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Mediating the homeostatic response during light & cytotoxic induced retinal damage

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Mediating the homeostatic response during light & cytotoxic induced retinal damage

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MEDIATING THE HOMEOSTATIC RESPONSE DURING LIGHT & CYTOTOXIC INDUCED
RETINAL DAMAGE

By

Joshua P. Kramer

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
Biology of Aging

August 2021

Copyright 2021 Joshua P. Kramer
ii
Acknowledgements

There are no words to truly express how grateful I am to have been given the opportunity
to mentor under so many talented individuals at USC, the Buck Institute, and UCSF. The wide
variety of people of all ages, backgrounds and training provided me the support I needed to get
where I am today through their technical expertise, resource assistance, and contagious
enthusiasm for science. The time I spent in each place drastically changed my perspective on
how great research is conducted. As I enter the final stretch of my PhD experience, I have found
a new appreciation for all the help, mentorship, and advice I have been given. I would especially
like to thank my mentors, Dr. Deepak Lamba and Dr. Henri Jasper, who have stood by and
supported me through both my mistakes and successes and have helped steer my work in the
right direction when things did not go as planned. I would not be where I am today without their
careful and patient guidance and advice.

I want to thank my parents, Fred and Anne Kramer, and my brother, Ben, for always
encouraging me to continue to nourish my zeal for research through the frustrations and
challenges that come with doctoral work, no matter how difficult they may seem. While I was
unable to stay in contact as closely as I wished, your words and kindness enabled me to continue
following my dreams toward the future.

I would like to show appreciation to the fellow students of the Biology of Aging Program.
Sharing our struggles and building comradery was one of the defining experiences of my time at
both USC and the Buck Institute. I know you will all go on to do great things, and I am thankful to
both have been a part of your journey and you a part of mine.
iii

I want to give a special thanks to the vice dean of the Biology of Aging program, Dr. Kelvin
Davies, for his help and advice he has given me throughout the program. I will never forget how
kind and considerate he was in helping my girlfriend and I find a safe place to stay at USC after
we had been scammed out of an apartment that we booked sight unseen when on the other side
of the country. His kindness and warm demeanor helped welcome us into the state we have come
to call home.

I also want to thank all the individual members and administrators of the Lamba and Jasper
Labs who have continued to help advise and mentor me throughout this process. You have all
always been willing to lend a hand, discuss scientific minutia, or offer constructive criticism on a
presentation. Thank you to Shereen Chew for always being there to help with protocol details or
lend a hand on experiments. Thank you to Katie Chirco for insightful suggestions and strategy
that I continue to use to this day. Thank you to Sangeetha for all of your suggestions and advice
to help refine my ideas and experimental design. Your help was invaluable and always
appreciated.

Finally, to my partner, Kelly Brown, you have been my rock and my support throughout
this entire process, always there to encourage me every late night, early morning, practice session
and research struggle. Your positive and cheerful outlook has kept me working at my best
throughout the hard times along the way. I am forever grateful for all you have done to help me
reach this point.
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Table of Contents

Acknowledgements ii
List of Figures vi
List of Tables vii
Abbreviations viii
Abstract x
Chapter I: Introduction 1
Strengths of Studying Visual System 1
Overview of Mouse Eye/Retina 3
Overview of Fly Eye/Retina 5
Retinal Immune Cell Diversity 9
Support Cell Functionality 11
Sources of Retinal Inflammation 13
Light Exposure 13
UV Irradiation 14
NMDA Cytotoxicity 16
Retinal Mechanisms in the Healing Process 16
Inflammatory Retinal Homeostasis 16
Dpp/TGFβ Superfamily Pathway/Interactions 22
Dpp 22
BMP 23
TGFβ 24
Dpp/TGFB Superfamily Cell-Injury Interactions 25
Cell Replacement 26
Support Cell Activation 26
Mobilization of Immune Cells 27
Antioxidant Response 28
Inhibition of Apoptosis 28
Bridging Gaps between Model Organisms 28
Drosophila Melanogaster as a Model Organism for Retinal Injury 29
M. musculus as a Model Organism to Retinal Injury 30
Limitations of Flies/Mice as Model Organisms 30
Translational Research 31
Chapter II: Dpp/TGFβ-superfamily play a dual role in Mediating Retinal Damage Response 32
Results 32
Hemocyte derived Dpp controls damaged retinal tissue response 32
Immune cell mouse BMP/TGFB is expressed in a timed manner 38
v

Modulation of BMP signaling protects against induced retinal apoptosis 40
Inhibition of TGFB signaling protects against induced retinal apoptosis 41
Microglia activation can be modulated by manipulating BMP/TGFB pathways 43
Discussion 43
Materials and Methods 47
Mouse Genetics & Damage Models 47
Drosophila Genetics & Damage Models 48
Chapter III: Conclusion and Final Thoughts 49
Hemocyte derived Dpp controls the Drosophila damaged retinal tissue response 49
Under UV radiation damage, Sax/Smox inhibit apoptosis and Mad/Tkv promote it 49
BMP4 activation/TGFβ inhibition protect against damage ind. retinal apoptosis 50
Supplemental Information 52
References 59

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List of Figures
Figure 1.1: Brief overview of Mouse Eye. .................................................................................... 4
Figure 1.2: Brief Overview of Mouse Retina. ............................................................................... 5
Figure 1.3: Simplified Drosophila Unit Eye (Ommatidium)........................................................... 8
Figure 1.4: Radiation in the Retina…………….. ........................................................................ 15
Figure 1.5: Inflammatory Retinal Homeostasis. ......................................................................... 21
Figure 1.6: Canonical Dpp Pathway in Drosophila. ................................................................... 23
Figure 1.7: Simplified Canonical BMP/TGFβ Pathway in Mammals. ......................................... 25
Figure 2.1: Dpp inhibition controls JNK and UV driven apoptosis in Drosophila. ...................... 33
Figure 2.2: Following UV driven whole eye radiation damage, Sax and Smox RNAi inhibit
apoptosis, while Mad and Tkv RNAi promote it.. ....................................................................... 36
Figure 2.3: BMP and TGFβ activation occurs in a time dependent manner following light
stress and damage.
.................................................................................................................................................... 40
Figure 2.4: Modulation of the BMP/TGFβ signaling pathway affects tissue damage induced
retinal apoptosis and immune activation.................................................................................... 42
Figure 3.1: Diagram of Dpp and TGFβ/BMP signaling act as central controllers of
photoreceptor death and survival after acute damage within the retina.
.................................................................................................................................................... 51
Figure S3.1: Representative images and quantitation of control pMad+ Elav+ cells. …..….….. 52
Figure S3.2: Quantitation of Individual Drosophila eyes. ............................................................54
Figure S3.3: Quantitation of Individual Drosophila eyes continued. .......................................... 56
Figure S3.4: Comparing UV exposed Sep::G4;Smox and MadRNAi individual progeny
DCP1+ Elav+ cells to UV Sep WT.................................................................................. 57
Figure S3.5: Representative images of pSmad 1/5/8 (BMP) and pSmad 2/3
(TGFB) animals .............................................................................................................. 58
vii
List of Tables
Table 1.1: Benefits of Investigating Visual System ................................................................... 2
Table 1.2: Immune Cells within the Choroid and Retina under
Homeostatic Conditions................. ………………………………...………………………...11
Table 1.3: Activated Microglial Types ........................................................................................17
Table 1.4: M2c Microglial Procedural Roles in Retinal Repair.................................................. 19
Table 1.5: Chick BMP4, Receptor & Inhibitor Expression Timeline.......................................... 27

viii
Abbreviations
JNKK - Jun N-terminal Kinase Kinase
Hep - Hemipterous
BMP – Bone Morphogenic Protein
TGFβ - Transforming Growth Factor Beta
NMDA - N-methyl D-aspartic Acid
Dpp - Decapentaplegic Protein
Tkv - Thickveins
Mad - Mothers Against Dpp
Smad - Small Mothers against Dpp
Smox - Smad on X
Wg - Wingless
Hh - Hedgehog
JAK/STAT - Janus kinases, Signal Transducer and Activator of Transcription proteins
CNS - Central Nervous System
ONL - Outer Nuclear Layer
GCL - Ganglion Cell Layer
CRALBP - Cellular Retinaldehyde–Binding Protein
GABA - Gamma Aminobutyric Acid
ASCL1 - Achaete-Scute Complex Homolog 1
GFAP - Glial Fibrillary Acidic Protein
GDFs - Growth Differentiation Factors
GDNFs - Glial Derived Neurotrophic Factors
Pros - Prospero
Prox1 - Prospero homeobox protein 1
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Pax2 - Paired box protein
Oli - Oligodendrocyte lineage protein
Olig1,2 - Oligodendrocyte lineage protein 1,2
ABCA - ATP-binding cassette, subclass A
RPE - Retinal Pigmented Epithelium
Kir4.1 - Inwardly Rectifying Potassium channel 4.1
UV - Ultraviolet
RGC - Retinal Ganglion Cell
cAMP - cyclic Adenosine Monophosphate
CREB - cAMP Response Element Binding
PRR - Pattern Recognition Receptor
DAMP - Damage Associated Molecular Pattern Molecules
ATP - Adenosine Triphosphate
NO - Nitric Oxide (NO)
ROI - Reactive Oxygen Intermediates
CD163 - Cluster of Differentiation 163
CD206 - Cluster of Differentiation 206
IL - Interleukin
LPS - Lipopolysaccharide
Pax6 - Paired Box Protein 6
INL - Inner Nuclear Layer
GDF - Growth Differentiation Factor
GDNF - Glial Cell Line-Derived Neurotrophic Factor

x

Abstract
Visual perception through the eye is crucial for most animals’ survival in the wild. This
remains true among humans, who devote two-thirds of all electrical activity and fifty percent of all
neural tissue in the brain to vision (Sells & Fixott, 1957; Kolster, 2012; Sheth et al., 2016).

