vs. Enalapril + 1 mM H 2O 2 ns 0.295
ACE2
NT vs. Enalapril ns 0.059
1 mM H 2O 2 vs. Enalapril + 1 mM H 2O 2 * 0.006
Figure 4.6: Gene expression changes in RAS components with hydrogen peroxide treatment after ACE
inhibition. Trends in expression suggest dose-response with cellular damage.

120
After ACE inhibition using enalapril, AT1R was found to be upregulated at 1 hour when
compared to untreated cells, where the difference were not statistically significant. After 1 mM H 2O 2
treatment, AT1R expression was downregulated. However, in ARPE19 treated with both enalapril and
H 2O 2, significant upregulated of AT1R relative to H 2O 2 alone (p=0.033). Gene expression of AT2R was
found to be downregulated after ACE inhibition relative to untreated cells. 1 mM H 2O 2 appears to
upregulate AT2R expression, which was amplified by Enalapril treatment. MasR was found to be
unaffected by Enalapril treatment relative to untreated. 1 mM H 2O 2 treatment appears to have no effect
on MasR, both in Enalapril-treated cells and in controls. Findings are summarized in Figure 4.7.

Treatment Group AT1R (Mean ± StDev) AT2R (Mean ± StDev) MasR (Mean ± StDev)
NT 1.000 ± 1.590 1.000 ± 1.014 1.000 ± 0.567
Enalapril 5.519 ± 1.625 0.335 ± 0.204 0.674 ± 0.360
1 mM H 2O 2 0.093 ± 0.104 4.146 ± 0.526 0.303 ± 0.091
Enalapril + 1 mM H 2O 2 1.074 ± 0.694 100.3 ± 141.8 1.269 ± 0.578

Gene Comparisons Summary Adjusted P Value
AT1R
NT vs. Enalapril * 0.012
1 mM H 2O 2 vs. Enalapril + 1 mM H 2O 2 * 0.033
AT2R
NT vs. Enalapril ns 0.882
1 mM H 2O 2 vs. Enalapril + 1 mM H 2O 2 * 0.0003
MasR
NT vs. Enalapril ns 0.862
1 mM H 2O 2 vs. Enalapril + 1 mM H 2O 2 ns 0.068
Figure 4.7: Gene expression changes in RAS receptors after hydrogen peroxide treatment. Trends in expression
suggest dose-response with cellular damage.

121
Losartan
ARPE-19 cells were incubated with 1 µM of Losartan, a commercially used Angiotensin Receptor
Blocker (ARB), for 30 minutes to completely inhibit AT1R (Burnier, 2001). Cells were then treated with
increasing concentrations of hydrogen peroxide (0 mM, 0.5 mM, 1 mM, 2 mM) to explore the gene
expression changes that would occur with this cellular damage model after blocking activation of
pathogenic RAS receptors. After ARB treatment, ACE1 was found to be significantly upregulated at 1
hour relative to untreated. Using 1 mM of hydrogen peroxide treatment, ACE1 was upregulated similar
to no hydrogen peroxide treatment. Losartan alone also upregulated gene expression for ACE2
compared to untreated cells, and 1 mM of H 2O 2 was able to further upregulate ACE2 expression, as
summarized in Figure 4.8.
Treatment Group ACE (Mean ± StDev) ACE2 (Mean ± StDev)
NT 1.000 ± 0.075 1.000 ± 0.490
Losartan 2.295 ± 1.450 8.085 ± 12.069
1 mM H 2O 2 1.096 ± 0.678 1.221 ± 0.452
Losartan + 1 mM H 2O 2 2.424 ± 0.914 2.267 ± 1.741

Gene Comparisons Summary Adjusted P Value
ACE
NT vs. Losartan * 0.007
1 mM H 2O 2 vs. Losartan + 1 mM H 2O 2 ns 0.174
ACE2
NT vs. Losartan ns 0.257
1 mM H 2O 2 vs. Losartan + 1 mM H 2O 2 ns 0.052
Figure 4.8: Gene expression changes in RAS enzymes with hydrogen peroxide treatment after AT1R inhibition.
Trends in expression suggest dose-response with cellular damage.

122
After ARB treatment, AT1R was found to be significantly upregulated at 1 hour for no H 2O 2 as
well as using 1 mM of H 2O 2. After ARB treatment, gene expression of AT2R was found to be upregulated
relative to untreated cells. 1 mM of H 2O 2 treatment was able to upregulate gene expression of AT2R and
amplified by Losartan treatment. MasR was similarly upregulated by ARB treatment. Furthermore, gene
expression of MasR was reduced with H 2O 2 treatment while upregulated using the combination
treatment. Findings are summarized in Figure 4.9.

Effect of A779
ARPE-19 cells were incubated with 1 µM of A779 for 30 minutes to determine what effect would
MasR inhibition (R. A. Santos et al., 2003) have on ARPE19 response toward H 2O 2 insult. ARPE19 were
treated with H 2O 2 from 0.5 to 2 mM. The addition of 1mM A779 was used to determine the effect of
MasR mediated effects. A779 treatment was found to have no effect on ACE1. However, ACE1 was
upregulated at one hour using A779 in combination with 1mM of H 2O 2. Gene expression for ACE2 was
Treatment Group AT1R (Mean ± StDev) AT2R (Mean ± StDev) MasR (Mean ± StDev)
NT 1.000 ± 1.590 1.000 ± 1.014 1.000 ± 0.567
Losartan 139.2 ± 207.5 4.362 ± 4.406 1.733 ± 1.637
1 mM H 2O 2 0.093 ± 0.104 4.146 ± 0.526 0.303 ± 0.091
Losartan + 1 mM H 2O 2 62.61 ± 61.95 68.64 ± 74.60 1.035 ± 1.397

Gene Comparisons Summary Adjusted P Value
AT1R
NT vs. Losartan * <0.001
1 mM H 2O 2 vs. Losartan + 1 mM H 2O 2 * <0.001
AT2R
NT vs. Losartan ns 0.239
1 mM H 2O 2 vs. Losartan + 1 mM H 2O 2 * 0.006
MasR
NT vs. Losartan ns 0.114
1 mM H 2O 2 vs. Losartan + 1 mM H 2O 2 * 0.050
Figure 4.9: Gene expression changes in RAS receptors after hydrogen peroxide treatment after AT1R inhibition.
Trends in expression suggest dose-response with cellular damage.

123
also found to be upregulated with A779 relative to untreated cells. However, gene expression was
unchanged relative to untreated after H 2O 2 treatment, Figure 4.10.
AT1R was found to be significantly upregulated at 1 hour relative to untreated cells and
upregulated in combination with 1 mM H 2O 2, although lower than A779 alone. No changes in gene
expression of AT2R were found at 1 hour with A779, and 1 mM of H 2O 2 upregulated AT2R, but A779
modulation of AT2R gene expression appears to be unaffected by H 2O 2. MasR expression was
upregulated by A779 relative to untreated. 1 mM H 2O 2 downregulated MasR expression while
combination with A779 significantly upregulated MasR. Findings are summarized in Figure 4.11 below.

Treatment Group AT1R (Mean ± StDev) AT2R (Mean ± StDev) MasR (Mean ± StDev)
NT 1.000 ± 1.590 1.000 ± 1.014 1.000 ± 0.567
A779 146.8 ± 253.3 4.362 ± 4.406 1.247 ± 0.524
1 mM H 2O 2 0.093 ± 0.104 4.146 ± 0.526 0.303 ± 0.091
A779 + 1 mM H2O2 62.61 ± 61.95 68.64 ± 74.60 16.58 ± 24.78

Gene Comparisons Summary Adjusted P Value
AT1R
NT vs. A779 * 0.013
1 mM H 2O 2 vs. A779 + 1 mM H2O2 * 0.003
AT2R
NT vs. A779 * 0.006
1 mM H 2O 2 vs. A779 + 1 mM H2O2 * 0.010
MasR
NT vs. A779 ns 0.124
1 mM H 2O 2 vs. A779 + 1 mM H2O2 * 0.007
Figure 4.11: Gene expression changes in RAS components with hydrogen peroxide treatment after Mas
inhibition. Trends in expression suggest dose-response with cellular damage.

Treatment Group ACE (Mean ± StDev) ACE2 (Mean ± StDev)
NT 1.000 ± 0.075 1.000 ± 0.490
A779 0.651 ± 0.163 2.376 ± 2.381
1 mM H 2O 2 1.096 ± 0.678 1.221 ± 0.452
A779 + 1 mM H 2O 2 2.927 ± 3.858 0.647 ± 0.690

Gene Comparisons Summary Adjusted P Value
ACE
NT vs. A779 ns 0.999
1 mM H 2O 2 vs. A779 + 1 mM H 2O 2 ns 0.572
ACE2
NT vs. A779 ns 0.121
1 mM H 2O 2 vs. A779 + 1 mM H 2O 2 ns 0.230
Figure 4.10: Gene expression changes in RAS components with hydrogen peroxide treatment after Mas
inhibition. Trends in expression suggest dose-response with cellular damage.

124

PD123319
ARPE-19 cells were incubated with 1 µM of PD123319, for 30 minutes to completely inhibit
AT2R. Cells were then treated with increasing concentrations of hydrogen peroxide (0 mM, 0.5 mM, 1
mM, 2 mM) to explore the gene expression changes that would occur with this cellular damage model
after blocking activation of pro-regenerative RAS receptors. After PD123319 treatment, alone and in
combination with H 2O 2, no changes were found in ACE1 expression, shown in Figure 4.12. PD123319
downregulated ACE2 expression when used alone, but upregulated ACE2 expression when A779 was
used in combination with 1 mM H 2O 2. With PD123319 treatment, expression for ACE2 was
downregulated relative to untreated. 1 mM treatment with hydrogen peroxide treatment had no real
effect on ACE2, but combination treatment with PD123319 upregulated ACE2.
No changes were observed in AT1R expression when treated with PD123319 alone. However,
AT1R expression was found to be significantly upregulated using combination of 1 mM H 2O 2. No changes
in gene expression of AT2R were found at 1 hour using PD123319 alone but were found to be
upregulated using combination 1 mM H 2O 2 and PD123319. MasR was found to be downregulated with
PD123319 treatment at 1 hour. Combination PD123319 and 1 mM H 2O 2 was found to have no impact on
gene expression of MasR relative to untreated. Findings are summarized in Figure 4.13 below.
Treatment Group ACE (Mean ± StDev) ACE2 (Mean ± StDev)
NT 1.000 ± 0.075 1.000 ± 0.490
PD123319 0.670 ± 0.451 0.128 ± 0.097
1 mM H 2O 2 1.096 ± 0.678 1.221 ± 0.452
PD123319 + 1 mM H2O2 1.140 ± 0.036 4.644 ± 6.254

Gene Comparisons Summary Adjusted P Value
ACE
NT vs. PD123319 ns 0.385
1 mM H 2O 2 vs. PD123319 + 1 mM H2O2 ns 0.433
ACE2
NT vs. PD123319 ns 0.095
1 mM H 2O 2 vs. PD123319 + 1 mM H2O2 ns 0.142
Figure 4.12: Gene expression changes in RAS enzymes with hydrogen peroxide treatment after AT2R inhibition.
Trends in expression suggest dose-response with cellular damage.

125

MLN4760
ARPE-19 cells were incubated with 10 µM of MLN4760, for 30 minutes to completely inhibit
ACE2. After MLN4760 treatment, alone and in combination with H 2O 2, no changes were found in ACE1
expression. MLN4760 appeared to have no effect on ACE2 expression when used alone, but
Treatment Group ACE (Mean ± StDev) ACE2 (Mean ± StDev)
NT 1.000 ± 0.075 1.000 ± 0.490
MLN4760 1.388 ± 0.593 0.587 ± 0.270
1 mM H 2O 2 1.096 ± 0.678 1.221 ± 0.452
MLN4760 + 1 mM H 2O 2 0.848 ± 0.507 0.241 ± 0.129

Gene Comparisons Summary Adjusted P Value
ACE
NT vs. MLN4760 ns 0.451
1 mM H 2O 2 vs. MLN4760 + 1 mM H 2O 2 ns 0.430
ACE2
NT vs. MLN4760 ns 0.247
1 mM H 2O 2 vs. MLN4760 + 1 mM H 2O 2 * 0.029
Figure 4.14: Gene expression changes in RAS enzymes with hydrogen peroxide treatment
after ACE2 inhibition. Trends in expression suggest dose-response with cellular damage.

Treatment Group AT1R
(Mean ± StDev)
AT2R
(Mean ± StDev)
MasR
(Mean ± StDev)
NT 1.000 ± 1.590 1.000 ± 1.014 1.000 ± 0.567
PD123319 1.162 ± 0.117 0.544 ± 0.533 0.109 ± 0.079
1 mM H 2O 2 0.093 ± 0.104 4.146 ± 0.526 0.303 ± 0.091
PD123319 + 1 mM H 2O 2 228.6 ± 321.7 47.09 ± 64.26 0.444 ± 0.403

Gene Comparisons Summary Adjusted P
Value
AT1R
NT vs. PD123319 ns 0.288
1 mM H 2O 2 vs. PD123319 + 1 mM H 2O 2 * 0.019
AT2R
NT vs. PD123319 ns 0.665
1 mM H 2O 2 vs. PD123319 + 1 mM H 2O 2 ns 0.296
MasR
NT vs. PD123319 ns 0.058
1 mM H 2O 2 vs. PD123319 + 1 mM H 2O 2 ns 0.361
Figure 4.13: Gene expression changes in RAS components with hydrogen peroxide treatment after
AT2R inhibition. Trends in expression suggest dose-response with cellular damage.

126
downregulated ACE2 expression when cells were treated with MLN4760 in combination with 1 mM
H 2O 2, as seen in Figure 4.14.
AT1R was found to be upregulated relative to untreated cells after 1 hour of MLN4760
treatment. While gene expression of AT1R was unchanged from untreated after combination treatment
with 1 mM concentrations of H 2O 2, it was a statistically significant difference from H 2O 2 treatment alone.
Upregulation of AT2R occurred using MLN4760 alone but was downregulated in combination with 1 mM
H 2O 2. MasR expression was found to be unchanged when using MLN4760 alone and in combination with
1 mM H 2O 2. Findings are summarized in Figure 4.15.
Treatment Group AT1R
(Mean ± StDev)
AT2R
(Mean ± StDev)
MasR
(Mean ± StDev)
NT 1.000 ± 1.590 1.000 ± 1.014 1.000 ± 0.567
MLN4760 11.19 ± 10.28 3.404 ± 3.428 1.111 ± 0.726
1 mM H 2O 2 0.093 ± 0.104 4.146 ± 0.526 0.303 ± 0.091
MLN4760 + 1 mM H 2O 2 1.697 ± 0.479 0.487 ± 0.425 0.633 ± 0.820

Gene Comparisons Summary Adjusted P
Value
AT1R
NT vs. MLN4760 * 0.002
1 mM H 2O 2 vs. MLN4760 + 1 mM H 2O 2 * 0.010
AT2R
NT vs. MLN4760 ns 0.507
1 mM H 2O 2 vs. MLN4760 + 1 mM H 2O 2 * 0.023
MasR
NT vs. MLN4760 ns 0.110
1 mM H 2O 2 vs. MLN4760 + 1 mM H 2O 2 ns 0.398
Figure 4.15: Gene expression changes in RAS receptors with hydrogen peroxide treatment
after ACE 2inhibition. Trends in expression suggest dose-response with cellular damage.

127
4.3.2.2 RT-PCR for Genes of RAS Receptor Activation
To confirm the activation of RAS receptors, gene expression of downstream signals NADPH
oxidase 2 (NOX-2), monocyte chemoattractant protein-1 (MCP-1), and Src homology 2-containing
protein-tyrosine phosphatase-1 (SHP-1) were probed. NOX-2 was found to be upregulated when
pretreated with Losartan and A779 prior to 1 mM H 2O 2 treatment. MCP-1 was found to be upregulated
with Losartan pretreatment but downregulated with every other inhibitor prior to 1 mM H 2O 2
treatment. SHP-1 was also found to be upregulated with Losartan pretreatment, while all other
inhibitors had no effect on SHP-1 expression prior to peroxide. These findings as summarized in Figure
4.16, suggests the possibility of compensatory upregulation after AT1R inhibition with losartan, as other
inhibitors did not have the same effect at 1 hour and 1 mM peroxide treatment.

Figure 4.16: Gene expression signals downstream of RAS receptors with RAS inhibitor pretreatment, followed
by 1 mM H 2O 2 treatment.

0.01
0.1
1
10
100
1000
H2O2 Concentration
Fold Change from Untreated
NOX-2
NT Control Enalapril Losartan A779 PD123319 MLN4760
0.01
0.1
1
10
100
1000
H2O2 Concentration
Fold Change from Untreated
SHP-1
NT Control Enalapril Losartan A779 PD123319 MLN4760
0.01
0.1
1
10
100
1000
H2O2 Concentration
Fold Change from Untreated
MCP-1
NT Control Enalapril Losartan A779 PD123319 MLN4760

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4.3.2.3 RT-PCR for Genes of Extracellular Matrix Remodeling
Control
Gene expression of metalloproteinases were probed to determine if there were any similarities between
H 2O 2 -induced damage and the observed extracellular matrix remodeling in Chapters 2 and 3. MMP-2
was found to be unchanged until 2 mM of H 2O 2 at 1 hour. MMP-3 was found to be downregulated, until
upregulation occurred using the 2 mM H 2O 2 treatment at 1 hour. MMP-9 was found to be
downregulated with 1 mM H 2O 2 and upregulated with 2 mM H 2O 2 at 1 hour. TIMP-2 was found to be
upregulated using 2 mM H 2O 2 at 1 hour as well. Findings are summarized in Figure 4.17.

Figure 4.17: Gene expression changes in MMP's after hydrogen peroxide treatment. Trends in expression
suggest dose-response with cellular damage.

0 mM
0.5 mM
1 mM
2 mM
0.1
1
10
100
H2O2 Concentration
Fold Change from Untreated
MMP2
**
0 mM
0.5 mM
1 mM
2 mM
0.1
1
10
100
H2O2 Concentration
Fold Change from Untreated
MMP9
0 mM
0.5 mM
1 mM
2 mM
0.1
1
10
100
H2O2 Concentration
Fold Change from Untreated
MMP3
*
0 mM
0.5 mM
1 mM
2 mM
0.1
1
10
100
H2O2 Concentration
Fold Change from Untreated
TIMP2
*

129
Genes of ECM Remodeling After RAS Inhibition
Looking at the 1-hour time point of 1 mM combination treatment with the RAS inhibitors as
representative of the entire data set, gene expression of MMP-2 was found to be upregulated with
Losartan but downregulated using A779 and MLN4760. No changes were observed with the other
inhibitors. MMP-3 was found to be upregulated with Losartan but downregulated with A779, PD123319,
and MLN4760. No changes were observed with Enalapril. MMP-9 was upregulated using Losartan,
PD123319, and MLN4760, while no changes were observed with Enalapril and A779 treatment. TIMP-2
was found to be upregulated using Enalapril, Losartan, and A779, while no changes were observed with
PD123319 and MLN4760 treatment. Findings are summarized in Figure 4.18.

Figure 4.18: Gene expression changes in MMP’s when inhibited prior to hydrogen peroxide treatment. Trends
in expression suggest dose-response with cellular damage.

NT Control Enalapril Losartan A779 PD123319 MLN4760
0.01
0.1
1
10
100
1000
1 mM H2O2
Fold Change from Untreated
MMP2
NT Control Enalapril Losartan A779 PD123319 MLN4760
0.01
0.1
1
10
100
1000
1 mM H2O2
Fold Change from Untreated
MMP9
NT Control Enalapril Losartan A779 PD123319 MLN4760
0.01
0.1
1
10
100
1000
1 mM H2O2
Fold Change from Untreated
MMP3
NT
Control
Enalapril
Losartan
A779
PD123319
MLN4760
0.01
0.1
1
10
100
1000
1 mM H2O2
Fold Change from Untreated
TIMP2

130
4.3.3.1 LC-MS/MS RAS Metabolomics
Collected cell media was analyzed for changes in RAS peptides after H2O2 treatment with and
without RAS inhibition, as summarized in Table 4.4. The 1 mM H2O2 treatment group was selected as
representative of RAS changes due to its previously established usage in ARPE-19 assays (Hu, Wu, Zhao,
& Wang, 2019). Ang(1-10) and Ang(1-9) were found to be increased in all inhibitor treatment groups
relative to 1 mM H2O2 alone. Interestingly, Ang II was only found in lower concentrations in the
PD123319 inhibitor group compared to the 1 mM treatment group, while all other inhibitors still had
higher amounts of Ang II. Meanwhile, all inhibitor groups had lower levels of Ang (1-7) than 1 mM of
H 2O 2 treatment, except for the Losartan group, which was found to have higher levels of Ang(1-7).
Paired t-test analyses were conducted between the levels of bioactive peptides between 1 mM
H 2O 2 that treatment groups and RAS inhibition groups, as summarized in Table 4.5. Statistical
significance was met for concentrations of Ang(1-9) within the combination H 2O 2 + A779 treatment

RAS Peptide Concentrations (ng/mL) (Mean ± StDev)– 1 Hr
Treatment Ang(1-10) Ang(1-9) Ang II Ang(1-7)
No Treatment 1.034 ± 0.353 0.096 ± 0.085 0.137 ± 0.035 0.111 ± 0.092
1mM H 2O 2 7.846 ± 10.105 0.831 ± 0.516 2.054 ± 1.819 0.031 ± 0.007
Enalapril + 1mM H 2O 2 1.437 ± 0.502 0.144 ± 0.108 1.581 ± 1.075 0.031 ± 0.001
Losartan + 1mM H 2O 2 0.926 ± 0.188 0.052 ± 0.006 1.871 ± 1.457 0.032 ± 0.005
A779 + 1mM H 2O 2 1.845 ± 1.831 1.367 ± 0.711 2.298 ± 0.035 0.063 ± 0.015
PD123319 + 1mM H 2O 2 20.046 ± 25.828 2.530 ± 1.570 5.975 ± 7.054 0.095 ± 0.004
MLN4760 + 1mM H 2O 2 29.942 ± 46.091 2.688 ± 4.286 7.008 ± 7.431 0.196 ± 0.285
Table 4.4: Cell media RAS peptides after H 2O 2 treatment with RAS enzyme and receptor inhibition.
RAS Peptide Concentrations (Adjusted P Values) – 1 Hr
Treatment Groups Ang(1-10) Ang(1-9) Ang II Ang(1-7)
NT vs. 1 mM H 2O 2 0.48 0.82 0.45 0.06
NT vs. Enalapril + 1 mM H 2O 2 0.66 0.50 0.47 0.06
NT vs. Losartan + 1 mM H 2O 2 0.40 0.42 0.05* 0.10
NT vs. A779 + 1 mM H 2O 2 0.34 0.04* 0.22 0.81
NT vs. PD123319 + 1 mM H 2O 2 0.91 0.29 0.41 0.24
NT vs. MLN4760 + 1 mM H 2O 2 0.42 0.54 0.55 0.67
Table 4.5: One-way ANOVA of RAS peptides after H 2O 2 treatment with RAS enzyme and receptor inhibition.
Multiple comparisons with the untreated group.

131
group. While not statistically significant, trends were observed in Ang(1-7) levels when inhibitors for the
pathogenic arm of RAS were used in contrast to inhibitors of the protective arm (bolded).
To verify the enzymatic activity of ACE and ACE2, peptide ratios were calculated using
product/reactant. The ratio of Ang II/Ang(1-10) is indicative of ACE activity and suggests that Ang II
production had increased with Enalapril and Losartan pretreatment, but decreased with A779,
PD123319, and MLN4750. However, the ratio of Ang(1-7)/Ang(1-9), which is also metabolized by ACE,
suggests that all metabolism of Ang(1-9) had been blunted. The ratio of Ang(1-7)/Ang II, which would be
indicative of ACE2 activity, had shown a decrease across all inhibition treatments. Conversely, Ang(1-
9)/Ang(1-10), which is also a proxy of ACE2 activity, showed increases in the treatment groups that
inhibited the protective arm of RAS, while the ratios were decreased in the pathogenic arm, relative to
untreated. Findings are summarized in Figure 4.19.

RAS Peptide Ratios (Mean ± StDev)
Treatment Ang II /Ang(1-10) Ang(1-7)/Ang II Ang(1-7)/Ang(1-9) Ang(1-9)/Ang(1-10)
NT 0.154 ± 0.101 0.935 ± 0.933 1.208 ± 0.223 0.088 ± 0.054
1mM H 2O 2 0.435 ± 0.016 0.025 ± 0.018 0.056 ± 0.051 0.250 ± 0.212
Enalapril + 1mM H 2O 2 1.107 ± 0.001 0.025 ± 0.012 0.308 ± 0.195 0.104 ± 0.062
Losartan + 1mM H 2O 2 1.876 ± 0.125 0.025 ± 0.015 0.618 ± 0.063 0.057 ± 0.014
A779 + 1mM H 2O 2 1.141 ± 0.030 0.037 ± 0.015 0.585 ± 0.463 0.994 ± 0.383
PD123319 + 1mM H 2O 2 0.341 ± 0.028 0.033 ± 0.022 0.044 ± 0.016 0.240 ± 0.137
MLN4760 + 1mM H 2O 2 0.625 ± 0.034 0.023 ± 0.015 0.123 ± 0.047 0.098 ± 0.067
Figure 4.19: Effect of RAS enzyme and receptor inhibitor on RAS peptide ratios in cell media after one hour of
H 2O 2 treatment.