Within the eye, the retina is the main light sensing tissue. It is responsible for translating
an organisms’ perception of the world into neural impulses in the brain. The retina consists of
several basic structures; photoreceptor cells, support cells, pigmented epithelium, and resident
immune cells, all of which work together to sustain the organ (Barbot et al., 2017; Baden et al.,
2019). Any injury that damages the individual structures of the retina must be handled rapidly to
maintain its function.

Retinal homeostasis depends on elaborate management of cell survival and apoptosis in
response to injury. The result of this outcome is crucial for the organism’s long-term survival as
most retinal cells are irreplaceable. However, the signaling mechanisms that control these
processes remain poorly understood. Decapentaplegic (Dpp), the first identified secreted
morphogen, is required for tissue formation, growth, scope, and maturation in the Drosophila eye.
Due to previous findings (Kelsey et al., 2012; Ayyaz et al., 2015; Neves et al., 2016), we
hypothesized that retinal morphogenic factors involved in growth and maturation such as Dpp
may also play a role in tissue repair and recovery.

We first chose to resolve the question of whether Drosophila immune cells, ninety five
percent of which are hemocytes, drove expression of Dpp under retinal injury. To accomplish this,

xi
we used a previously characterized fly line overexpressing a constitutively active form of the Jun
N-terminal Kinase Kinase (JNKK) Hemipterous (Hep) under the control of the photoreceptor and
cone cell driver Sep-Gal4 (Therrien et al., 1999; Jasper et al., 2002; Kanda et al., 2004; Luo et
al., 2007; Kelsey et al., 2012). When Sep-Hep
ACT
was crossed to flies expressing Sax-RNAi or
Smox-RNAi under the control of UAS, a significant increase in surviving photoreceptors compared
to control flies was observed. Over-expression of the negative feedback inhibitor of Dpp signaling,
Daughters against Dpp (Dad; Tsuneizumi et al., 1997; Kamiya et al., 2008), also promoted
survival. Conversely, we found that knocking down Tkv and Mad resulted in a significant decrease
in surviving photoreceptors. When Sep-Hep
ACT
was crossed to flies expressing Hemolectin-
Dpp
RNAi
apoptosis was also significantly reduced.

To answer the question of how eye injury drives Dpp expression, we used a Drosophila
model for irradiation-induced damage, collecting glass multimer reporter driver (GMR-Gal4)
(Moses and Ruben, 1991) to express RNAi targeting Dpp pathway components in all postmitotic
cells of the retina. We find that the BMP orthologue Decapentaplegic (Dpp) is released by immune
cells after irradiation and stimulates apoptosis shortly after damage through the type I receptor
Thickveins (Tkv) and the signal transducer Mothers against Dpp (Mad). At a later stage, the
alternative type I receptor Saxophone, and its target Smad on X (Smox) promote tissue repair
and photoreceptor survival.

In mammals, Dpp homologues include various members of the bone morphogenic protein
(BMP) and transforming growth factor beta (TGFβ) family. Their downstream targets include
Smad 1/5/8 (activated by BMP) and Smad 2/3 (activated by TGFβ). BMP 2/4 have been shown
to act in an anti-inflammatory manner, biasing macrophages to their anti-inflammatory M2 subtype
(Talati et al., 2014; Dzik, et al., 2014; Ji et al., 2014; Shinohara et. al., 2010). These proteins also

xii
stimulate Müller glia proliferation, promote survival of retinal ganglion cells after damage, and
decrease microglial activation in vivo (Ueki et. al., 2012; Ueki et. al., 2013; Mitchell et. al., 2013;
Thompson et al., 2019). However, the timing and mechanism(s) by which BMP/TGFβ signaling
modulates repair in the mammalian retina is not fully understood. It is also unclear how these two
homologous pathways interact in this tissue.

To determine if the dual role of BMP signaling is conserved in the mammalian retina, we
induced separately either light damage or excitotoxicity in C57BL/6J mice. We found BMP
downstream signaling is activated in retinal support cells, the Müller glia. We then coupled this
treatment with supplementation of BMP4 or inhibition of BMP using small molecules. We found
that under both injury models, BMP4 promotes tissue survival under damage, while inhibition of
BMP negatively affects tissue survival after light damage.

Lastly, we sought to determine how innate immune cells of the eye were affected by the
BMP/TGFβ pathways. We accomplished this by analyzing changes in morphology which have
been shown to reflect activation (Karperien et al., 2013). We found that, in comparison to control,
with nearly all immune cells displaying highly ramified morphology, inhibition of BMP led to
significant reduction in ramified morphology, while supplementation led to a notable increase.

In conclusion, our data identify novel evolutionarily conserved mechanisms by which
retinal homeostasis is maintained. My work is first known to show that Dpp and TGFβ/BMP
signaling act as central controllers of photoreceptor death and survival after acute damage within
the retina.

1
Chapter I: Introduction
The following chapter contains portions adapted from an in-review article (J. Kramer, Neves,
Koniikusic, Jasper and Lamba, 2021).
Strengths of Studying Visual System
As most organisms are heavily reliant on vision for survival, visual ability correlates strongly
with health. Although the eyes only occupy 0.1 percent of the total body, injury that damages this
organ can have wide ranging effects that impact all tissues of the visual system (Blaivas, 2003;
Moraczewski, 2007). Additionally, neurodegenerative diseases that impact the brain and spinal cord
can first reveal themselves in the eye (London et al., 2013). Disease and damage that decrease
sight encompass more than just the eye. Vision loss can impact mobility, independence, overall
quality of life, and is correlated with increased risk of injury, reduced mental health, cognition,
employment, and social/educational accomplishment in both children and adults (Ray et al., 2008;
Chen et al., 2017, Safi et al., 2017; Wang et al., 2017). In addition, typical late term interventions
can be extremely cost prohibitive, out of reach to those of modest income (Busbee et al. 2003).
Therefore, addressing issues of the eye through mechanistic studies can have a significant impact
on both the health and wellness outcomes of an individual over a lifetime.

Providing care and therapy to the eye is not only useful to prevent these outcomes. The eye
can also be used to provide therapeutic interventions with convenient access to the brain and
nervous system. Photoreceptors are irreplaceable cells sharing many similarities with other
invaluable neuronal subtypes in the brain, and potential treatments of neuronal diseases can be
quantified noninvasively through many readily available tests. Additionally, symptoms in central
nervous system diseases often occur first in the eye (London et al., 2013). Thus, the eye can be an

2
ideal candidate for clinical trials and preliminary drug studies to address disorders of both the eye
and central nervous system (Hudry et al., 2019). Understanding the correlation of repair
mechanisms within the eye and retina may also elucidate parallels to neural repair in the brain (Wong
et al., 2020).

Both the eye and brain have a common cellular lineage of the embryonic neural tube during
development (Stenkamp, 2015). This shared formative connection also exists in the neurons and
glia of mice and flies, represented by two of the most common eye animal models used in
mechanistic studies, Mus Musculus and Drosophila Melanogaster (Sprecher et al., 2008; Jeibmann
et al., 2009; Hickman et al, 2017). As we increase our understanding of the relationship between
different mechanisms in the visual system of these and other model organisms and their shared
gene networks, more frequent discoveries of evolutionary conserved pathways of retinal
homeostasis and repair will occur. The ultimate goal of this work is to help pave the way toward
translational retinal interventions that both treat and mitigate future eye injury.

Table 1.1: Benefits of Investigating Visual System
Type of Utility Description
Eye as window to CNS
Useful to early diagnose specific CNS illnesses and
calibrate potential treatments
Irreplaceable photoreceptors are
similar to other brain neurons
Ease of study; Noninvasive quantitative tests can
be done on photoreceptor treatment to diagnose
impact
Correlated to long term quality of life
improvement
Ocular interventions have significant impact on long
term earning potential and happiness/satisfaction
Privileged immune state
Blood-retinal barrier keeps out larger molecules and
provides a better environment for drug application

3
Overview of Mouse Eye/Retina
The mature mouse visual system is very similar to that of other mammals. In brief, it
develops from a single layered retinal neuroepithelium region of the embryonic neural tube
expressing several eye field transcription factors. This region evaginates bilaterally from the
neural folds of the developing forebrain, forming the optic grooves (Kondoh, 2002). The area
becomes the optic vesicles with the eventual closure of the neural tube, initiated by the master
control homeobox gene small eye - known homologue in humans (Pax6) and Drosophila
(eyeless), forming the basis of bilateral eyes (Ashery-Padan et al., 2001; Heavner et al., 2012).

The optic vesicles, extending from the forebrain toward the surface ectoderm, form into
the optic cup, with bone morphogenic protein (BMPs) agonists and antagonists guiding proper
optic cup development (Wordinger et al., 2007; Teraoka et al., 2009). Blood vessels gain access
to the optic cup through a dip in the inferior surface known as the choroidal fissure, which
eventually fuses enclosing the vessels within the eye. The basal layer of the optic cup forms the
retinal pigment epithelium (RPE) and the apical layer forms the neural retina, which will eventually
consist of photoreceptors, ganglion cells, bipolar cells, horizontal cells, amacrine cells and Müller
glia. The mid region of the optic cup matures into the iris and ciliary body. The ectoderm thickens
and the lens vesicle separates from it to form the lens at the open end of the optic cup. The surface
ectoderm then produces the corneal epithelia, conjunctiva, the eyelids, and lacrimal apparatus
after sensing the presence of the lens. The remaining ocular tissue matures from the
mesenchyme.

4

Figure 1.1: Brief Overview of Mouse Eye. Diagram of structures of the mouse eye and where
they are located. From left to right; Cornea, Vitreous, Lens, Sclera, Muscle, Retina, Choroid.
(NEI-medialibrary-2817499.png. November 7
th
, 2019. Public Domain).

Vision in adult mice operates by light entering the eye through the cornea, where it refracts
and passes through the pupil and is further refracted by the lens. The resulting image is projected
inversely on the retina. The retina detects this through its photoreceptor cells. Photoreceptors are
divided into cones and rods, which have differences in both function and location in the retina. In
humans, cones are found mostly in the center of the retina, and are divided into short - blue,
middle - green, or long - red by the wavelength range of light they absorb (Ortín-Martínez et al.,
2014). Rods reside throughout the retina except the very central foveal region and specialize in

5
absorbing low levels of light. These photoreceptors translate this detection into electrical signal
that is transmitted to the bipolar cells, which in turn transfer this information to ganglion cells which
project to the brain (De Mayo, 2014).

Figure 1.2: Brief Overview of Mouse Retina. Magnified view of mouse retina, showing neuronal
structure, layering and network. From top to bottom; Retinal Pigment Epithelium (brown), Rod
Photoreceptor (green), Cone Photoreceptor (blue), Müller Glia (pink), Horizontal cell (light blue),
Bipolar cell (grey), Ganglion cell (red).

Overview of Fly Eye/Retina
The adult Drosophila visual network has significant homology in both cell type and function
with the mammalian ocular system. However, early development of the fly visual system is distinctly
different.

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The foundations of the Drosophila eye form early in embryonic development as an
ectodermal region that curls into the interior of the embryo. This walled-off region connects directly
to the forming larval photoreceptor neurons known as bolwig’s organ, and the dual tissues are
moved through the mechanism of head involution, like a sock being turned inside out, into their final
position. Firmly attached to the developing brain lobe, the organ now known as the eye imaginal
disk begins formation of the optic stalk. The stalk will be used by all future photoreceptors to connect
their axons to the brain. Conversely, glial cells from the brain grow outward to the eye disk along
Bolwig’s nerve. The eye imaginal disk then grows asymmetrical, divided by a peripodial membrane,
and one side becomes columnar epithelium, where neurogenesis will take place (VanHook and
Letsou, 2007).

Beginning with the second instar stage of larval development, the number of cells in the eye
field increases from 130 to 1,500 by the beginning of the third instar. During the third instar, the eye
field, which consists of 10,000 individual cells, ceases mitosis and a wave of maturation is induced
by Decapentaplegic (Dpp) and tuned by Wingless (Wg), which act in opposition. This activity, known
as the morphogenetic furrow, crosses the eye imaginal disk, and causes a G1 cell cycle arrest,
shifting cells to a pre-proneural state where they can respond to Hedgehog (Hh) and Atonal signaling
to convert pre-photoreceptor R-cells into mature photoreceptors. Once the photoreceptor neurons
are created they form axons that grow toward the brain lobes. R1-6 axons end in the lamina, while
R7-8 axons terminate in the medulla (Silies et al., 2010). This leads to formation of the approximately
760 unit ommatidium of the Drosophila eye. The eye becomes fully postmitotic soon after the third
instar as the fly transitions to the pupa state.

7
Each individual ommatidium unit is made up of a hexagonal structure of eight photoreceptor
neurons - R1 through 8, four non neuronal lens secreting cone cells, two primary pigment cells, three
tertiary pigment cells and three bristle cells. The basal region of the ommatidium is covered by a
transparent cornea. The pigment cells act to isolate the visual field of one ommatidium from another,
improving Drosophila vision, as each ommatidium unit eye is responsible for sending the brain one
picture element, which gets processed first at the lamina before getting sent to the brain (Land et
al., 2012). The eye field generates more cells then are required, all of which resolve into either
excess secondary or tertiary pigment cells. During the mid-pupal stage a final wave of programmed
cell death occurs to eliminate the surplus cells (Kumar, 2013).

In the adult Drosophila eye, light transmits through the cornea and cone where it is refracted
and is then detected by the eight individual photoreceptor cells depending upon its wavelength, and
the resulting light translated in adjacent compartment known as the rhabdomere, resulting in a seven
pixel inverted image being formed in each unit eye. The unit eyes are aligned such that the field of
view within an ommatidium is the same as that between them before being sent to the brain for full
resolution (Land, 2012).
.

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Figure 1.3: Simplified Drosophila Unit Eye (Ommatidium). Cross-sectional diagram of
individual unit eye covering basic structures. Bristle shaft (black), cornea (light blue), cone glial
cell (yellow), rhabdomere (orange), primary pigmented cell (inner dark blue region), secondary
pigment cell (outer dark blue region), Adapted from (Land, 2012). photoreceptors R1-7 (green),
photoreceptor R8 (middle bottom of orange line), cone cell foot (bottom yellow region). Adapted
from (Song et al., 2009).

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Retinal Immune Cell Diversity
Within the retina resides a variety of immune cell types, each performing a different role in
the maintenance of the eye. However, under normal conditions, the immune diversity within the
retinal tissue is less than other regions of the body.

In Drosophila, more than ninety percent of all blood cells, known as hemocytes, are
plasmatocytes (Gold & Brückner, 2016). Plasmatocytes, similar to macrophages in mammals, are
phagocytic immune cells focused on placement of extracellular matrix, clearance of cellular waste,
invading foreign bodies, and aiding in the repair of damaged tissues. A tiny proportion of these
hemocytes are crystal cells, which induce melanization during injury and infection (Lebestky, 2000).
They maintain their populations through self-renewing hematopoietic pockets in muscles adjacent
to the eye. Under inflammatory conditions, hemocytes activate lamellocyte formation via release of
unpaired 2 and 3 and activation of the JAK/STAT pathway. Lamellocytes primarily are responsible
for encapsulation of foreign bodies (Stofanko et al., 2010).

In the mouse, the majority of innate immune cells in the retina are microglia. Microglia are
yolk sac originated cells of macrophage lineage. Under normal conditions they exist in a crystalline
pattern adjacent to bipolar cells in the middle of the retina - spaced between the nerve fiber/ganglion
cell layer and inner/outer plexiform layers. Under homeostatic conditions microglia in this region are
ramified, with long branching processes that extend and retract repeatedly to monitor their
immediate environment. In both the CNS and retina, they are typically marked as CD11b+, Iba-1+,
CX3CR1+, Tmem119+, CD45 low (Lee et al., 2008; Eme-Scolan & Dando, 2020). Along with
phagocytic activity, microglia manage and prune neural connections while maintaining
communication with support cells such as the Müller glia, astrocytes, and each other (Rathnasamy
et al., 2018).

10

Due to the exclusion of large molecular weight molecules, proteins, erythrocytes and
leukocytes by the blood retinal barrier, the retina operates under similar conditions to that of the
brain, showing a reduced inflammatory immune response to foreign substances. In accordance with
this, microglia are more likely to apoptosis under overactivation than other specialized immune cells
utilized by the body outside the eye (Rathnasamy et al., 2018). Microglia also actively inhibit
macrophage penetration during wound resolution (Plemel et al., 2020).

Below the retina lies the region of eye called the choroid, a highly vascularized region of
diffuse connective tissue that provides nutrients and growth factors to the retina. Within the choroid
reside other immune cells of the eye, macrophages, dendritic cells and mast cells (Chinnery et al.,
2017). Macrophages of the choroid primarily act in a phagocytic role, patrolling the expanse of the
tissue and digesting foreign bodies, pathogens, dead and dying cells, and protein aggregates
(Plernel et al., 2020). Dendritic cells within the choroid obtain, refine, and present antigens to B and
T cells. Lastly, mast cells, originating from hematopoietic progenitors, mediate vascular
homeostasis, venom detoxification, and help regulate the function of T cells, B cells, fibroblasts,
macrophages, and dendritic cells through the release of an array of proteases, histamines,
cytokines, chemokines, and growth factors (Bousquet et al., 2015).

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Table 1.2: Immune Cells within the Choroid and Retina under Homeostatic Conditions
Choroid Retina
Macrophages Microglia
Dendritic Cells -
Mast Cells -

Table 1.2: Immune Cells within the Choroid and Retina under Homeostatic Conditions. The
retina is a complex and varied tissue with multiple venues for immune response using the innate,
adaptive, and complement system to respond to retinal threats. However, under homeostatic
conditions, the vast majority of innate immune cells within the retina are Microglia.