NT 1 mM Enalapril Losartan A779 PD123319 MLN4760
0.1
1
10
Peptide Ratio
AngII/Ang(1-10)
NT 1 mM Enalapril Losartan A779 PD123319 MLN4760
0.01
0.1
1
10
Peptide Ratio
Ang(1-7)/Ang(1-9)
NT 1 mM Enalapril Losartan A779 PD123319 MLN4760
0.01
0.1
1
10
Peptide Ratio
Ang(1-7)/AngII
NT 1 mM Enalapril Losartan A779 PD123319 MLN4760
0.01
0.1
1
10
Peptide Ratio
Ang(1-9)/Ang(1-10)

132

4.3.3.2 Integrated RAS: Gene Expression and Peptides
Angiotensin peptides were arranged to map the peptide, enzyme, and receptor levels.
Upregulated signals are represented in green, while downregulation is represented in red, as seen in
Figure 4.20. In the 1 mM H 2O 2 treatment group, ACE1 and ACE2 were both found to be upregulated,
while AT1R was downregulated and AT2R was upregulated. MasR was observed to be downregulated
relative to untreated. Downstream, MCP-1 was found to be downregulated, matching expectations from
downregulation of AT1R and upregulation of AT2R.

Figure 4.20: RAS components Gene expression and peptide levels changes after H 2O 2
induction
NT
Ang(1-10)
797.5 pM
Ang(1-9)
81.4 pM
Ang(1-7)
123.4 pM
AngII
129.9 pM
ACE
ACE2
ACE
ACE2
AT1
AT 2 Mas
MCP-1 SHP-1 NOX-2
Ang(1-10)
583.3 pM
Ang(1-9)
21.3 pM
Ang(1-7)
67.3 pM
AngII
71.9 pM
ACE
ACE2
ACE
ACE2
AT1
AT 2 Mas
MCP-1 SHP-1 NOX-2
1 mM H
2
O
2

133
Combination Losartan and H 2O 2 treated cells were previously identified as having significantly
different levels of Ang(1-10) when compared to H 2O 2 alone 1 hour after treatment. All genes and
peptides were found to be upregulated, as seen in Figure 4.21. However, when compared against 1 mM
H2O2 alone, the data would suggest the increases in gene expression to occur in AT1R and MasR are
derived from Losartan treatment, influencing the downstream activation of NOX-2, MCP-1, and SHP-1.
Combination MLN4760 and H 2O 2 treated cells were also identified as having significantly
different levels of Ang(1-10) when compared to H 2O 2 alone 1 hour after treatment. Gene expression of
ACE2 and MCP-1 were downregulated, while all other genes were found to be unchanged from
untreated, as seen in Figure 4.22. Levels of Ang(1-10) were found to be significantly higher than those of
1 mM peroxide treatment alone, while levels of Ang(1-9), Ang II, and Ang(1-7) were lower. The
downregulated gene expression of ACE2 confirms the relative decrease in Ang(1-9) and Ang(1-7) levels.
As the levels of Ang(1-10) increased with no observed change in ACE, and decreased levels of Ang II
suggest binding of Ang II to AT2R as MCP-1 was found to be downregulated. Interestingly, these results

Figure 4.21: Gene expression changes in RAS components in response to H 2O 2 induction, and the
impact of losartan on the RAS metabolism after AT1R inhibition.

Losartan
Losartan + 1 mM H
2
O
2
Ang(1-10)
818.3 pM
Ang(1-9)
57.2 pM
Ang(1-7)
40.9 pM
AngII
100.3 pM
ACE
ACE2
ACE
ACE2
AT1 AT2 Mas
MCP-1 SHP-1 NOX-2
Ang(1-10)
715 pM
Ang(1-9)
44 pM
Ang(1-7)
36 pM
AngII
1788 pM
ACE
ACE2
ACE
ACE2
AT1 AT2 Mas
MCP-1 SHP-1 NOX-2
Losartan Losartan

134
would also suggest an alternative pathway that is activated by the tissue damage to generate Ang(1-7)
from the excess Ang(1-10) produced, rather than the canonical Ang(1-7)/AngII axis.
TGF-β Response to Peroxide Damage
Gene expression for TGF-β was found to be significantly upregulated after 2 mM H 2O 2 induction
after 1 hour. The addition of either ACEi or ARB was able to decrease TGF-β expression relative to H 2O 2
treatment alone. When cells were treated with enalapril and H 2O 2 for 1 hour, the data suggests this
combination was able to prevent upregulation of TGF-β. The use of ACEi suggest that RAS modulation
can prevent TGF-b expression and thus reduce the fibrotic wound healing on RPE. This suggest that the
administration of enalapril may have benefits in individuals undergoing SRT treatment, especially in
patient populations with hypertension and AMD.

Figure 4.22: Gene expression changes in RAS components with hydrogen peroxide treatment after ACE2
inhibition.

MLN4760
Ang(1-10)
978.8 pM
Ang(1-9)
80.2 pM
Ang(1-7)
30.3 pM
AngII
120.0 pM
ACE
ACE2
ACE
ACE2
AT1
AT
2 Mas
MCP-1 SHP-1 NOX-2
Ang(1-10)
1147.7 pM
Ang(1-9)
79.77 pM
Ang(1-7)
62.0 pM
AngII
102.0 pM
ACE
ACE2
ACE
ACE2
AT1
AT
2 Mas
MCP-1 SHP-1 NOX-2
MLN4760 + 1 mM H
2
O
2
MLN4760 MLN4760

135
4.3.1.3 TGF-β Response: R:GEN vs Ang II Dose Escalation
To determine if the upregulation in AngII that occurred post-R:GEN treatment was likely to
induce fibrosis via TGF- β, the ocular concentrations measured in Chapter 3 were converted to molar
units and overlapped with the dose escalation data conducted in ARPE-19 cells. As seen in Figure 4.24,

Figure 4.23: Gene expression changes in TGF-β with H 2O 2 treatment after RAS inhibition.

NT Control Enalapril Losartan
0.1
1
10
100
2 mM H2O2
Fold Change from Untreated
TGF-B - 1 Hr
NT
Control
Enalapril
Losartan

Figure 4.24: Plot comparing TGF- β1 expression and molar Ang II concentrations of dose escalation treatment
and R:GEN treatment.

0.01 0.1 1 10 100 1000 10000
0.00001
0.0001
0.001
0.01
0.1
1
10
100
Concentration (nM)
Fold Change from Untreated
Dose-Response TGFB1
24 Hr Ang II Treatment
Ang II - 100% Intensity
Ang II - 50% Intensity

136
the concentrations of AngII measured from the ocular fluid of CxCr5 -/- mice were much lower than the
1µM concentration required to induce increased in TGF- β gene expression, suggesting the increase in
TGF- β after R:GEN treatment is more likely to have been from the disruption of ECM and release of
MMP’s.

4.4 Discussion
In this chapter, the effect of AngII and H 2O 2 was used to mimic R:GEN induced tissue damage on
RAS and MMP in ARPE-19 (Lazzarini et al., 2022). Additionally, RAS enzyme and receptor inhibition at
concentrations capable of complete saturation to dissect the molecular mechanism of tissue injury
response.
Specific inhibition of the RAS components targeting ACE1, ACE2, AT1R, AT2R and Mas were
employed to determine the impact of angiotensin peptide following H 2O 2 induced tissue damage. This will
help us understand the role of angiotensin receptors in relations to overall impact of wound healing. In
Chapters 2 and 3, it was established that R:GEN treatment modulated the expression of RAS metabolic
enzymes and receptors at early timepoints (1 hours) after treatment. To better characterize the
immediate effects of damage on the RPE in combination with RAS modulation, we sought to parallel the
1 hour time point utilized in our prior animal studies. It is expected that the changes in peptide and
receptor levels observed in this in vitro study will provide additional insights into the biological impact
that the RAS has on wound healing in the eye.
Gene expression and metabolomics were utilized to provide insights related to our molecular
probing. This approach builds upon our reported animal studies with the additional use of inhibitors to
shunt peptides down particular arms of the pathway to determine RAS activation.

137
4.4.1 Angiotensin II Upregulates Genes in Dose-Response Relationship
It has previously been reported that Ang II/AT1R activation can upregulate TGF- β, which in turn
promotes pro-fibrotic wound healing (Border & Noble, 1998). The effect of Ang II treatment on ARPE-19
can upregulate TGF-b gene expression. While a concentration dependent effect of Ang II on TGF- β
expression was expected, however TGF-β did not upregulated until 1 µM of Ang II. However, 1 µM is
significantly greater than the AngII concentrations observed in mice after R:GEN treatment, suggesting
the observed TGF- β action from R:GEN is not due to Ang II binding to AT1R but rather from the release
of MMP’s and TGF- β from damaged extracellular matrix (Song, Klepeis, Nugent, & Trinkaus-Randall,
2002). It would be surmised that the amount of Ang II induced after R:GEN is unlikely to activate the
pathogenic arm of RAS and thus contribute to profibrotic healing. Additionally, both ACE and ACE2 were
found to be significantly upregulated in response to Ang II treatment, indicating that the ARPE-19 cells
compensate for the higher AngII levels by enhance clearance. Increased RAS enzyme production to may
be an adaptive cellular response to reduce the stress caused by high levels of Ang II. Furthermore,
upregulation of AT1R gene expression by Ang II induction was previously confirmed in literature (Mitra,
Gao, & Zucker, 2010).
4.4.2 Hydrogen Peroxide Treatment Alters RAS in ARPE-19 Cells
H 2O 2 has previously been utilized to study cell damage responses in ARPE-19 (Hu et al., 2019). In
this study, we explored the effect of damage on human RPE with regards to the RAS. RAS peptides have
previously been reported to play a role in corneal wound healing process, for which activation of AT1R
can promote fibrosis while activation of AT2R and MasR promote non-fibrotic healing (Abdallah et al.,
2016; K. Rodgers et al., 1997). To probe the hypothesis that shifting RAS towards the pro-regenerative
arm will lead to non-fibrotic wound healing, we utilized inhibitors specific for RAS enzymes and
receptors to direct peptides towards either the pathogenic or pro-regenerative arm.

138
ARPE-19 cells were treated with H 2O 2 up to 2 mM, upregulating ACE, ACE2, and AT2R.
Simultaneously, AT1R and MCP-1 were downregulated. While gene expression was shown to be wildly
altered with RAS inhibitors, with regards to RAS peptides, only Losartan and MLN4760 treated cells had
produced concentrations statistically significantly different from hydrogen peroxide treatment alone.
Surprisingly, all genes, including AT1R, were upregulated with Losartan treatment, although it has
previously been reported that Losartan increases the mRNA expression of AT1R, suggesting there may
be some degree of compensatory upregulation when inhibiting such a critical receptor (T. Wang, Lian,
Cai, Lin, & Xie, 2018). Increased Ang(1-10), Ang(1-9), Ang II, and Ang(1-7) biosynthesis confirmed the
upregulation of ACE and ACE2.
Losartan treatment increased all RAS peptides. While, ACE2 inhibition via MLN4760 was found
to increase the formation of Ang(1-10) while decreasing the formation of downstream metabolites.
Treatment with MLN4760 was able to downregulate the gene expression of ACE2 and MCP-1 when
compared to untreated ARPE-19. Downregulation of ACE2 expression was confirmed with lower
concentrations of Ang(1-9) and Ang(1-7). In addition, Ang II decreased was accompanied by
accumulation of Ang(1-10) indicating that Ang II is being utilized the activate its downstream receptors.
However, because there is no upregulation of NOX-2 and MCP-1 is downregulated, it is possible that Ang
II is primarily activating AT2R.
RAS and Implications for Fibrosis
To initiate and explore the effect that RAS has on fibrotic wound healing, gene expression for
TGF-β was measured in ARPE-19 cells treated with H2O2 in a dose-escalation manner until a maximum 2
mM concentration. Hydrogen peroxide has previously been reported to be an effective inducer of
apoptosis in cells of all types, ranging from 100 µM to 1.6 mM in concentrations (Xiang, Wan, Guo, &
Guo, 2016). For our purposes, we wanted to generate microbubbles to emulate R:GEN’s mechanism of

139
action. We hypothesized that the catalase activity of ARPE-19 would rapidly decompose H 2O 2 to form
oxygen, simulating R:GEN’s subthreshold laser.
Our findings show that high doses of H 2O 2 is capable of inducing upregulation of TGF-β. As
expected, the addition of Enalapril decreased TGF-β expression relative to hydrogen peroxide treatment
alone. Unexpectedly, TGF- β was still upregulated with Losartan treatment in hour 1, as shown in Figure
4.16. Surprisingly, TGF- β was not upregulated at 1 hour with A779 but was with PD123319 (not shown),
suggesting that AT2R activation may be more critical than MasR in reducing expression of TGF-β in the
immediate phase of wound healing. Overall, these findings suggest that RAS modulation does play a role
in downstream mechanisms of wound healing on RPE and may have implications for optimizing the
efficacy of SRT in future development.
Challenges commonly exist when translating in vitro and in vivo studies. For example, while RPE
are normally pigmented in vivo, inducing pigmentation in ARPE-19 is a known challenge, therefore limiting
our ability to use the R:GEN as the laser is specific for the melanosomes of the eye. Rather, hydrogen
peroxide (H2O2) was used in our in vitro model as a widely accepted method for causing oxidative damage
and stress in cell-based models (Cao, Liu, Tuo, Shen, & Chan, 2010). Although catalase, the enzyme
responsible for catalyzing H 2O 2 to H 2O and O 2, has previously been shown to be extremely active in ARPE-
19, it was unable to decompose peroxides rapidly enough to form microbubbles within the RPE to better
parallel the activity of R:GEN (George, 1947; Kurz, Karlsson, Brunk, Nilsson, & Frennesson, 2009).
However, because oxidative stress is a known driver of AMD pathogenesis, these findings provide
additional evidence that therapeutic benefit may come from modulation of RAS. Enalapril was shown to
ameliorate the upregulation of TGF-β relative to control, suggesting ACE inhibition may aid RPE’s ability
to mitigate a known driver of disease progression. To supplement these findings, future studies may
consider repeating this study design, but culture ARPE-19 with chronic low-dose H 2O 2 treatment to better
mimic real world chronic oxidative stress.

140
4.5 Conclusion
In this chapter, H 2O 2 mediated ocular damage on of RAS and MMP components were evaluated
in human RPE cell line, ARPE-19. While challenges exist in having ARPE-19 cells perfectly match the
morphology and pigmentation that would have allowed us to test R:GEN in vitro, gene expression
changes during the acute phase of tissue repair and confirm modulation of RAS components occurs prior
to MMP. Using either RAS enzyme or receptor inhibitors allowed us to identify how shifts within RAS
metabolism may dictate downstream mechanisms of wound healing. Surprisingly, signals downstream
of RAS receptors indicated primarily AT2R activation when ARPE-19 cells were treated with H 2O 2.
Furthermore, while Enalapril and Losartan are both FDA approved drugs and likely used in patient
demographics that include AMD, their opposing effect on TGF- β expression in the immediate phase of
damage suggests that studies utilizing longer time points may be warranted to fully characterize the
effects that RAS modulation may have.

141
4.6 Chapter 4 References
Abdallah, W. F., Louie, S. G., Zhang, Y., Rodgers, K. E., Sivok, E., G, S. d., & Humayun, M. S. (2016).
NorLeu3A(1-7) Accelerates Clear Corneal Full Thickness Wound Healing. Invest Ophthalmol Vis
Sci, 57(4), 2187-2194. doi:10.1167/iovs.15-18515

Border, W. A., & Noble, N. A. (1998). Interactions of transforming growth factor-beta and angiotensin II
in renal fibrosis. Hypertension, 31(1 Pt 2), 181-188. doi:10.1161/01.hyp.31.1.181

Boulay, G., Servant G Fau - Luong, T. T., Luong Tt Fau - Escher, E., Escher E Fau - Guillemette, G., &
Guillemette, G. Modulation of angiotensin II binding affinity by allosteric interaction of polyvinyl
sulfate with an intracellular domain of the DuP-753-sensitive angiotensin II receptor of bovine
adrenal glomerulosa. (0026-895X (Print)).

Browatzki, M., Larsen, D., Pfeiffer, C. A., Gehrke, S. G., Schmidt, J., Kranzhofer, A., . . . Kranzhofer, R.
(2005). Angiotensin II stimulates matrix metalloproteinase secretion in human vascular smooth
muscle cells via nuclear factor-kappaB and activator protein 1 in a redox-sensitive manner. J
Vasc Res, 42(5), 415-423. doi:10.1159/000087451

Burnier, M. (2001). Angiotensin II type 1 receptor blockers. Circulation, 103(6), 904-912.
doi:10.1161/01.cir.103.6.904

Cao, X., Liu, M., Tuo, J., Shen, D., & Chan, C. C. (2010). The effects of quercetin in cultured human RPE
cells under oxidative stress and in Ccl2/Cx3cr1 double deficient mice. Exp Eye Res, 91(1), 15-25.
doi:10.1016/j.exer.2010.03.016

George, P. (1947). Reaction Between Catalase and Hydrogen Peroxide. Nature, 160(4054), 41-43.
doi:10.1038/160041a0

Gonzalez Amaya, J. A., Cabrera, D. Z., Matallana, A. M., Arevalo, K. G., & Guevara-Pulido, J. (2020). In-
silico design of new enalapril analogs (ACE inhibitors) using QSAR and molecular docking models.
Informatics in Medicine Unlocked, 19, 100336. doi:https://doi.org/10.1016/j.imu.2020.100336

Gross, D. M., Sweet, C. S., Ulm, E. H., Backlund, E. P., Morris, A. A., Weitz, D., . . . Stone, C. A. (1981).
Effect of N-[(S)-1-carboxy-3-phenylpropyl]-L-Ala-L-Pro and its ethyl ester (MK-421) on
angiotensin converting enzyme in vitro and angiotensin I pressor responses in vivo. J Pharmacol
Exp Ther, 216(3), 552-557. Retrieved from https://www.ncbi.nlm.nih.gov/pubmed/6259322

Hu, X., Wu, X., Zhao, B., & Wang, Y. (2019). Scutellarin protects human retinal pigment epithelial cells
against hydrogen peroxide (H2O2)-induced oxidative damage. Cell Biosci, 9, 12.
doi:10.1186/s13578-019-0276-0

Joshi, S., Balasubramanian, N., Vasam, G., & Jarajapu, Y. P. (2016). Angiotensin converting enzyme
versus angiotensin converting enzyme-2 selectivity of MLN-4760 and DX600 in human and
murine bone marrow-derived cells. European journal of pharmacology, 774, 25-33.
doi:10.1016/j.ejphar.2016.01.007

142
Kaczara, P., Sarna, T., & Burke, J. M. (2010). Dynamics of H2O2 availability to ARPE-19 cultures in models
of oxidative stress. Free Radic Biol Med, 48(8), 1064-1070.
doi:10.1016/j.freeradbiomed.2010.01.022

Kurz, T., Karlsson, M., Brunk, U. T., Nilsson, S. E., & Frennesson, C. (2009). ARPE-19 retinal pigment
epithelial cells are highly resistant to oxidative stress and exercise strict control over their
lysosomal redox-active iron. Autophagy, 5(4), 494-501. doi:10.4161/auto.5.4.7961

Lazzarini, R., Nicolai, M., Lucarini, G., Pirani, V., Mariotti, C., Bracci, M., & Mattioli-Belmonte, M. (2022).
Oxidative stress in retinal pigment epithelium impairs stem cells: a vicious cycle in age-related
macular degeneration. Mol Cell Biochem, 477(1), 67-77. doi:10.1007/s11010-021-04258-3

Mitra, A. K., Gao, L., & Zucker, I. H. (2010). Angiotensin II-induced upregulation of AT(1) receptor
expression: sequential activation of NF-kappaB and Elk-1 in neurons. American journal of
physiology. Cell physiology, 299(3), C561-C569. doi:10.1152/ajpcell.00127.2010

Park, W. H. (2014). Anti-apoptotic effect of caspase inhibitors on H2O2-treated HeLa cells through early
suppression of its oxidative stress. Oncol Rep, 31(5), 2413-2421. doi:10.3892/or.2014.3084

Rodgers, K., Abiko, M., Girgis, W., St Amand, K., Campeau, J., & diZerega, G. (1997). Acceleration of
dermal tissue repair by angiotensin II. Wound Repair Regen, 5(2), 175-183. doi:10.1046/j.1524-
475X.1997.50210.x

Santos, R. A., Simoes e Silva, A. C., Maric, C., Silva, D. M., Machado, R. P., de Buhr, I., . . . Walther, T.
(2003). Angiotensin-(1-7) is an endogenous ligand for the G protein-coupled receptor Mas. Proc
Natl Acad Sci U S A, 100(14), 8258-8263. doi:10.1073/pnas.1432869100

Shentu, X.-C., Ping, X.-Y., Cheng, Y.-L., Zhang, X., Tang, Y.-L., & Tang, X.-J. (2018). Hydrogen peroxide-
induced apoptosis of human lens epithelial cells is inhibited by parthenolide. International
journal of ophthalmology, 11(1), 12-17. doi:10.18240/ijo.2018.01.03

Song, Q. H., Klepeis, V. E., Nugent, M. A., & Trinkaus-Randall, V. (2002). TGF-beta1 regulates TGF-beta1
and FGF-2 mRNA expression during fibroblast wound healing. Mol Pathol, 55(3), 164-176.
doi:10.1136/mp.55.3.164

Swinney, D. C. (2011). Chapter 18 - Molecular Mechanism of Action (MMoA) in Drug Discovery. In J. E.
Macor (Ed.), Annual Reports in Medicinal Chemistry (Vol. 46, pp. 301-317): Academic Press.

Wang, T., Lian, G., Cai, X., Lin, Z., & Xie, L. (2018). Effect of prehypertensive losartan therapy on AT1R
and ATRAP methylation of adipose tissue in the later life of high-fat-fed spontaneously
hypertensive rats. Mol Med Rep, 17(1), 1753-1761. doi:10.3892/mmr.2017.8081

Xiang, J., Wan, C., Guo, R., & Guo, D. (2016). Is Hydrogen Peroxide a Suitable Apoptosis Inducer for All
Cell Types? BioMed Research International, 2016, 7343965. doi:10.1155/2016/7343965

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Chapter 5: Summary and Concluding Remarks
Age-related macular degeneration (AMD) is one of the leading causes of blindness in the United
States and the world, with nearly 45 million cases per year (Y. Wang et al., 2021). As an age-related
disease, AMD has been found to be multifactored in its progression, increasing the complexity of
identifying novel targets and developing effective treatments (van Lookeren Campagne et al., 2014).
While anti-VEGF treatments are available to manage the “wet” form of AMD, approximately 80% to 85%
of AMD patients suffer from the “dry” form of the disease, for which an FDA-approved therapeutic does
not yet exist and signals a significant unmet need (Ambati & Fowler, 2012). The development of
therapeutics for dAMD is currently centered around drusen, the clinical marker of AMD, for which
increase in size of these lipoprotein deposits indicate disease progression. As such, drusen regression is
believed to slow/treat the disease.
This dissertation investigate and characterize the molecular response after R:GEN, a selective
retinal therapy (SRT) laser, ocular application as a potential therapeutic modality for dAMD.
Photocoagulation, a form of laser treatment previously tested for AMD where its use was found reduce
drusen deposits, but also commonly resulted in fibrosis in the eye, negating its usefulness for this
application (Kotoula et al., 2009). In comparison, SRT, uses pulses of light energy to form microbubbles
within cells and burst them without causing excessive damage to the surrounding tissue, avoiding
adverse events in the eyes of AMD patients (Theodore Smith, 2019).
Outside of its prominent role in regulating blood pressure, the RAS has been shown to play a
role in the pathogenesis of atherosclerosis and promotion of wound healing. Prior research has shown
the “pathogenic arm” of RAS or the Ang II/AT1R axis to promote plaque formation and promote pro-
fibrotic wound healing (Bargagli et al., 2020; Silva et al., 2020). However, a “protective arm” of RAS also
exists for which include the following components Ang II/AT2R and Ang(1-7)/MasR. The activation of
Protective Arm of RAS can counterbalance the activity of the pathogenic arm, reducing plaque formation

144
and promoting anti-fibrotic activity. The mounting evidence that RAS can mitigate plaque formation and
fibrotic wound healing suggested the efficacy of SRT might be tied to this pathway. After a review of the
literature, virtually no studies have been conducted to determine the effect of lasers on ocular RAS and
how modulation of this pathway impact drusen regression and disease pathogenesis of dry AMD. To
address this gap in the knowledge and building upon findings not yet published by our research group, I
aimed to determine the implications of RAS modulation in the eyes of AMD mouse models.
In Chapter 2, I have investigated the effects of R:GEN treatment on the eyes of Cx3Cr1-knockout
mice, a hard drusen mouse model. The parameters of laser treatment were determined from our pilot
studies in C57/BL, for which application of 100% intensity R:GEN laser was deemed safe and sufficient
for activation of RAS enzymes and receptors. The safety of treatment was assessed by utilizing fundus
imaging, fluorescein angiography, optical coherence tomography (OCT), and immunohistochemistry
techniques. RPE lesions from R:GEN were shown close by day 7 of treatment, as leakage of fluorescein
could no longer be detected. Hematoxylin and eosin (H&E) sections underscored the eye’s ability to heal
the laser-induced lesions as pigment filled in the void induced by R:GEN treatment. Furthermore, H&E
showed that R:GEN did not affect the photoreceptor layer of the eye, highlighting the safety and
selectivity of the laser. Fundus autofluorescence (FAF) was utilized to determine the presence and
regression of drusen-like deposits in the Cx3Cr1 -/- mice, and while not found to be statistically
significant, did show a decreasing trend in the number of fluorescent spots detected, indicating that
R:GEN was capable of regressing drusen in this model (Luhmann et al., 2009). To probe the effect of
R:GEN on local RAS activity, ocular fluids were collected at the end of each time point and analyzed for
the presence of peptides using multiplexed liquid chromatography-mass spectrometry (LC-MS) method
that I had optimized and validated. Utilizing a metabolomics approach, I found evidence that R:GEN
would activate the protective arm of RAS, where the peptide ratio of Ang(1-7)/Ang II was indicative of
high ACE2 activity (Reddy et al., 2019). Furthermore, using RT-PCR, I found that there was an