Support Cell Functionality
Support cells play a crucial role in preserving the functions of the retina under normal
conditions. In mammals, this role in the retina is taken by astrocytes and Müller glia. Müller glia have
a particularly important role in communication with microglia to maintain homeostasis and operation
of the retina. In Drosophila, the primary support cell role is taken by ommatidial cone cells.
Although Müller glia nuclei are based in the inner nuclear layer, their processes branch out
across the width of the retina and extend to both the reaches of the outer nuclear layer (ONL) and
the outer edge of the ganglion cell layer (GCL), covering and acting as a pillar and brace for virtually
the entire retina. Their processes provide guidance and support for all the neurons of the retina -
photoreceptors, horizontal cells, amacrine cells, ganglion cells, horizontal cells and bipolar cells,
even helping cone and rod photoreceptors better detect light with their funnel like geometry and

12
guide their axons to maintain crosstalk and strengthen frequently used synaptic connections
(Franze, 2007).
Müller glia also communicate with the retinal innate immune cells, microglia, and play an
essential role in controlling the microenvironment of the retina over a lifetime in the face of increasing
inflammation. To perform this function they absorb neurotransmitters, control calcium gradients,
stow glycogen for later use, and clear waste (Goldman, 2014). This waste removal can include
ammonia, carbon dioxide, and spent neurotransmitters such as glutamate, GABA, dopamine, and
taurine.
In mice, previous work has shown that inhibition of histone deacetylase coinduced with
ASCL1 overexpression can drive regeneration of neurons from Müller glia (Jorstad, 2017). Work in
human paradigms have also shown Müller glia can also be induced to act as a multipotent progenitor
and source for rod photoreceptors (Giannelli et al., 2011; Bhatia et al., 2011).
Like Müller glia, astrocytes also provide structural support for the varied cells of the retina.
During development, astrocytes first reach the developing retina from the optic nerve. Their
distinctive morphological arrangement is of a pancake cell body with spoke-like processes. They act
to provide glial and axonal sheaths to retinal neurons, reinforce and help direct the growth of blood
vessels adjacent to the blood retinal barrier, and share glucose with surrounding neurons. Astrocytes
stain with antibodies against GFAP (Helmbacher, 2020)
In Drosophila, the glia subtype known as ommatidial cone cells, also called semper cells,
robustly express Drosophila homologue glial effectors and receptors. Like Müller glia in vertebrate
models, cone cells provide structural support to the unit eye, maintain neuronal activity and
neurotransmission, while preventing light driven retinal degeneration. Cone cells do this using a

13
three-pronged support pathway; Prospero/Prox1, Pax2, and Oli/Olig1,2 (Charlton-Perkins et al.,
2017). Pigment cells play a secondary role in supporting the ommatidium. They reduce aberrant
light diffusion, support the visual cycle and help maintain ion homeostasis within the unit eye
(Chaturvedi, 2014).
Sources of Retinal Inflammation
Inflammation in the retina can be due to a mis regulation of genetic/metabolic components
or infection, trauma, exposure to toxins, or as a natural consequence of light exposure while the
organism observes its surroundings.

Light Exposure
The visual system functions to perceive, place, and interpret the world around the organism.
Due to this, the retina, responsible for the detection of visible light, is the most important specialized
structure in the visual system, enabling organisms to discern their environment. It is also very
susceptible to light induced radiation damage. However, in patients over their mid-20’s, most low
intensity UV radiation can’t reach the retina due to the yellowing of the lens, as misfolded crystallin
protein accumulates in the lens and 3-hydroxykynurenine, which normally absorbs at 365 nm, is
increasingly photo modified. However, within the visible spectra of (400-700 nm), high intensity blue
light, ranging from (400-450 nm), is still able to penetrate the retina, and it is that exposure that is
most dangerous.

When visible light hits the eye, it is converted into electrical signals by 11-cis-retinal in
photoreceptors changing into all-trans retinal. The trans-retinal is shuttled by ABCA transport to the
RPE, where RPE65 converts it back to its 11-cis form and returns it to the photoreceptor. However,
the all-trans retinal is susceptible to oxidation, which naturally forms during high intensity light

14
exposure. This produces superoxide radicals and hydroperoxides that go on to induce protein and
lipid oxidation, crosslinking, and eventually ABCA transport dysfunction. This cascades into the
RPE, where the lysosome aggregates these indigestible byproducts, leading to pro-inflammatory
signaling and RPE dysfunction. If the transport system malfunction is severe enough this will lead
to photoreceptor apoptosis.

This digression may lead to deposit formation on top of Bruch’s membrane, the most inner
region of the choroid, which reduces its permeability and drives crosslinking from indigestible
oxidation products (Bloom et al., 2011). This cascade leads to increased angiogenic signaling which
mobilizes macrophages and microglia to the region, leading towards a cyclical increase in
inflammation and damage.

In addition to the negative effects of intense visible light on the photoreceptors and RPE,
excessive observable light also impacts Müller glia. Under thirty minutes of 405 nm blue light
exposure at 8 mW/cm
2
, rat Müller glia cells have been shown to experience overexpression of the
glial water channel aquaporin-4, mis regulation of Kir4.1 protein, and an engorgement of Müller glia
cells corresponding to reduction in potassium flow (Iandiev et al., 2008)

UV Irradiation
UV irradiation as a component of light exposure, has significant impacts on the health of the
retina, and high intensity exposure can drive photoreceptor cell death. All three categories of UV
radiation, A (315-400 nm), B (280-315 nm), and C (100-280 nm), have considerable injurious
consequences on the outcome of the eye. However, UV-B/C mostly impact the anterior regions of
the eye. Upon even low intensity short duration exposure, the cornea will become hyperemic and

15
inflamed. Lens exposure can precipitate formation of singlet oxygen driven oxidative stress,
denaturing collagen, and driving cataract formation (Cejková et al., 2010). UVA and B exposure is
generally infrequent in the normal population. Conversely, exposure to natural sources of UVA such
as the sun is much more frequent and can play an impactful role on the continued homeostasis of
the retina, although it generally does not result in mechanical or thermal damage (Behar-Cohen et
al., 2013). Like the previous section on high intensity blue light, UVA photons are similar enough in
wavelength that their effects on the internal structures of the retina are comparable, if magnified.
Successful management of these acute injuries requires a robust immune response (Glickman,
2011).

Figure 1.4: Radiation in the Retina. The cornea absorbs the majority of UVC, with UVA & B
radiation mostly absorbed by the lens. Portions of UVA radiation penetrate to the back of the eye,
where they can cause permanent damage to the retina and supporting cells.

16

NMDA Cytotoxicity
Induction of carefully titrated N-methyl D-aspartic acid (NMDA) into the retina, acting as an
excitotoxin, is a well characterized experimental model that induces swift retinal ganglion cell
(RGC) decline and apoptosis. The mechanism inducing cell death in the RGC is through abundant
inducement of glutamate receptor NMDA, which incites excitotoxicity through driving large
amounts of calcium ions into RGCs. This influx generates proteases, endonucleases, and
phospholipases to injure cell cytoskeleton, membranes and DNA. It also induces apoptosis via
cAMP response element binding (CREB) inhibition, mimicking several retinal diseases (Evangelho
et al., 2017; Christensen et al., 2019).
Retinal Mechanisms in the Healing Process
Inflammatory Retinal Homeostasis
(portions of the following section were adapted from (Kramer, Chirco and Lamba, 2019))

Under an inflammatory or injury condition in the retina, resident immune cells, and in some
cases circulating immune cells, are mobilized to the site of injury by activation of pattern recognition
receptors (PRRs) from damage associated molecular pattern molecules (DAMPs) released by dying
cells into the extracellular space. Some DAMPs include heat shock proteins, adenosine triphosphate
(ATP), and uric acid. PRRs receptors include Toll-like nucleotide-binding oligomerization domain
(NOD)-like, retinoic acid-inducible gene (RIG)-I-like, advanced glycation end-product and G-protein-
coupled (Gong et al., 2020). This binding trigger generation of mitogen-activated protein kinases
(MAPKs), interferon-α, nuclear factor kappa B (NF-κB), and cJun N-terminal kinase (JNK) (Israel,
2010). These inflammation inducers recruit immune cells to the region.

17

Table 1.3: Activated Microglial Types
Activated Microglial Types Description
M1 Pro-apoptotic NO/ROI producing
M2a Immune response against parasites
M2b Immune regulating subtype
M2c Anti-inflammatory subtype

Table 1.3: Activated Microglial Types. There are many microglial subtype populations that
occur in vivo, and microglia can belong to multiple categories. However, they most often
specialize into these four subtypes under the injury/resolution response.

Immune cells transform from the ramified patrolling form to either the M1, pro-apoptotic nitric
oxide (NO) or reactive oxygen intermediates (ROI) producing, or M2, anti-apoptotic group -
expressing CD163 and CD206, depending upon exposure to relative proportions of existing factors
and predetermined biases. The M2a immune cell, activated by IL-4, IL-10 and IL-13 and expressing
L-4R, Dectin-1 and CD209 are rarely seen in the retina and usually involved in the immune response
against parasites. The M2b immune type, known as the immune regulating subtype, is activated by
IL-1 and LPS produces IL-1, IL-6 and TNF alpha. The M2c is produced in the presence of IL-10 and
is known as a deactivated or anti-inflammatory immune cell (Zhou et al., 2017). Acting by subtype,
these arriving immune cells secrete soluble proteins, cytokines, and chemokines to induce
apoptosis, resolve injury, and promote healing.

As stated above, the mononuclear immune cells including microglia and macrophages are
critical innate immune responders to any retinal damage. The retinal inflammation process is
moderated by microglia, which upon sensing the injury becomes activated M1 microglia (Karlstetter
et al., 2015). Intracellular miRNA-155 expression has been shown to be important to keep them in

18
an M1 state (Hu et al., 2014). These M1 microglia migrate to the source of injury, proliferate, and
phagocytose debris, working with nearby astrocytes to release inflammatory cytokines and
chemokines such as TNFα, IL-1β, IL-6, IL-8, glutamate, reactive oxygen species, and nitric oxide,
all of which can result in the death of nearby susceptible neurons (Cuenca et al., 2014). Upon
activation, microglia have also been shown to initiate crosstalk with nearby Müller glia, which can
induce a gliotic response. As the initial damage-signaling attenuates, M1 microglia transition to an
M2 anti-inflammatory phenotype or undergo apoptosis to make way for new M2-polarized microglia.
The M2 transition process normally occurs by activation of intrinsic molecular switches such as
STAT6, IFR-4, PPARγ, and C/EBPB, along with miRNA-124 (Hu et al., 2014).

There are three distinct M2 expression profiles: M2a, M2b, and M2c. Beginning
chronologically, M2a microglia primarily participate in inflammation inhibition and secretion of anti-
inflammatory cytokines (IL-4, IL-13, and IL-10; (Cuenca et al., 2014)). Once this process is initiated,
M2c microglia initiate restoration and repair of damaged surviving cells by (1) secreting neurotrophic
factors such as the calcium binding protein oncomodulin (Martinez et al., 2014) and IGF-1 to induce
neurogenesis, (2) secreting vascular endothelial growth factor (VEGF) to stimulate angiogenesis,
and (3) promoting oligodendrogenesis and remyelination (Su et al., 2013). M2b microglia, which
form via the final microglia subdivision, can activate complement, are involved in the memory
immune response, and can help to stimulate or reduce inflammation as necessary. At the conclusion
of this polarization process, microglia return to a ramified resting state.

The M2 transition typically occurs under states of acute injury or stress and can help return
the retina to homeostasis. However, if the insult persists, as in the case of inherited retinal
degenerations or age-related macular degeneration, the inflammatory response becomes

19
increasingly toxic to the cells and surrounding tissue within the microenvironment (Karlstetter et al.,
2015).

Table 1.4: M2c Microglial Procedural Roles in Retinal Repair
Step Role
1 Secrete neurotrophic factors
2 Secrete VEGF
3 Promote Oligodendrogenesis and remyelination

Table 1.4: M2c Microglial Procedural Roles in Retinal Repair. Rapid microglial management
of inflammation and resolution is an important part of homeostasis. Defined structures in retinal
repair orient microglia to phased resolution of inflammatory conditions.

Müller glia are also a primary factor in inflammatory retinal homeostasis. They are extremely
sensitive to damage within the retina, not only to inflammatory factors, but also due to their keen
ability to sense changes in intercellular connections and neurotransmitter alterations. Müller cells
typically undergo reactive gliosis in response to damage, infection, or other inflammatory conditions
within the retina. Müller glia experience brief reactive gliosis and overexpress glial fibrillary acidic
protein (GFAP) (Groeger et al., 2012). After this, a number of Müller glia cells express PAX6 and
their nuclei move toward the upper INL but, in mammals, do not proliferate (Joly et al., 2011).
However, these cells express bFGF, GDNF, NGF, and PEDF, along with other growth factors, into
the injured region of the retina (Groeger et al., 2012; Forrester et al., 2020). Lastly, some Müller glia
are known to use phagocytic activity to remove the damaged retinal environment of debris post injury
(Sakami et al., 2018). Chronic gliosis causes further damage through the overproduction of
proinflammatory cytokines and nitric oxide.

20
Swift inflammation and resolution is essential for tissue repair. Persistent inflammation can
turn acute stressors into chronic disease (Nagashima et al., 2021). The order and manner in which
the interaction between immune cells and Müller glia maintain the fitness of injured tissue and
determine their outcome is still not fully understood. Dpp and the TGFβ superfamily are signaling
molecules known to activate under inflammatory conditions and we hypothesize also take part in
this process.

21

Figure 1.5: Inflammatory Retinal Homeostasis. In the coordinated system of the retina, many
cells are involved in resolving retinal inflammation. This figure shows the close connection
between neuronal cell types from the ONL to the GCL layers, and that the Müller glia cells closely
monitor these connections while aiding microglia in the removal of debris. From left to right;
patrolling ramified microglia (orange[1]), amoeboid microglia (orange), astrocyte (light green[3]),
ganglion cell[4]/bipolar cell[5]/photoreceptor[6] (blue[top to bottom]), Müller glia (dark green[7]),
inflammatory cytokines (light blue), growth factors (yellow), phagocytosis of apoptotic debris
(pink[8,9]). Adapted from (Rübsam and Parikh, 2018).

Inflammation in Drosophila is generally mediated by plasmatocytes. Under injury such as
acute bright light or UV conditions, Drosophila photoreceptors R1-8 and their associated
rhabdomere compartments are vulnerable to excitotoxic injury. This damage can be mitigated by
pros expression mediated by cone glial cells (Charlton-Perkins et al., 2017). Drosophila retinal glial

22
cells have also recently been shown to use phagocytosis under neuronal injury conditions to clear
the damaged region of debris and restore homeostatic equilibrium (Velarde et al., 2021).

Dpp/TGFβ Superfamily Pathway/Interactions
Dpp
In Drosophila, decapentaplegic (Dpp) is the earliest discovered secreted morphogen, is
required for tissue formation and advancement of the fifteen imaginal discs. The imaginal disks are
precursor tissues that will become appendages and other body parts in Drosophila. Dpp has been
found to also play a role in determining the proliferation and scope of many organs. Flies with Dpp
gene dysfunction do not grow tissues properly.

In the developing larval retina between the second and third instar, Dpp acts under a
concentration gradient, opposed by Wg, to initiate and drive the morphogenetic furrow. Dpp is a
ligand of the TGFβ superfamily. It signals through type II receptors wishful thinking (wit) and punt,
when phosphorylated and activated bind to type I receptors thickvein (Tkv) and saxophone (Sax).
These type I and II receptors phosphorylate mothers against dad (Mad) and smad on x (Smox),
which individually homodimerize and form a trimeric complex with co-smad Medea and each
independently translocate to the nucleus. It is inhibited by the I-smad, Dad before translocation into
the nucleus (Li et al., 2007; Kamiya et al., 2008; Eusebio et al., 2018).

A coordination of Tkv and Sax signaling has recently been described in the tissue damage
response of the fly intestinal epithelium, where Sax/Smox signaling mediates activation of intestinal
stem cell (ISC) proliferation in response to enteropathogen infection, while Tkv/Mad signaling
promotes the return to quiescence of these cells (Ayyaz et al., 2015). This response is coordinated

23
by the control of Tkv protein turnover in ISCs. Tkv turnover is reduced during ISC activation, allowing
for Tkv protein accumulation and replacement of Sax in Tkv/Punt complexes (Cai et al., 2019).

Figure 1.6: Canonical Dpp Pathway in Drosophila. Dpp ligand (in green) signals through type
II receptors wishful thinking (wit) and punt, when phosphorylated and activated bind to type I
receptors thickvein (Tkv) and saxophone (Sax). These type I and II receptors phosphorylate
mothers against dad (Mad) and smad on x (Smox), which individually homodimerize and form a
trimeric complex with co-smad Medea and each independently translocate to the nucleus. It is
inhibited by the I-smad, Dad before translocation into the nucleus. Adapted from (Upadhyay et
al., 2017).

BMP
Bone morphogenic proteins (BMP) are part of the TGF-β superfamily of proteins, which
includes TGF-βs, GDFs, GDNFs, activins, inhibins, and several other hormones. In the canonical
pathway, BMPs begin signal transduction by attaching to type I (BMPR1A/BMPR1B) or type II
(BMPR2) serine/threonine kinase receptors and establishing a heterotetrameric complex. The

24
constitutively active type II receptor then trans-phosphorylates the type I receptor, and the type I
receptor phosphorylates the receptor-regulated Smads (R-Smads) Smad1/5/8. Phosphorylated
Smad1/5/8 binds to co-Smad (Smad4), and the complex translocates to the nucleus where it
activates coactivators or corepressors that control gene expression. Inhibitory Smads, Smad6/7,
can competitively bind to co-Smad4 to inhibit downstream gene activation ((Kuczma & Kraj, 2015;
Fatima, 2021).
TGFβ
In the canonical pathway, the three TGFβ isoforms, precursor TGFβ1, TGFβ2, and TGFβ3,
bind to form a latent TGFβ complex ligand before being secreted. After extracellular activation, TGFβ
ligands bind to the TGFβ type II receptor (TGFβRII), which upon activation, phosphorylates and
dimerizes with the TGFβ type I receptor (TGFβRI) The constitutively active type II receptor then
trans-phosphorylates the type I receptor, and the type I receptor phosphorylates the R-Smads,
Smad2 and Smad3. The R-Smads dimerize and form a heterotrimeric complex with Smad4 which
translocates into the nucleus (Neuzillet et al., 2013).

25

Figure 1.7: Simplified Canonical BMP/TGFβ Pathway in Mammals. BMP ligand binds to
phosphorylated BMP receptor 2 and phosphorylates the receptor-regulated Smads (R-Smads)
Smad1/5/8. Phosphorylated Smad1/5/8 binds to co-Smad (Smad4), and the complex
translocates to the nucleus unless inhibited by Inhibitory Smad Smad6/7. Adapted from (Kuczma
& Kraj, 2015). TGFβ ligands bind to the TGFβ type II receptor (TGFβRII), which phosphorylates
the R-Smads, Smad2/3. The R-Smads dimerize and form a heterotrimeric complex with Smad4
which translocates into the nucleus, unless inhibited by Smad 6/7 (Neuzillet et al., 2013).