145
upregulation of Mas receptor expression. In the presence of Ang(1-7), this can activate the protective
arm of RAS. The ability to activate the protective arm of RAS highlights how R:GEN laser therapy can
activate molecular mechanisms that promotes minimally fibrotic healing.
Correlation analysis between the change in FAF counts and RAS components, revealed negative
correlations between FAF counts and RAS components such as Ang(1-7), MasR, and the Ang(1-9)/Ang(1-
10) ratio. Overall, these findings support the hypothesis that R:GEN could activate beneficial for dAMD,
where its application can promote the regression of the drusen-like deposits through activating the
protective arm of RAS. However, the unexpected downregulation of ACE, ACE2, and AT2R immediately
after treatment suggests that when R:GEN is used at the 100% intensity level, this stimulation level may
be excessive for RPE and trigger a shut down of some components of RAS, setting the stage for a
reduction in dose in chapter 3.
Building on the findings from Chapter 2, we repeated our comprehensive analysis of R:GEN,
used a soft drusen mouse model (Cxcr5-/-) where treatment strategies used reduced laser intensity
(e.g., 50% and 100% intensity). Similar to Chapter 2, we verified that R:GEN selectively targeted the RPE,
as shown in the H&E slides, where disruption of the RPE was visible immediately after treatment while
the photoreceptors remained undisturbed. These results underscored the safety of R:GEN, even when
used in a soft drusen model. Again, FA showed the RPE could heal from R:GEN induced lesions from
R:GEN by Day 7 of treatment, for which fluorescein leakage was undetected. While soft drusen is a
greater risk factor for progression of AMD, the lack of distinct borders in the deposits of our Cxcr5-/-
were a severe limitation for FAF analysis, leaving us to rely on the correlations identified in the previous
animal study as a proxy of efficacy (Nita et al., 2014). Again, we observed evidence for activation of the
protective arm of RAS, where Ang(1-9), Ang(1-7) AT2R, and the ratio of Ang(1-7)/Ang II were found to be
upregulated after treatment, at early timepoints. Interestingly, we found that gene expression of MasR
was varied with laser intensity, supporting our decision to include an additional treatment arm with less

146
energy. We see the 100% intensity arm led to a pronounced immediate upregulation of MasR in the RPE
followed by downregulation, whereas 50% intensity treatment led to relatively lower but more
sustained upregulation of MasR. By definition, 50% intensity is the amount of energy required to induce
positive FA. This suggests that subthreshold activation may be all that is necessary to induce molecular
events that will promote ocular healing. Furthermore, I identified a dose-response relationship between
laser intensity and ocular peptide production by calculating the area under the curve (AUC) in the acute
phase of wound healing, which is defined as the first three timepoints (e.g., 0.04, and 0.16 and 3 day).
The AUC for Ang(1-7) and Ang(1-5) ocular concentrations were found to be greater at 50% in our soft
drusen model, as well as the metabolomic peptide ratios indicative of ACE and ACE2 activity. The work
conducted in this chapter confirms that R:GEN is safe in the eyes of our soft drusen model, in alignment
with our healthy normal mice and hard drusen mice. In addition, we were able to confirm our
hypothesis that R:GEN was able to stimulate the protective arm of RAS, increasing the production and
activity of RAS components identified to correlate with the drusen regression of chapter 2. As expected
from the hard drusen mouse study, the evidence suggests that a lower intensity of 50% resulted in a
greater effect in our desired molecular signals in our disease models. Unexpectedly, ACE2 was still
downregulated in the RPE after treatment, suggesting a potential alternative pathway engaging in the
metabolism RAS peptides and increasing the production of downstream metabolites, like Ang(1-7).
In Chapter 4, I devised an in vitro model using ARPE-19 to mimic R:GEN-induced RPE to further
probe the role of RAS in ocular wound healing. Using inhibitors of RAS enzymes and receptors, I
dissected and elucidate the molecular mechanism of immediate tissue damage on RAS metabolism. Cells
were pre-treated with inhibitors for 30 minutes to completely inhibit their targets prior to H 2O 2
treatment, a method previously used to induce damage to ARPE-19 cells for studying damage and
oxidative stress (Kaczara et al., 2010). Gene expression and extracellular metabolomic changes were
analyzed to provide additional context to our conducted animal studies. As expected, upregulation of

147
ACE and ACE2 occurred with H 2O 2. Interestingly, H 2O 2 induced upregulation of AT2R, similar to what was
observed in soft drusen mice, but simultaneously downregulated AT1R, similar to the effect of R:GEN in
our hard drusen model. Gene expression of ARPE-19 cells with inhibitors were in line with expectations
from literature (Costerousse, Allegrini, Clozel, Menard, & Alhenc-Gelas, 1998). However, while the use of
inhibitors in combination with H 2O 2 induced changes within peptide concentrations when compared to
H 2O 2 alone, metabolomics analysis indicated that only Losartan, an angiotensin receptor blocker (ARB),
and MLN4760, an ACE2 inhibitor had any statistically significant differences. ACE inhibitors and ARBs are
FDA-approved therapeutics commonly used in older populations, previously reported to be 38.7% of
individuals 65 and older (Murphy, Drawz, & Foley, 2019). Because ACEi/ARB users are in an age group
that overlaps with those of AMD populations, the observed modulations in RAS indicate that further
study with combination ACEi/ARB and R:GEN may be warranted. Specifically, Enalapril appeared to
augment the production of Ang(1-9) with H 2O 2, while Losartan was able to increase the production of
Ang(1-9) and Ang(1-7), potentially shunting the RPE towards activation of the protective arm of RAS
after tissue damage.
Finally, our previous animal studies all showed an initial upregulation of TGF- β, a normal
response as part of wound healing but known to cause fibrosis when continuously upregulated
(Eickelberg, 2001; Meng et al., 2016). Because Ang II is known to upregulate TGF- β as part of the Ang
II/AT1R pathogenic arm of RAS, it was critical to determine whether R:GEN stimulation of RAS peptide
production would potentially drive fibrosis (Daniel, 2008). To mitigate these concerns, I incubated ARPE-
19 cells with escalating doses of Ang II from 1 nM to 1 µM, for which the highest dose was found to
upregulate TGF- β gene expression. In comparison, when the Ang II concentrations measured in the eye
were converted to molar, we observed these concentrations were significantly lower than 1 nM, even
though TGF- β was upregulated. These results would suggest that TGF- β expression observed in our
R:GEN animal studies are from the normal tissue remodeling response and unlikely to come from RAS

148
modulation, once again emphasizing the safety of R:GEN as it is unlikely to cause fibrosis commonly
observed in photocoagulation (Guyer, D'Amico, & Smith, 1992; Lovestam-Adrian & Agardh, 2000).
In the presented studies, I have shown selective retinal therapy, and more specifically the R:GEN
laser to be safe for use in the eye. Through a combination of imaging and bioanalytical techniques, I was
able to find correlations between drusen regression in our hard drusen mouse model and components
of the Renin-Angiotensin System, highlighting the possibility that this pathway may play a critical role in
slowing disease progression. I also identified a dose-response relationship between R:GEN intensity and
RAS signals in our soft drusen mouse model, indicating the importance of refining the treatment “dose”
when translating to clinical use, such that efficacy can be maximized for disease patients as their
response may vary from that of healthy controls. Lastly, I evaluated how inhibition of RAS enzymes and
receptors may affect the wound healing response immediately after damage.
To build upon the work presented in this dissertation, patients who are undergoing selective
retinal therapy with R:GEN may benefit from the addition of ACEi or ARB to tilt the balance towards the
protective arm of RAS. To further understand this combination therapy, a non-rodent model, like a
drusen-forming monkey model may needed since the larger eyes may mitigate some of the extraction
challenges that occur with mice (Edwards & Malek, 2007). Lastly, additional techniques, such MALDI-
TOF imaging for proteins known to be present in drusen may be considered to better determine the
efficacy of R:GEN treatment (Farkas, Sylvester, Archer, & Altona, 1971; Johnson, Ozaki, Staples, Erickson,
& Anderson, 2000).

149
References
Abdallah, W. F., Louie, S. G., Zhang, Y., Rodgers, K. E., Sivok, E., G, S. d., & Humayun, M. S.
(2016). NorLeu3A(1-7) Accelerates Clear Corneal Full Thickness Wound Healing. Invest
Ophthalmol Vis Sci, 57(4), 2187-2194. doi:10.1167/iovs.15-18515

Allen, T. J., & Jandeleit-Dahm, K. A. (2005). Preventing atherosclerosis with angiotensin-
converting enzyme inhibitors: emphasis on diabetic atherosclerosis. Curr Drug Targets
Cardiovasc Haematol Disord, 5(6), 503-512. doi:10.2174/156800605774962068

AlQudah, M., Hale, T. M., & Czubryt, M. P. (2020). Targeting the renin-angiotensin-aldosterone
system in fibrosis. Matrix Biol, 91-92, 92-108. doi:10.1016/j.matbio.2020.04.005

Amaroli, A., Pasquale, C., Zekiy, A., Utyuzh, A., Benedicenti, S., Signore, A., & Ravera, S. (2021).
Photobiomodulation and Oxidative Stress: 980 nm Diode Laser Light Regulates
Mitochondrial Activity and Reactive Oxygen Species Production. Oxid Med Cell Longev,
2021, 6626286. doi:10.1155/2021/6626286

Ambati, J., & Fowler, B. J. (2012). Mechanisms of age-related macular degeneration. Neuron,
75(1), 26-39. doi:10.1016/j.neuron.2012.06.018

Aono, J., Suzuki, J., Iwai, M., Horiuchi, M., Nagai, T., Nishimura, K., . . . Higaki, J. (2012). Deletion
of the angiotensin II type 1a receptor prevents atherosclerotic plaque rupture in
apolipoprotein E-/- mice. Arterioscler Thromb Vasc Biol, 32(6), 1453-1459.
doi:10.1161/ATVBAHA.112.249516

Baradaran-Rafii, A., Sarvari, M., Alavi-Moghadam, S., Payab, M., Goodarzi, P., Aghayan, H. R., . . .
Arjmand, B. (2020). Cell-based approaches towards treating age-related macular
degeneration. Cell Tissue Bank. doi:10.1007/s10561-020-09826-3

Bargagli, E., Refini, R. M., d'Alessandro, M., Bergantini, L., Cameli, P., Vantaggiato, L., . . . Landi,
C. (2020). Metabolic Dysregulation in Idiopathic Pulmonary Fibrosis. Int J Mol Sci, 21(16).
doi:10.3390/ijms21165663

Barzilai, N., Crandall, J. P., Kritchevsky, S. B., & Espeland, M. A. (2016). Metformin as a Tool to
Target Aging. Cell Metab, 23(6), 1060-1065. doi:10.1016/j.cmet.2016.05.011

Bergandi, L., Skorokhod, O. A., Franzone, F., La Grotta, R., Schwarzer, E., & Nuzzi, R. (2018).
Induction of oxidative stress in human aqueous and vitreous humors by Nd:YAG laser
posterior capsulotomy. Int J Ophthalmol, 11(7), 1145-1151. doi:10.18240/ijo.2018.07.12

150
Border, W. A., & Noble, N. A. (1998). Interactions of transforming growth factor-beta and
angiotensin II in renal fibrosis. Hypertension, 31(1 Pt 2), 181-188.
doi:10.1161/01.hyp.31.1.181

Boulay, G., Servant G Fau - Luong, T. T., Luong Tt Fau - Escher, E., Escher E Fau - Guillemette, G.,
& Guillemette, G. Modulation of angiotensin II binding affinity by allosteric interaction
of polyvinyl sulfate with an intracellular domain of the DuP-753-sensitive angiotensin II
receptor of bovine adrenal glomerulosa. (0026-895X (Print)).

Branton, M. H., & Kopp, J. B. (1999). TGF-beta and fibrosis. Microbes Infect, 1(15), 1349-1365.
doi:10.1016/s1286-4579(99)00250-6

Browatzki, M., Larsen, D., Pfeiffer, C. A., Gehrke, S. G., Schmidt, J., Kranzhofer, A., . . .
Kranzhofer, R. (2005). Angiotensin II stimulates matrix metalloproteinase secretion in
human vascular smooth muscle cells via nuclear factor-kappaB and activator protein 1 in
a redox-sensitive manner. J Vasc Res, 42(5), 415-423. doi:10.1159/000087451

Burnier, M. (2001). Angiotensin II type 1 receptor blockers. Circulation, 103(6), 904-912.
doi:10.1161/01.cir.103.6.904

Burns, R. P., & Feeney-Burns, L. (1980). Clinico-morphologic correlations of drusen of Bruch's
membrane. Trans Am Ophthalmol Soc, 78, 206-225.