Dpp/TGFβ Superfamily Cell-Injury Interactions
Earlier literature on associations between the Dpp/TGFβ Superfamily and the retina are
extensive. It reveals a significant role for the ligand in proliferation, differentiation, maturation, and
in limiting the size of organs. However, its part in retinal injury is less well characterized. The
proceeding sections will seek to establish previous work in each category if it exists and clarify as
needed.

26
Cell Replacement
Beyond its distinct and defined role in lens and eye development, prior work conducted on
adult retinal stem cells (RSCs) has shown that secretion of BMP2/4 by the adult lens and cornea
maintain cell quiescence in vitro (Balenci et al., 2013). Work in the zebrafish has shown that
repression of TGFβ signaling by co-repressors Six3b and Tgif1 is necessary for sufficient Müller glia
proliferation (Lenkowski et al., 2013), implying a detrimental effect of TGFβ post photoreceptor
injury.

Support Cell Activation
Previous research in the chick has shown promising results in activation of retinal progenitor
cells via BMP 4 ligand to induce regeneration post retinectomy (Haynes et al., 2007). It has also
been shown that survival of retinal ganglion cells can be promoted by BMP driven phosphorylation
of Smad1/5/8 (Ueki and Reh, 2012). Additional work by the Reh lab showed that postnatal day
twelve or adult trp53 mice explanted retina exposed to EGF drive Müller glia replication via the
BMP/Smad 1/5/8 pathway (Ueki and Reh, 2013). Activation of retinal pigment cells by BMP-6 has
been shown to inhibit MAP kinase activity and oxidative stress induced injury (Chen et al., 2018).
Mammalian astrocytes have been shown to react to TGFβ signaling by inducing gliosis and
producing extracellular matrix, driving scar formation (Reichenbach and Bringmann, 2013). In the
central nervous system, TGFβ inhibition prevents scar formation (Moon et al., 2001).

27
Table 1.5: Chick BMP4, Receptor & Inhibitor Expression Timeline
Time BMP4 BMPR1A BMPRII Noggin

Dorsal/Ventral/
Posterior
Retina
Dorsal/Ventral/
Posterior
Retina
Dorsal/Ventral
/Posterior
Retina
Dorsal/Ventral/
Posterior
Retina
E4 ^/^/^ ^/^/^ ^/^/^ ^/^/^
E7 ^/+/^ ^/^/^ ^/+/^ 0/+/0
3d PR ^/+/^ ^/^/^ ^/^/^ 0/0/0
E9 +/0/+ +^/^ +/^/+ 0/0/+
5d PR ^/^/^ +/+/+ ^/^/+ ^/^/^
E11 ^/^/^ ^/^/^ 0/+/+ 0/0/0
7d PR 0/+/+ ^/^/^ +/0/+ ^/^/^

Table 1.5: Chick BMP4, Receptor & Inhibitor Expression Timeline. Expression of BMP
molecules during retina development and regeneration. Relative levels of dorsal (upper retina,
midway from lens), ventral (lower retina, midway from lens), and posterior (back of retina farthest
from lens). ^ represents high expression, + represents low expression, and 0 represents no
expression. E - embryonic day. d PR - days post retrieval. Adapted from (Haynes et al., 2007).

Mobilization of Immune Cells
After UV damage, hemocytes are recruited to the retina, where they are activated by the
Pdgf1 orthologue Pvf1 in response to activation of the Dpp signal transducer Schnurri (Shn) in
damaged retinal cells (Yao et. al., 2006; Kelsey et. al., 2012). Pvf1, in turn, induces the neurotrophic
factor Mesencephalic Astrocyte Derived Neurotrophic Factor (MANF), in hemocytes, regulating their
transition to an anti-inflammatory, pro-repair phenotype (Neves et al., 2016). This response is
conserved in mice (Neves et al., 2016), but the precise role of Dpp/Shn signaling in coordinating the
tissue repair response remains unclear.

28
Interestingly, while application of BMP-7 in prediabetic cardiomyopathy drove macrophages
to an M2 anti-inflammatory state (Urbina et al., 2014), BMP-7 was found to drive inflammatory
activation of microglia and drive retinal inflammation in an acute model of damage in vivo
(Dharmarajan et al., 2017).

Antioxidant Response
Previous studies have shown that TGFβ acts as a powerful fibrogenic cytokine and is
overexpressed in most fibrotic illnesses. Beyond consequence of downstream activation of the
phospho-Smad2/3 pathway, TGFβ signaling also inhibits mitochondrial function, reduces expression
of glutathione (GSH) and other antioxidant enzymes to drive increased oxidative stress in tissue (Liu
& Gaston Pravia, 2010; Liu & Desai, 2015).

Inhibition of Apoptosis
Work in rats cultured brain media with rat adrenal medullary PC12 pheochromocytoma
neuronal cells and mouse N1E-115 neuroblastoma cells has shown that BMP2, an upstream ligand
of phosphorylated Smad 1/5/8, acts in both a neurotrophic and neuroprotective manner against
nocodazole driven neurodegeneration (Saglam et al., 2014). When using chronic models of retinal
disease such as diabetic retinopathy disease models in aged mice, researchers have found
promising results in regression via administration of BMP-7 (Urbina et al., 2014; Sugimoto et al.,
2017).

Bridging Gaps between Model Organisms
Both Drosophila and Mus musculus have strengths and weaknesses when being used to
analyze complications of retinal diseases in a rapid and quantitative manner. While recognizing the

29
discrete advantages and boundaries of each model to investigate conserved mechanistic links, it
can be very cost and time effective to use both models simultaneously to better characterize
potential findings (Warr et al., 2018).

Drosophila Melanogaster as a Model Organism for Retinal Injury
Drosophila is an excellent model for studying the interaction between immune cells and
photoreceptors during retinal tissue repair. It is an extremely versatile model organism, granting
researchers the ability to rapidly and in real time, use RNA interference to observe how retinal tissue
homeostasis is maintained in the absence of inhibited downstream pathways. The time to generate
new fly lines of approximately twelve days is brief and rapid, the requirements for lab equipment and
storage are very cost effective, and the number and ways to manipulate animals for different
genotypes of interest is robust.

The Drosophila eye is also available and receptive to acute UV irradiation during the discrete
window of observation twenty-four hours post pupation when retinal cells are post-mitotic (Jassim
et al., 2003), and many other well characterized retinal damage models are also available (Kang
and Ryoo, 2009; Catalini et al., 2021; Velarde et al., 2021). Genetic or UV-C induced damage
directed to the post-mitotic Drosophila pupal retina can induce reproducible, quantifiable damage
that persists to adulthood and is sensitive to genetic perturbations, allowing dissection of pathways
that mediate photoreceptor apoptosis and control survival (Jassim et al., 2003; Kelsey et al., 2012;
Neves et al., 2016). Persistent DNA damage induced by UV has been shown to promote initiator
caspase activity in photoreceptors and thus apoptosis (Brodsky et al., 2000; Jin et al., 2000; Ollmann
et al., 2000; Sogame et al., 2003; Zhou and Steller, 2003; Hay and Guo, 2006; Steller, 2008; Denton
et. al, 2013; White et al., 2017).

30

Seventy five percent of all human diseases have genetic homologues in flies (Mirzoyan et
al., 2019). Lastly, like the mouse and human retina's reliance on microglia under homeostatic
equilibrium, the eyecup is mostly dependent on innate immunity for defense from pathogens, injury,
toxins, and foreign objects (Akhtar ‐Schäfer et al., 2018).

M. musculus as a Model Organism for Retinal Injury
While Drosophila models have significantly advanced study of specific pathways within
the eye, some investigators assert mammalian models are required to construct a robust
correlation with human retinal diseases. Due to having ninety nine percent genetic similarity,
comparable anatomy and physiology to humans, the mouse is one of the most common
mammalian models in retinal research (Krebs et al., 2017). It is small, easily managed, and with
proper facilities, maintained at relatively low cost (Bryda, 2013). Due to their extensive use in
biology, a number of mutant strains and genetic strains are readily available. Additionally, a large
and ever-growing variety of publicly available genetic databases using mouse disease models
such as the Mouse Genome Database (MGC) exist and are easily accessible (Blake et al., 2021).
Furthermore, clinical trials expect mammalian data to decide safety and effectiveness, and mice
represent the bulk of such data sets (Guo et al., 2019).

Limitations of Flies & Mice as Model Organisms
While both flies and mice organisms offer significant knowledge into diverse biological
activities within the injured retina, each model has their unique drawbacks. While the fly shares
many homologous pathways to humans, the mouse likeness is significantly more robust.
However, the mouse retinal disease models have of yet been unable to replicate all forms of

31
retinal disease, or even more common ones such as age related drusen accumulation (Cheng et
al., 2020). Neither fly nor mouse has a macula, or a region of highly elevated cone density that
would be like the fovea (Volland et al., 2015). Additionally, often neither model, nor others
commonly used - such as rat, rabbit, or dog, perfectly translates to human diseases, regardless
how careful or extensive the study. Eighty nine percent of novel drugs tested with mice fail in
human trials, with more than half due to unexpected toxicity (Van Norman, 2019). There is a
tremendous need for models with better translatability to human retinal diseases.

Translational Research
The main strength of using two evolutionarily distinct models such as the fly and mouse is in
deciphering conserved mechanisms and homologous gene networks that are essential in flies, mice,
and humans. This approach can both provide cost savings by enabling reduced usage of more
costly, larger primate models, while still obtaining results that are translatable to a wide variety of
species along the evolutionary chain. Importantly, translational drug development requires room for
failure. Using multiple, less time consuming and expensive animal models can enable increased
chance of a successful discovery and potential reproducibility in humans (Stieglitz, 2020).
Increasing throughput to drive successful, translational discoveries can be approached in
several different ways. Burgeoning retinal disease model systems such as human retinal organoids
and organ-on-a-chip technology provide tremendous hope for the future. Recent work merging these
two models enables the ability to monitor the interaction of mature photoreceptor segments, RPE,
and vasculature-like perfusion in vitro (Achberger et al., 2019).

32
Chapter II: Dpp/TGFβ-superfamily play a dual role in Mediating Retinal Damage Response

The following chapter is adapted from (J. Kramer et al., 2021).
Hemocyte derived Dpp controls the damaged retinal tissue response in Drosophila
Our previous findings had identified Shn as a critical mediator of the retinal damage response
in flies (Kelsey et al., 2012), yet the role of Dpp signaling in that response remained unclear. To start
assessing a possible role for Dpp signaling components in retinal apoptosis, we performed genetic
interaction experiments using a previously characterized fly line overexpressing a constitutively
active form of the Jun N-terminal Kinase Kinase (JNKK) Hemipterous (Hep) under the control of the
photoreceptor and cone cell driver Sep-Gal4 (Therrien et al., 1999; Jasper et al., 2002; Kanda et al.,
2004; Luo et al., 2007; Kelsey et al., 2012). In this line (Sep-Hep
ACT
), Hep
ACT
expression is initiated
during development in postmitotic photoreceptor and cone cells in the third instar larval eye disc,
inducing photoreceptor apoptosis through activation of the pro-apoptotic gene hid (Luo et al., 2007;
Nielsen et al., 2008).
When Sep-Hep
ACT
was crossed to flies expressing Sax-RNAi or Smox-RNAi under the
control of UAS, a significant increase in surviving photoreceptors compared to control flies (crossed
to UAS::mCherry-RNAi) was observed (Fig. 2.1A,A’). Over-expression of the negative feedback
inhibitor of Dpp signaling, Daughters against Dpp (Dad; Tsuneizumi et al., 1997; Kamiya et al.,
2008), also promoted survival. Conversely, we found that knocking down Tkv and Mad resulted in a
significant decrease in surviving photoreceptors (Fig. 2.1A,A’).

33

Figure 2.1: Dpp inhibition controls JNK and UV driven apoptosis in Drosophila. (1A,A’)
Representative images and quantitation of ratio of surviving photoreceptors to total in
w;Sep::Gal4,UAS::HepACT flies crossed with RNAi lines of each genotype. Representative
images and quantification of adult GMR::Gal4; /UAS::RNAi progeny following knockdown of is
compared through ratio of UV-exposed to unexposed eye (1B-B’). Representative images and
quantitation of the pMad (in maturing GMR::Gal4;Dad-nGFP/UAS::RNAi) and nuclear Smox (in
GMR::Gal4;Smox::FLAG::GFP/UAS::RNAi) (1D,D’) expression in Elav+ (1C,C’) progeny eye
with and without irradiation at 27, 30, 36 and 48hours post UV. Representative images and

34
quantitation of total DCP1+ Elav+ cells (1E,E’) or Dad-nGFP+/Elav+ cells (1E’’,F’”) in
GMR::Gal4;Dad-nGFP/UAS::mCherryRNAi progeny with and without irradiation at various time
points post exposure. Representative images and quantitation of total DCP1+ Elav+ cells (1F,F’)
or DadnGFP+Elav+ cells (1F”,F’”) in maturing GMR::Gal4;Dad-nGFP/ Hml::DppRNAi progeny
compared to controls. Scale Bar: 20 μm. Error bars indicate s.e.m. P-values from Student’s ttest.
*p<0.05, **p<0.01, **p<0.001.

To assess whether this antagonistic effect of Sax/Smox and Tkv/Mad signaling was also
observed during UV-induced cell death, we used an assay in which the headcase of pupae is
removed at 24 hours after puparium formation, and animals are irradiated on one side with 17.5
micro Joules of UV-C radiation (Jassim et al., 2003; Luo et al., 2007). In this model, hemocytes are
attracted to the retina after UV-C induced damage and are critical to limit excessive cell death. When
animals emerge after development, the extent of apoptosis can be measured by quantifying the size
of the irradiated eye in relation to the non-irradiated control eye. We used the glass multimer reporter
driver (GMR-Gal4) (Moses and Ruben, 1991) to express RNAi targeting Dpp pathway components
in all postmitotic cells of the retina, and found that Sax or Smox knockdown inhibits UV-induced
tissue loss, while Tkv or Mad knockdown promote it (Fig. 2.1B,B’). A similar protective effect was
observed when we used a hemocyte-specific driver (hemolectin::Gal4; Goto et al., 2003) (Fig. 2.1B)
in support of previous studies from our lab and others demonstrating that hemocytes secrete Dpp
(Ayyaz et al., 2015; Dey et. al., 2016; Eleftherianos et. al., 2016; Chakrabarti et. al., 2016; Tian et
al., 2017; Gervais and Bardin, 2017). These data further support a model in which Sax/Smox
signaling promotes apoptosis and Tkv/Mad signaling inhibits apoptosis in the retina.
To directly investigate the effects of Dpp signaling in photoreceptors, we irradiated
Drosophila pupa, waited an additional 24 hours as previously described (Jassim et. al., 2003; Kelsey
et. al., 2012), and dissected retinae during the post maturation stage (27, 30, 36 and 48 hours post
UV exposure; equivalent to 51, 54, 60 and 72 hours post puparium formation) for analysis. We used

35
immunohistochemistry against phospho-Mad (pMad) to assess activation of the Tkv/Mad pathway
in Elav+ photoreceptors (Suh et. al., 2002) during each post maturation time point (Fig. 1C). After
UV treatment, pMad activity in Elav+ cells peaked at 36 hours (Fig. 1C’). We further assessed Smox
nuclear localization using a Smox::GFP knockin line in which GFP-tagged Smox is expressed from
the endogenous Smox locus (Spokony and White, 2013) in non UV control condition (Fig. S3.1B,B’).
Nuclear localization of Smox represents activation of the Sax/Smox response (Ayyaz et al., 2015).
Compared to pMad, Smox undergoes nuclear translocation at the earliest time point we assessed,
27 hours post UV exposure (Fig. 1D,D’). To assess whether the kinetics of Sox/Smox and Tkv/Mad
activation correlates with the induction of apoptosis, we stained for cleaved Drosophila caspase-1
(DCP1), a short-prodomain caspase and crucial driver of cell death (Xu et al., 2006). When
compared to control animals, DCP1 activity peaks at 36 hours post UV exposure (Fig. 1E,E’).
Expression of a Dad-nGFP reporter (Ninov et al., 2010; Malzer et al., 2018) also peaked in Elav+
cells at 36 hours following irradiation (Fig. 1E’’,E’’’).

36

Figure 2.2: Following UV driven whole eye radiation damage, Sax and Smox RNAi inhibit
apoptosis, while Mad and Tkv RNAi promote it. Representative images and quantitation of
total DCP1+ Elav+ cells (2-A’; 2B-B’) or DadnGFP+ Elav+ cells (2A’’-S2A’’’; 2B’’-S2B’’’) in
maturing GMR::Gal4;Dad-nGFP/ Sax-RNAi and SmoxRNAi progeny compared to controls.
Representative images and quantitation of total DCP1+ Elav+ cells (2C-C’; 2D-D’) or DadnGFP+
Elav+ cells (2C’’-S2C’’’; 2D’’-S2D’’’) in maturing GMR::Gal4;Dad-nGFP/ Tkv-RNAi and Mad-RNAi
progeny compared to controls. Representative images and quantitation of total DCP1+ Elav+
cells (2-E-E’; 2F-F’) in maturing Sep::Gal4;/ Smox-RNAi and Mad-RNAi progeny compared to

37
controls. Scale Bar: 20 μm. Error bars indicate s.e.m. P-values from Student’s t-test. *p<0.05,
**p<0.01, **p<0.001.
These kinetics are consistent with the hypothesis that Mad activation induces Dad
transcription to inhibit Sax/Smox signaling in the eye, thus limiting apoptosis. To test this hypothesis,
we examined the effects of Tkv/Mad and Sax/Smox perturbations on UV-induced apoptosis.
Knocking down Dpp in immune cells significantly reduced apoptosis (as determined by anti-DCP1
staining) when compared to controls (Fig 2.1F,F’), while also resulting in a significant increase in
Dad activity at 48 hours post UV (Fig. 2.1F’’,F’’’).
Knockdown of Dpp in immune cells under non UV state did not significantly differ from control
levels of apoptosis (Fig. S3.1C,C’). Knockdown of Sax or Smox, on the other hand, resulted in a
significant decrease in apoptosis at the 30 and 36 hour time points (Fig. 2A,A’-B,B’) compared to
control, associated with a significant increase in Dad-nGFP expression (Fig. 2A’’-B’’)(Fig. S3A,A’-
B,B’). Knocking down Tkv or Mad, in turn, significantly increased the number of DCP1+ cells at
every time point (Fig. 2C,C’-D,D’), and decreased Dad-nGFP reporter activity (Fig. 2C’’-D’’; S3C,C’-
D,D’). These observations were recapitulated when Tkv/Mad and Sax/Smox perturbations were
targeted directly to photoreceptors and cone cells by Sep::Gal4 rather than to the whole retina (Fig.
2E,E’-F,F’; S4). DCP1+ cell count under control Sep::Gal4 had similar results to GMR::Gal4 under
UV and non UV states (Fig. S3D,D’,D’’).
Our data suggest a bimodal response of photoreceptors to Dpp after UV irradiation that is
controlled by Sax and Tkv signaling. Additionally, suppression of Dad expression by Smox and
subsequent activation of Dad expression by Mad contribute to the regulation of photoreceptor cell
death and survival.