Buschini, E., Fea, A. M., Lavia, C. A., Nassisi, M., Pignata, G., Zola, M., & Grignolo, F. M. (2015).
Recent developments in the management of dry age-related macular degeneration. Clin
Ophthalmol, 9, 563-574. doi:10.2147/OPTH.S59724

Caley, M. P., Martins, V. L., & O'Toole, E. A. (2015). Metalloproteinases and Wound Healing. Adv
Wound Care (New Rochelle), 4(4), 225-234. doi:10.1089/wound.2014.0581

Caligiuri, G., Nicoletti, A., Poirier, B., & Hansson, G. K. (2002). Protective immunity against
atherosclerosis carried by B cells of hypercholesterolemic mice. The Journal of clinical
investigation, 109(6), 745-753. doi:10.1172/JCI7272

Cao, X., Liu, M., Tuo, J., Shen, D., & Chan, C. C. (2010). The effects of quercetin in cultured
human RPE cells under oxidative stress and in Ccl2/Cx3cr1 double deficient mice. Exp
Eye Res, 91(1), 15-25. doi:10.1016/j.exer.2010.03.016

Chiu, A. T., Herblin, W. F., McCall, D. E., Ardecky, R. J., Carini, D. J., Duncia, J. V., . . .
Timmermans, P. B. M. W. M. (1989). Identification of angiotensin II receptor subtypes.
Biochemical and Biophysical Research Communications, 165(1), 196-203.
doi:https://doi.org/10.1016/0006-291X(89)91054-1

151
Costerousse, O., Allegrini, J., Clozel, J. P., Menard, J., & Alhenc-Gelas, F. (1998). Angiotensin I-
converting enzyme inhibition but not angiotensin II suppression alters angiotensin I-
converting enzyme gene expression in vessels and epithelia. J Pharmacol Exp Ther,
284(3), 1180-1187. Retrieved from https://www.ncbi.nlm.nih.gov/pubmed/9495881

Crabb, J. W., Miyagi, M., Gu, X., Shadrach, K., West, K. A., Sakaguchi, H., . . . Hollyfield, J. G.
(2002). Drusen proteome analysis: an approach to the etiology of age-related macular
degeneration. Proc Natl Acad Sci U S A, 99(23), 14682-14687.
doi:10.1073/pnas.222551899

Crotty, S. (2011). Follicular Helper CD4 T Cells (TFH). Annual Review of Immunology, 29(1), 621-
663. doi:10.1146/annurev-immunol-031210-101400

Curcio, C. A. (2018). Soft Drusen in Age-Related Macular Degeneration: Biology and Targeting
Via the Oil Spill Strategies. Invest Ophthalmol Vis Sci, 59(4), AMD160-AMD181.
doi:10.1167/iovs.18-24882

Curcio, C. A., Johnson, M., Huang, J. D., & Rudolf, M. (2010). Apolipoprotein B-containing
lipoproteins in retinal aging and age-related macular degeneration. J Lipid Res, 51(3),
451-467. doi:10.1194/jlr.R002238

Curcio, C. A., Millican, C. L., Bailey, T., & Kruth, H. S. (2001). Accumulation of cholesterol with
age in human Bruch's membrane. Invest Ophthalmol Vis Sci, 42(1), 265-274.

Daniel, C. (2008). Blocking of angiotensin II is more than blocking of transforming growth factor-
beta. Kidney Int, 74(5), 551-553. doi:10.1038/ki.2008.290

Danser, A. H., Derkx Fh Fau - Admiraal, P. J., Admiraal Pj Fau - Deinum, J., Deinum J Fau - de
Jong, P. T., de Jong Pt Fau - Schalekamp, M. A., & Schalekamp, M. A. (1994). Angiotensin
levels in the eye. (0146-0404 (Print)).

Dolman, C. L., & MacLeod, P. M. (1981). Lipofuscin and its Relation to Aging. In S. Fedoroff & L.
Hertz (Eds.), Advances in Cellular Neurobiology (Vol. 2, pp. 205-247): Elsevier.

Donoso, L. A., Kim, D., Frost, A., Callahan, A., & Hageman, G. (2006). The role of inflammation in
the pathogenesis of age-related macular degeneration. Surv Ophthalmol, 51(2), 137-
152. doi:10.1016/j.survophthal.2005.12.001

Edwards, A. O., & Malek, G. (2007). Molecular genetics of AMD and current animal models.
Angiogenesis, 10(2), 119-132. doi:10.1007/s10456-007-9064-2

Eickelberg, O. (2001). Endless healing: TGF-beta, SMADs, and fibrosis. FEBS Lett, 506(1), 11-14.
doi:10.1016/s0014-5793(01)02875-7

152
Epstein, I. J., Aziz, H. A., Young, R. C., & Berrocal, A. M. (2013). Macular hard exudates and scar
formation after laser photocoagulation in retinopathy of prematurity. J Pediatr
Ophthalmol Strabismus, 50 Online, e30-32. doi:10.3928/01913913-20130625-01

Fang, J., Yang, J., Wu, X., Zhang, G., Li, T., Wang, X., . . . Wang, L. (2018). Metformin alleviates
human cellular aging by upregulating the endoplasmic reticulum glutathione peroxidase
7. Aging Cell, 17(4), e12765. doi:10.1111/acel.12765

Farkas, T. G., Sylvester, V., Archer, D., & Altona, M. (1971). The histochemistry of drusen. Am J
Ophthalmol, 71(6), 1206-1215. doi:10.1016/0002-9394(71)90964-0

Fernandez-Godino, R., Garland, D. L., & Pierce, E. A. (2016). Isolation, culture and
characterization of primary mouse RPE cells. Nat Protoc, 11(7), 1206-1218.
doi:10.1038/nprot.2016.065

Ferris, F. L., Davis, M. D., Clemons, T. E., Lee, L. Y., Chew, E. Y., Lindblad, A. S., . . . Age-Related
Eye Disease Study Research, G. (2005). A simplified severity scale for age-related
macular degeneration: AREDS Report No. 18. Arch Ophthalmol, 123(11), 1570-1574.
doi:10.1001/archopht.123.11.1570

Figueroa, M. S., Regueras, A., & Bertrand, J. (1994). Laser photocoagulation to treat macular
soft drusen in age-related macular degeneration. Retina, 14(5), 391-396.
doi:10.1097/00006982-199414050-00001

Frederick, P. A., & Kleinman, M. E. (2014). The Immune System and AMD. Curr Ophthalmol Rep,
2(1), 14-19. doi:10.1007/s40135-013-0037-x

Gehrs, K. M., Anderson, D. H., Johnson, L. V., & Hageman, G. S. (2006). Age-related macular
degeneration--emerging pathogenetic and therapeutic concepts. Ann Med, 38(7), 450-
471. doi:10.1080/07853890600946724

George, P. (1947). Reaction Between Catalase and Hydrogen Peroxide. Nature, 160(4054), 41-
43. doi:10.1038/160041a0

Girmens, J. F., Sahel, J. A., & Marazova, K. (2012). Dry age-related macular degeneration: A
currently unmet clinical need. Intractable Rare Dis Res, 1(3), 103-114.
doi:10.5582/irdr.2012.v1.3.103

Gonzalez Amaya, J. A., Cabrera, D. Z., Matallana, A. M., Arevalo, K. G., & Guevara-Pulido, J.
(2020). In-silico design of new enalapril analogs (ACE inhibitors) using QSAR and
molecular docking models. Informatics in Medicine Unlocked, 19, 100336.
doi:https://doi.org/10.1016/j.imu.2020.100336

153
Gross, D. M., Sweet, C. S., Ulm, E. H., Backlund, E. P., Morris, A. A., Weitz, D., . . . Stone, C. A.
(1981). Effect of N-[(S)-1-carboxy-3-phenylpropyl]-L-Ala-L-Pro and its ethyl ester (MK-
421) on angiotensin converting enzyme in vitro and angiotensin I pressor responses in
vivo. J Pharmacol Exp Ther, 216(3), 552-557. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/6259322

Gunn, M. D., Ngo, V. N., Ansel, K. M., Ekland, E. H., Cyster, J. G., & Williams, L. T. (1998). A B-
cell-homing chemokine made in lymphoid follicles activates Burkitt's lymphoma
receptor-1. Nature, 391(6669), 799-803. doi:10.1038/35876

Guo, L., Hussain, A. A., Limb, G. A., & Marshall, J. (1999). Age-dependent variation in
metalloproteinase activity of isolated human Bruch's membrane and choroid. Invest
Ophthalmol Vis Sci, 40(11), 2676-2682. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/10509665

Guttenplan, K., Blum, J., & Bennett, M. (2018). A Role for Microglia in Retinal Development. J
Neurosci, 38(43), 9126-9128. doi:10.1523/JNEUROSCI.1813-18.2018

Guyer, D. R., D'Amico, D. J., & Smith, C. W. (1992). Subretinal fibrosis after laser
photocoagulation for diabetic macular edema. Am J Ophthalmol, 113(6), 652-656.
doi:10.1016/s0002-9394(14)74789-0

Guymer, R. H., Wu, Z., Hodgson, L. A. B., Caruso, E., Brassington, K. H., Tindill, N., . . . Laser
Intervention in Early Stages of Age-Related Macular Degeneration Study, G. (2019).
Subthreshold Nanosecond Laser Intervention in Age-Related Macular Degeneration: The
LEAD Randomized Controlled Clinical Trial. Ophthalmology, 126(6), 829-838.
doi:10.1016/j.ophtha.2018.09.015

H. J. Motulsky. GraphPad Statistics Guide. Retrieved from
http://www.graphpad.com/guides/prism/7/statistics/index.htm

Hampe, C. S. (2012). B Cell in Autoimmune Diseases. Scientifica, 2012, 215308.
doi:10.6064/2012/215308

Harman, D. (1956). Aging: a theory based on free radical and radiation chemistry. J Gerontol,
11(3), 298-300. doi:10.1093/geronj/11.3.298

Hector, S. M., & Sørensen, T. L. (2017). Circulating monocytes and B-lymphocytes in neovascular
age-related macular degeneration. Clinical ophthalmology (Auckland, N.Z.), 11, 179-184.
doi:10.2147/OPTH.S121332

Hu, X., Wu, X., Zhao, B., & Wang, Y. (2019). Scutellarin protects human retinal pigment
epithelial cells against hydrogen peroxide (H2O2)-induced oxidative damage. Cell Biosci,
9, 12. doi:10.1186/s13578-019-0276-0

154
Huang, H., Liu, Y., Wang, L., & Li, W. (2017). Age-related macular degeneration phenotypes are
associated with increased tumor necrosis-alpha and subretinal immune cells in aged
Cxcr5 knockout mice. PLoS One, 12(3), e0173716. doi:10.1371/journal.pone.0173716

Jadhav, S. S., Sharma, N., Meeks, C. J., Mordwinkin, N. M., Espinoza, T. B., Roda, N. R., . . .
Rodgers, K. E. (2013). Effects of combined radiation and burn injury on the renin-
angiotensin system. Wound Repair Regen, 21(1), 131-140. doi:10.1111/j.1524-
475X.2012.00867.x

Janeway CA Jr, T. P., Walport M, et al. (2001). The complement system and innate immunity. In
G. Science (Ed.), Immunobiology: The Immune System in Health and Disease. 5th edition.
New York.

Jobling, A. I., Guymer, R. H., Vessey, K. A., Greferath, U., Mills, S. A., Brassington, K. H., . . .
Fletcher, E. L. (2015). Nanosecond laser therapy reverses pathologic and molecular
changes in age-related macular degeneration without retinal damage. FASEB J, 29(2),
696-710. doi:10.1096/fj.14-262444

Jobling, A. I., Waugh, M., Vessey, K. A., Phipps, J. A., Trogrlic, L., Greferath, U., . . . Fletcher, E. L.
(2018). The Role of the Microglial Cx3cr1 Pathway in the Postnatal Maturation of Retinal
Photoreceptors. J Neurosci, 38(20), 4708-4723. doi:10.1523/JNEUROSCI.2368-17.2018

Johnson, L. V., Ozaki, S., Staples, M. K., Erickson, P. A., & Anderson, D. H. (2000). A potential role
for immune complex pathogenesis in drusen formation. Exp Eye Res, 70(4), 441-449.
doi:10.1006/exer.1999.0798

Johnstone, M. T., Perez, A. S., Nasser, I., Stewart, R., Vaidya, A., Al Ammary, F., . . . Quist, W. C.
(2004). Angiotensin receptor blockade with candesartan attenuates atherosclerosis,
plaque disruption, and macrophage accumulation within the plaque in a rabbit model.
Circulation, 110(14), 2060-2065. doi:10.1161/01.CIR.0000143627.55926.4C

Joshi, S., Balasubramanian, N., Vasam, G., & Jarajapu, Y. P. (2016). Angiotensin converting
enzyme versus angiotensin converting enzyme-2 selectivity of MLN-4760 and DX600 in
human and murine bone marrow-derived cells. European journal of pharmacology, 774,
25-33. doi:10.1016/j.ejphar.2016.01.007

Kaczara, P., Sarna, T., & Burke, J. M. (2010). Dynamics of H2O2 availability to ARPE-19 cultures
in models of oxidative stress. Free Radic Biol Med, 48(8), 1064-1070.
doi:10.1016/j.freeradbiomed.2010.01.022

Kaemmerer, E., Schutt, F., Krohne, T. U., Holz, F. G., & Kopitz, J. (2007). Effects of lipid
peroxidation-related protein modifications on RPE lysosomal functions and POS
phagocytosis. Invest Ophthalmol Vis Sci, 48(3), 1342-1347. doi:10.1167/iovs.06-0549

155
Karwatowski, W. S., Jeffries, T. E., Duance, V. C., Albon, J., Bailey, A. J., & Easty, D. L. (1995).
Preparation of Bruch's membrane and analysis of the age-related changes in the
structural collagens. Br J Ophthalmol, 79(10), 944-952. doi:10.1136/bjo.79.10.944

Kim, J., Lee, Y. J., & Won, J. Y. (2021). Molecular Mechanisms of Retinal Pigment Epithelium
Dysfunction in Age-Related Macular Degeneration. Int J Mol Sci, 22(22).
doi:10.3390/ijms222212298

Klein, R., Myers, C. E., Lee, K. E., Gangnon, R. E., Sivakumaran, T. A., Iyengar, S. K., & Klein, B. E.
(2015). Small Drusen and Age-Related Macular Degeneration: The Beaver Dam Eye
Study. J Clin Med, 4(3), 424-440. doi:10.3390/jcm4030425

Kon, V., & Jabs, K. (2004). Angiotensin in atherosclerosis. Curr Opin Nephrol Hypertens, 13(3),
291-297. doi:10.1097/00041552-200405000-00005

Kotoula, M. G., Chatzoulis, D. Z., Tahmitzi, S., & Tsironi, E. E. (2009). Detection of macular
photocoagulation scars with confocal infrared reflection imaging. Ophthalmic Surg
Lasers Imaging, 40(4), 385-388. doi:10.3928/15428877-20096030-06

Küppers, R. (2005). Mechanisms of B-cell lymphoma pathogenesis. Nature Reviews Cancer, 5(4),
251-262. doi:10.1038/nrc1589

Kurz, T., Karlsson, M., Brunk, U. T., Nilsson, S. E., & Frennesson, C. (2009). ARPE-19 retinal
pigment epithelial cells are highly resistant to oxidative stress and exercise strict control
over their lysosomal redox-active iron. Autophagy, 5(4), 494-501.
doi:10.4161/auto.5.4.7961

Landsman, L., Bar-On, L., Zernecke, A., Kim, K. W., Krauthgamer, R., Shagdarsuren, E., . . . Jung,
S. (2009). CX3CR1 is required for monocyte homeostasis and atherogenesis by
promoting cell survival. Blood, 113(4), 963-972. doi:10.1182/blood-2008-07-170787

Lazzarini, R., Nicolai, M., Lucarini, G., Pirani, V., Mariotti, C., Bracci, M., & Mattioli-Belmonte, M.
(2022). Oxidative stress in retinal pigment epithelium impairs stem cells: a vicious cycle
in age-related macular degeneration. Mol Cell Biochem, 477(1), 67-77.
doi:10.1007/s11010-021-04258-3

Lee, M., Lee, Y., Song, J., Lee, J., & Chang, S. Y. (2018). Tissue-specific Role of CX3CR1 Expressing
Immune Cells and Their Relationships with Human Disease. Immune Netw, 18(1), e5.
doi:10.4110/in.2018.18.e5

Lennikov, A., Mukwaya, A., Saddala, M. S., & Huang, H. (2021). Deficiency of C-X-C chemokine
receptor type 5 (CXCR5) gene causes dysfunction of retinal pigment epithelium cells. Lab
Invest, 101(2), 228-244. doi:10.1038/s41374-020-00491-4

156
Li, M., Dolz-Marco, R., Messinger, J. D., Wang, L., Feist, R. M., Girkin, C. A., . . . Freund, K. B.
(2018). Clinicopathologic Correlation of Anti-Vascular Endothelial Growth Factor-Treated
Type 3 Neovascularization in Age-Related Macular Degeneration. Ophthalmology,
125(2), 276-287. doi:10.1016/j.ophtha.2017.08.019

Lovestam-Adrian, M., & Agardh, E. (2000). Photocoagulation of diabetic macular oedema--
complications and visual outcome. Acta Ophthalmol Scand, 78(6), 667-671.
doi:10.1034/j.1600-0420.2000.078006667.x

Luhmann, U. F., Robbie, S., Munro, P. M., Barker, S. E., Duran, Y., Luong, V., . . . MacLaren, R. E.
(2009). The drusenlike phenotype in aging Ccl2-knockout mice is caused by an
accelerated accumulation of swollen autofluorescent subretinal macrophages. Invest
Ophthalmol Vis Sci, 50(12), 5934-5943. doi:10.1167/iovs.09-3462

Ly, A., Nivison-Smith, L., Assaad, N., & Kalloniatis, M. (2017). Fundus Autofluorescence in Age-
related Macular Degeneration. Optometry and vision science : official publication of the
American Academy of Optometry, 94(2), 246-259. doi:10.1097/OPX.0000000000000997

Ma, B., Dang, G., Yang, S., Duan, L., & Zhang, Y. (2015). CX3CR1 polymorphisms and the risk of
age-related macular degeneration. Int J Clin Exp Pathol, 8(8), 9592-9596. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/26464724

Maeshima, K., Utsugi-Sutoh, N., Otani, T., & Kishi, S. (2004). Progressive enlargement of
scattered photocoagulation scars in diabetic retinopathy. Retina, 24(4), 507-511.
doi:10.1097/00006982-200408000-00002

Major, A. S., Fazio, S., & Linton, M. F. (2002). B-Lymphocyte Deficiency Increases Atherosclerosis
in LDL Receptor–Null Mice. Arteriosclerosis, Thrombosis, and Vascular Biology, 22(11),
1892-1898. doi:10.1161/01.ATV.0000039169.47943.EE

Martin, P. (1997). Wound healing--aiming for perfect skin regeneration. Science, 276(5309), 75-
81. doi:10.1126/science.276.5309.75

Martins, V. L., Caley, M., & O'Toole, E. A. (2013). Matrix metalloproteinases and epidermal
wound repair. Cell Tissue Res, 351(2), 255-268. doi:10.1007/s00441-012-1410-z

Meghraoui-Kheddar, A., Barthelemy, S., Boissonnas, A., & Combadiere, C. (2020). Revising
CX3CR1 Expression on Murine Classical and Non-classical Monocytes. Front Immunol,
11, 1117. doi:10.3389/fimmu.2020.01117

Mehta, P. K., & Griendling, K. K. (2007). Angiotensin II cell signaling: physiological and
pathological effects in the cardiovascular system. Am J Physiol Cell Physiol, 292(1), C82-
97. doi:10.1152/ajpcell.00287.2006

157
Meng, X. M., Nikolic-Paterson, D. J., & Lan, H. Y. (2016). TGF-beta: the master regulator of
fibrosis. Nat Rev Nephrol, 12(6), 325-338. doi:10.1038/nrneph.2016.48

Mitra, A. K., Gao, L., & Zucker, I. H. (2010). Angiotensin II-induced upregulation of AT(1)
receptor expression: sequential activation of NF-kappaB and Elk-1 in neurons. American
journal of physiology. Cell physiology, 299(3), C561-C569.
doi:10.1152/ajpcell.00127.2010

Murphy, D. P., Drawz, P. E., & Foley, R. N. (2019). Trends in Angiotensin-Converting Enzyme
Inhibitor and Angiotensin II Receptor Blocker Use among Those with Impaired Kidney
Function in the United States. Journal of the American Society of Nephrology : JASN,
30(7), 1314-1321. doi:10.1681/ASN.2018100971

Nagai, H., & Kalnins, V. I. (1996). Normally occurring loss of single cells and repair of resulting
defects in retinal pigment epithelium in situ. Exp Eye Res, 62(1), 55-61.
doi:10.1006/exer.1996.0007

Neumann, J., & Brinkmann, R. (2005). Boiling nucleation on melanosomes and microbeads
transiently heated by nanosecond and microsecond laser pulses. J Biomed Opt, 10(2),
024001. doi:10.1117/1.1896969

Nita, M., Grzybowski, A., Ascaso, F. J., & Huerva, V. (2014). Age-related macular degeneration in
the aspect of chronic low-grade inflammation (pathophysiological parainflammation).
Mediators Inflamm, 2014, 930671. doi:10.1155/2014/930671

Park, W. H. (2014). Anti-apoptotic effect of caspase inhibitors on H2O2-treated HeLa cells
through early suppression of its oxidative stress. Oncol Rep, 31(5), 2413-2421.
doi:10.3892/or.2014.3084

Patel, N., Ohbayashi, M., Nugent, A. K., Ramchand, K., Toda, M., Chau, K.-Y., . . . Chong, V.
(2005). Circulating anti-retinal antibodies as immune markers in age-related macular
degeneration. Immunology, 115(3), 422-430. doi:10.1111/j.1365-2567.2005.02173.x

Phipps, J. A., Vessey, K. A., Brandli, A., Nag, N., Tran, M. X., Jobling, A. I., & Fletcher, E. L. (2018).
The Role of Angiotensin II/AT1 Receptor Signaling in Regulating Retinal Microglial
Activation. Invest Ophthalmol Vis Sci, 59(1), 487-498. doi:10.1167/iovs.17-22416

Qin, S., Dong, N., Yang, M., Wang, J., Feng, X., & Wang, Y. (2021). Complement Inhibitors in Age-
Related Macular Degeneration: A Potential Therapeutic Option. J Immunol Res, 2021,
9945725. doi:10.1155/2021/9945725

Querques, G., Cicinelli, M. V., Rabiolo, A., de Vitis, L., Sacconi, R., Querques, L., & Bandello, F.
(2018). Laser photocoagulation as treatment of non-exudative age-related macular

158
degeneration: state-of-the-art and future perspectives. Graefes Arch Clin Exp
Ophthalmol, 256(1), 1-9. doi:10.1007/s00417-017-3848-x

Rabelo, L. A., Alenina, N., & Bader, M. (2011). ACE2–angiotensin-(1–7)–Mas axis and oxidative
stress in cardiovascular disease. Hypertension Research, 34(2), 154-160.
doi:10.1038/hr.2010.235

Ramadan, R., Dhawan, S. S., Binongo, J. N., Alkhoder, A., Jones, D. P., Oshinski, J. N., & Quyyumi,
A. A. (2016). Effect of Angiotensin II Type I Receptor Blockade with Valsartan on Carotid
Artery Atherosclerosis: A Double Blind Randomized Clinical Trial Comparing Valsartan
and Placebo (EFFERVESCENT). Am Heart J, 174, 68-79. doi:10.1016/j.ahj.2015.12.021

Raoul, W., Feumi, C., Keller, N., Lavalette, S., Houssier, M., Behar-Cohen, F., . . . Sennlaub, F.
(2008). Lipid-bloated subretinal microglial cells are at the origin of drusen appearance in
CX3CR1-deficient mice. Ophthalmic Res, 40(3-4), 115-119. doi:10.1159/000119860

Reddy, R., Asante, I., Liu, S., Parikh, P., Liebler, J., Borok, Z., . . . Louie, S. G. (2019). Circulating
angiotensin peptides levels in Acute Respiratory Distress Syndrome correlate with
clinical outcomes: A pilot study. PLoS One, 14(3), e0213096.
doi:10.1371/journal.pone.0213096

Rein, D. B., Wittenborn, J. S., Zhang, X., Honeycutt, A. A., Lesesne, S. B., Saaddine, J., & Vision
Health Cost-Effectiveness Study, G. (2009). Forecasting age-related macular
degeneration through the year 2050: the potential impact of new treatments. Arch
Ophthalmol, 127(4), 533-540. doi:10.1001/archophthalmol.2009.58

Ren, C., Liu, W., Yin, X., Zhang, B., & Lu, P. (2020). Renin-Angiotensin System Inhibitor Usage and
Age-Related Macular Degeneration among Hypertensive Patients: Results from the
National Health and Nutrition Examination Survey, 2005-2008. J Ophthalmol, 2020,
4252031. doi:10.1155/2020/4252031

Richert, E., Koinzer, S., Tode, J., Schlott, K., Brinkmann, R., Hillenkamp, J., . . . Roider, J. (2018).
Release of Different Cell Mediators During Retinal Pigment Epithelium Regeneration
Following Selective Retina Therapy. Invest Ophthalmol Vis Sci, 59(3), 1323-1331.
doi:10.1167/iovs.17-23163

Rodanant, N., Friberg, T. R., Cheng, L., Aurora, A., Bartsch, D., Toyoguchi, M., . . . Freeman, W. R.
(2002). Predictors of drusen reduction after subthreshold infrared (810 nm) diode laser
macular grid photocoagulation for nonexudative age-related macular degeneration. Am
J Ophthalmol, 134(4), 577-585. doi:10.1016/s0002-9394(02)01691-4

Rodgers, K., Abiko, M., Girgis, W., St Amand, K., Campeau, J., & diZerega, G. (1997).
Acceleration of dermal tissue repair by angiotensin II. Wound Repair Regen, 5(2), 175-
183. doi:10.1046/j.1524-475X.1997.50210.x

159
Rodgers, K. E., Espinoza, T., Felix, J., Roda, N., Maldonado, S., & diZerega, G. (2003).
Acceleration of healing, reduction of fibrotic scar, and normalization of tissue
architecture by an angiotensin analogue, NorLeu3-A(1-7). Plast Reconstr Surg, 111(3),
1195-1206. doi:10.1097/01.PRS.0000047403.23105.66

Rodgers, K. E., Roda, N., Felix, J. E., Espinoza, T., Maldonado, S., & diZerega, G. (2003).
Histological evaluation of the effects of angiotensin peptides on wound repair in
diabetic mice. Exp Dermatol, 12(6), 784-790. doi:10.1111/j.0906-6705.2003.00087.x

Romdhoniyyah, D. F., Harding, S. P., Cheyne, C. P., & Beare, N. A. V. (2021). Metformin, A
Potential Role in Age-Related Macular Degeneration: A Systematic Review and Meta-
Analysis. Ophthalmol Ther, 10(2), 245-260. doi:10.1007/s40123-021-00344-3

Ruan, Y., Jiang, S., & Gericke, A. (2021). Age-Related Macular Degeneration: Role of Oxidative
Stress and Blood Vessels. Int J Mol Sci, 22(3). doi:10.3390/ijms22031296

Rubner, R., Li, K. V., & Canto-Soler, M. V. (2022). Progress of clinical therapies for dry age-
related macular degeneration. Int J Ophthalmol, 15(1), 157-166.
doi:10.18240/ijo.2022.01.23

Salo, T., Makela, M., Kylmaniemi, M., Autio-Harmainen, H., & Larjava, H. (1994). Expression of
matrix metalloproteinase-2 and -9 during early human wound healing. Lab Invest, 70(2),
176-182. Retrieved from https://www.ncbi.nlm.nih.gov/pubmed/8139259

Santos, R. A., Simoes e Silva, A. C., Maric, C., Silva, D. M., Machado, R. P., de Buhr, I., . . .
Walther, T. (2003). Angiotensin-(1-7) is an endogenous ligand for the G protein-coupled
receptor Mas. Proc Natl Acad Sci U S A, 100(14), 8258-8263.
doi:10.1073/pnas.1432869100

Santos, R. A. S., Oudit, G. Y., Verano-Braga, T., Canta, G., Steckelings, U. M., & Bader, M. (2019).
The renin-angiotensin system: going beyond the classical paradigms. Am J Physiol Heart
Circ Physiol, 316(5), H958-H970. doi:10.1152/ajpheart.00723.2018

Sharma, R., Bose, D., Maminishkis, A., & Bharti, K. (2020). Retinal Pigment Epithelium
Replacement Therapy for Age-Related Macular Degeneration: Are We There Yet? Annu
Rev Pharmacol Toxicol, 60, 553-572. doi:10.1146/annurev-pharmtox-010919-023245

Shentu, X.-C., Ping, X.-Y., Cheng, Y.-L., Zhang, X., Tang, Y.-L., & Tang, X.-J. (2018). Hydrogen
peroxide-induced apoptosis of human lens epithelial cells is inhibited by parthenolide.
International journal of ophthalmology, 11(1), 12-17. doi:10.18240/ijo.2018.01.03

Silva, G. M., Franca-Falcao, M. S., Calzerra, N. T. M., Luz, M. S., Gadelha, D. D. A., Balarini, C. M.,
& Queiroz, T. M. (2020). Role of Renin-Angiotensin System Components in

160
Atherosclerosis: Focus on Ang-II, ACE2, and Ang-1-7. Front Physiol, 11, 1067.
doi:10.3389/fphys.2020.01067

Sivaprasad, S., & Chong, N. V. (2006). The complement system and age-related macular
degeneration. Eye (Lond), 20(8), 867-872. doi:10.1038/sj.eye.6702176

Song, Q. H., Klepeis, V. E., Nugent, M. A., & Trinkaus-Randall, V. (2002). TGF-beta1 regulates
TGF-beta1 and FGF-2 mRNA expression during fibroblast wound healing. Mol Pathol,
55(3), 164-176. doi:10.1136/mp.55.3.164

Suzuki, M., Kamei, M., Itabe, H., Yoneda, K., Bando, H., Kume, N., & Tano, Y. (2007). Oxidized
phospholipids in the macula increase with age and in eyes with age-related macular
degeneration. Mol Vis, 13, 772-778. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/17563727

Suzuki, Y., Ruiz-Ortega, M., & Egido, J. (2000). Angiotensin II: a double-edged sword in
inflammation. J Nephrol, 13 Suppl 3, S101-110. Retrieved from
https://www.ncbi.nlm.nih.gov/pubmed/11132026

Swinney, D. C. (2011). Chapter 18 - Molecular Mechanism of Action (MMoA) in Drug Discovery.
In J. E. Macor (Ed.), Annual Reports in Medicinal Chemistry (Vol. 46, pp. 301-317):
Academic Press.

Takeda, H., Katagata, Y., Hozumi, Y., & Kondo, S. (2004). Effects of angiotensin II receptor
signaling during skin wound healing. Am J Pathol, 165(5), 1653-1662.
doi:10.1016/S0002-9440(10)63422-0

Theodore Smith, R. (2019). Sub-threshold nanosecond laser (SNL) treatment in intermediate
AMD (IAMD). Ann Eye Sci, 4. doi:10.21037/aes.2018.12.04

Tuo, J., Bojanowski, C. M., Zhou, M., Shen, D., Ross, R. J., Rosenberg, K. I., . . . Chan, C. C. (2007).
Murine ccl2/cx3cr1 deficiency results in retinal lesions mimicking human age-related
macular degeneration. Invest Ophthalmol Vis Sci, 48(8), 3827-3836. doi:10.1167/iovs.07-
0051

van der Vliet, A., & Janssen-Heininger, Y. M. (2014). Hydrogen peroxide as a damage signal in
tissue injury and inflammation: murderer, mediator, or messenger? J Cell Biochem,
115(3), 427-435. doi:10.1002/jcb.24683

Van Haeringen, N. J. (1996). The renin-angiotensin system in the human eye. Br J Ophthalmol,
80(2), 99-100. doi:10.1136/bjo.80.2.99

161
van Lookeren Campagne, M., LeCouter, J., Yaspan, B. L., & Ye, W. (2014). Mechanisms of age-
related macular degeneration and therapeutic opportunities. J Pathol, 232(2), 151-164.
doi:10.1002/path.4266

Vavvas, D. G., Daniels, A. B., Kapsala, Z. G., Goldfarb, J. W., Ganotakis, E., Loewenstein, J. I., . . .
Miller, J. W. (2016). Regression of Some High-risk Features of Age-related Macular
Degeneration (AMD) in Patients Receiving Intensive Statin Treatment. EBioMedicine, 5,
198-203. doi:10.1016/j.ebiom.2016.01.033

Velez-Montoya, R., Oliver, S. C., Olson, J. L., Fine, S. L., Quiroz-Mercado, H., & Mandava, N.
(2014). Current knowledge and trends in age-related macular degeneration: genetics,
epidemiology, and prevention. Retina, 34(3), 423-441.
doi:10.1097/IAE.0000000000000036

Virgili, G., Michelessi, M., Parodi, M. B., Bacherini, D., & Evans, J. R. (2015). Laser treatment of
drusen to prevent progression to advanced age-related macular degeneration. Cochrane
Database Syst Rev(10), CD006537. doi:10.1002/14651858.CD006537.pub3

Wang, T., Lian, G., Cai, X., Lin, Z., & Xie, L. (2018). Effect of prehypertensive losartan therapy on
AT1R and ATRAP methylation of adipose tissue in the later life of high-fat-fed
spontaneously hypertensive rats. Mol Med Rep, 17(1), 1753-1761.
doi:10.3892/mmr.2017.8081

Wang, Y., Zhong, Y., Zhang, L., Wu, Q., Tham, Y., Rim, T. H., . . . Liu, L. (2021). Global Incidence,
Progression, and Risk Factors of Age-Related Macular Degeneration and Projection of
Disease Statistics in 30 Years: A Modeling Study. Gerontology, 1-15.
doi:10.1159/000518822

Wilkinson-Berka, J. L., Suphapimol, V., Jerome, J. R., Deliyanti, D., & Allingham, M. J. (2019).
Angiotensin II and aldosterone in retinal vasculopathy and inflammation. Exp Eye Res,
187, 107766. doi:10.1016/j.exer.2019.107766

Wilson, A. S., Hobbs, B. G., Shen, W. Y., Speed, T. P., Schmidt, U., Begley, C. G., & Rakoczy, P. E.
(2003). Argon laser photocoagulation-induced modification of gene expression in the
retina. Invest Ophthalmol Vis Sci, 44(4), 1426-1434. doi:10.1167/iovs.02-0622

Wolter, J. R., & Falls, H. F. (1962). Bilateral confluent drusen. Arch Ophthalmol, 68, 219-226.
doi:10.1001/archopht.1962.00960030223013

Wright, C. B., & Ambati, J. (2017). Dry Age-Related Macular Degeneration Pharmacology. Handb
Exp Pharmacol, 242, 321-336. doi:10.1007/164_2016_36

162
Xiang, J., Wan, C., Guo, R., & Guo, D. (2016). Is Hydrogen Peroxide a Suitable Apoptosis Inducer
for All Cell Types? BioMed Research International, 2016, 7343965.
doi:10.1155/2016/7343965

Xie, J. X., Hu, J., Cheng, J., Liu, C., & Wei, X. (2022). The function of the ACE2/Ang(1-7)/Mas
receptor axis of the renin-angiotensin system in myocardial ischemia reperfusion injury.
Eur Rev Med Pharmacol Sci, 26(6), 1852-1859. doi:10.26355/eurrev_202203_28330

Zarbin, M. A. (2004). Current concepts in the pathogenesis of age-related macular
degeneration. Arch Ophthalmol, 122(4), 598-614. doi:10.1001/archopht.122.4.598

Abstract (if available)

Abstract Age-related macular degeneration (AMD) is a leading cause of vision loss in the United States. While VEGF inhibitors are available for treating wet AMD (wAMD), there are currently no FDA-approved treatments for dry AMD (dAMD). The hallmark of AMD is the presence of drusen, lipid and protein amalgamations that accumulate in the subretinal space, for which their increase in size and volume is positively correlated with disease progression. This correlation has made drusen deposits, which are compositionally similar to atherosclerotic plaques, a primary target in developing treatments for dAMD, although the cause of accumulation is still unclear. Selective Retinal Therapy (SRT), is a treatment proposed for dAMD, for which subthreshold nano-lasers are used to selectively destroy RPE cells and initiate mechanisms of wound repair. SRT lasers have previously been shown to reduce drusen deposits in clinic, although the underlying mechanism beyond extracellular matrix turnover is unclear. The Renin-Angiotensin System (RAS) has previously been shown to play a role in wound healing from thermal injury and atherosclerosis, but a gap in the knowledge for whether stimulation of ocular RAS can be beneficial for drusen reduction in dAMD. We utilized R:GEN, a proprietary SRT laser, to demonstrate its safety and efficacy in vivo using two genetic knockout mouse models of AMD, and explore its ability to stimulate ocular RAS as part of its drusen reduction mechanism. This body of work provides evidence that RAS modulation may be beneficial for drusen reduction and dAMD and establishes the foundation for probing combination therapy.

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Asset Metadata

Creator Zhou, Eugene (author)

Core Title Laser induced modulation of ocular renin-angiotensin system for treatment of dry age-related macular degeneration

School School of Pharmacy

Degree Doctor of Philosophy

Degree Program Clinical and Experimental Therapeutics

Degree Conferral Date 2023-05

Publication Date 01/19/2024

Defense Date 06/23/2022

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag AMD,angiotensin,laser,OAI-PMH Harvest,selective retinal therapy

Format theses (aat)

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Louie, Stan (committee chair), Asante, Isaac (committee member), Asatryan, Liana (committee member)

Creator Email eugeneczhou@gmail.com,eugenezh@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-oUC25344

Unique identifier UC25344

Identifier etd-ZhouEugene-11425.pdf (filename)

Legacy Identifier etd-ZhouEugene-11425

Document Type Dissertation

Format theses (aat)

Rights Zhou, Eugene

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Type texts

Source 20230120-usctheses-batch-1002 (batch), University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

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