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BMP and TGFβ signaling are activated in a time dependent manner following retinal stress
and damage
As the Drosophila visual system has many similarities with the mammalian retina (Sanes and
Zipursky, 2010), we next asked whether the bimodal response of the two arms of the Dpp pathway
to damage in the Drosophila eye is conserved in mammals. We specifically focused on the
Transforming Growth Factor beta (TGFβ) superfamily, which has a close homology to the Drosophila
Dpp canonical pathway and has shown to play a role in the immune response in multiple systems
(Upadhyay et al., 2017; Sanjabi et al., 2017). Two important members of the family, Bone
Morphogenic Protein (BMP) and TGFβ, share close homology with Dpp (Hamaratoglu et al., 2014;
Upadhyay et al., 2017). Following binding-induced TGFβ type 1 receptor phosphorylation, they
trigger phosphorylation and nuclear translocation of Smads (Smad 1/5/8 for BMP pathway or Smad
2/3 for TGFβs), and transcriptional activation of Smad target genes. Based on our studies in
Drosophila, we hypothesized that the TGFꞵ/Smad2/3 arm of the pathway is responsible for the
inflammatory or apoptotic response, and the BMP/Smad1/5/8 arm for the anti-inflammatory pro-
repair response.
To test this hypothesis, and to elucidate the dynamics of TGFβ superfamily activation, we
used light-induced photoreceptor stress and damage models previously used by various labs
including ours (Grim and Remé, 2013; White et al, 2007; Neves et. al., 2016). Briefly, C57BL/6 mice
were exposed to high-intensity (10,000 lux) light for 1.5 hours to induce retinal stress without overt
retinal apoptosis. We then analyzed the phosphorylation of the mammalian homologues to Mad and
Smox, Smad1/5/8 and Smad2/3 (Kashima and Hata, 2017) comparing non-light exposed conditions
to control (Fig. S3.5A), 0, 6, 12, 24 and 36 hours post exposure. Upon exposing 3.5 month old
C57BL/6 mice to light, we observed elevated levels of phosphorylated Smad 1/5/8 (BMP response)
in the inner nuclear layer (INL) of the retina, starting at 6hr post-light exposure, and peaking at 12

39
hours post exposure (Fig 2.3A,A’). Phosphorylation of Smad 2/3 (TGFβ activation) peaked earlier,
at 6 hours post exposure (Fig 2.3B,B’).
Next, we asked if the response differs in the BALB/c mouse model, which are more sensitive
to light damage and exhibit widespread apoptosis in the retina after light exposure. BALB/c mice
were exposed to 5,000 lux light for 1hr to induce apoptosis as previously shown (Neves et al. 2016).
TUNEL assay was performed on retina 24 hours post exposure to confirm significant apoptosis (Fig.
S3.5B). Interestingly, even prior to light exposure, BALB/c mice exhibit persistent TGF-β activity
(pSmad 2/3) in the inner nuclear layer, suggesting substantial retinal stress under normal light
conditions (Fig 3C). Immediately after light exposure (0 hours), this activity increases transiently, but
by 6 hours, the pSmad2/3 signal returns to pre-light exposure levels and is absent once apoptosis
sets in at 24 hours (Fig. 2.3C’). As opposed to C57BL/6 mice, BALB/c mice exhibited no phospho-
Smad1/5/8 (BMP) activity in the inner retina either under control conditions or after light exposure at
any of the time points assessed in our study (Fig. S3.5D). These studies suggest that TGFβ signaling
may be involved in the early inflammatory response, while BMPs may play a role in a later protective
anti-apoptotic pro-repair response which BALB/c mice lack.
We and others have previously shown that the retinal damage response relies on factors
secreted by activated immune cells (Minhas et al., 2016; Okunuki et al., 2018; Akhtar-Schafer et al.,
2018; Okunuki et al., 2019). We next tested whether the light stress-induced BMP response is
impacted when immune cells are absent. We used CD11b::DTR mice wherein myeloid cells can be
inducibly ablated using intraperitoneally injected diphtheria toxin (DT) (Frieler et al., 2015;
McPherson et al., 2019). We have previously shown that light stress results in photoreceptor
apoptosis in CD11b::DTR mice despite them being on C57BL/6 background (Neves et. al. 2016).
DT induced immune cell loss in CD11b::DTR mice also resulted in loss of pSmad 1/5/8 in the inner
retina under apoptosis inducing light-damage condition (Fig. 3D,D’ in comparison to PBS injected
light damaged control retinas where phospho-Smad 1/5/8 activation was seen in CRALBP+ Müller

40
glial cells in (Fig. S3.5E). These studies suggest that immune cells participate in the damage-
induced BMP response in mammalian retinas.

Figure 2.3: BMP and TGFβ activation occurs in a time dependent manner following light
stress and damage. Representative retinal images and quantitation of control dark adapted 3.5
m/o C57BL/6 mice, or 0, 6, 12, and 24 hours post exposure to 10,000 lux for 1.5 hours stained
for pSmad 1/5/8 (in green, 3A-A’) and p-Smad 2/3 (in green,3B-B’). Similar analysis of pSmad
2/3 in light-damaged BALB/c mice retinal sections (3-C). Representative retinal images and
quantitation in 3.5 m/o CD11b::DTR mice treated with either diphtheria toxin or PBS control and
exposed to 10,000 Lux. Sections were stained pSmad 1/5/8 (green) (3D-D’). Arrows (red)
highlight pSmad expression in the inner retinal in all images. Scale Bar: 30 μm. Error bars indicate
s.e.m. Pvalues from Student’s t-test. *p<0.05, **p<0.01, **p<0.001. PE = post-exposure. DAPI
(blue) marks nuclei in all panels.

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Modulation of the BMP signaling pathway protects against damage induced retinal apoptosis
We hypothesized that BMP mediates an anti-inflammatory protective retinal damage
response and sought to test this hypothesis by asking whether modulation of BMP signaling would
impair retinal repair. We used the NMDA excitotoxic damage model previously used by multiple labs
to cause inner retinal damage (Karl et al., 2008; Lamba et al., 2008; Kuehn et al., 2017; Lambuk et
al., 2019) in addition to the light damage model. The excitotoxin NMDA damages the ganglion and
amacrine cells in the retina and has been used to model retinal damage due to glaucoma, retinal
ischemia and diabetic retinopathy (Niwa et al., 2016; Maekawa et al., 2017; Pirmardan et al., 2018;
Christensen et al., 2019). To directly monitor BMP activity, we used BRE-eGFP transgenic mice.
These mice report the transcriptional response of BMP-Smad activation through the BMP response
element (BRE) (Javier et al., 2012). In these mice, GFP expression was observed in CRALBP+
Müller glia following NMDA exposure (Fig. 2.4A).
To modulate the BMP signaling pathway, mice were injected intravitreally with either
recombinant BMP4 or the small molecule BMP inhibitor Dorsomorphin (Yu et al., 2007; Ueki et al.,
2012). Upon comparing various treatments, we confirmed that GFP activity increases following
BMP4 treatment and reduces with Dorsomorphin (Fig. 2.4A,A’). TUNEL staining at 48-hours post-
NMDA injection revealed that inhibition of BMP signaling increased apoptosis, while
supplementation of BMP4 inhibited NMDA-induced apoptosis (Fig. 2.4B,B’). To assess whether this
was a conserved retinal damage response or specific to NMDA damage, we assessed pathway
modulation under light-induced photoreceptor damage conditions ( 20,000 Lux for 2 hours (Zhang
et al., 2015)). TUNEL staining at 48 hours post exposure revealed that inhibition of BMP signaling
increases apoptosis and supplementation of recombinant BMP4 reduces it (Fig. 2.4C,C’), similar to
our observations in the NMDA damage model.

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Inhibition of TGFβ signaling pathway protects against damage induced retinal apoptosis
Since our data suggested that TGFβ may be involved in the earlier inflammatory response
to damage, we tested if inhibiting the pathway using the small molecule TGFβ inhibitor SB431542
(Tappeiner et al., 2016) would be protective. We carried similar NMDA and light-damage
experiments as above and co-treated the mice with 25 μM of SB431542 intravitreally. In both
damage models, we observed that inhibition of the TGFβ pathway reduced retinal apoptosis
compared to control mice (Fig 2.4B,C). Interestingly, we also observed a small reduction in BRE-
GFP activity following TGFβ inhibition (Fig 4A) suggesting cross-regulation between the pathways.

Figure 2.4: Modulation of the BMP/TGFβ signaling pathway affects tissue damage induced
retinal apoptosis and immune activation. 3.5 m/o BRE-eGFP reporter mice intravitreally
injected with 25mM NMDA were treated with either; PBS control, mBMP4, Dorsomorphin (BMP
Inhibitor), or SB431542 (TGFβ Inhibitor). At 48 hours, BRE-GFP (green) colocalization was
assessed and quantified with Müller glial marker, CRALBP (white) and plotted following various
treatments in comparison to control NMDA (4AA’). TUNEL assay (red) was performed under
above conditions, with total TUNEL+ cells plotted in comparison to control (4B-B’). Effects of
various treatments tested in light damage conditions following 20,000 Lux exposure for 1.5 hours

43
with retina collected 48 hours post exposure. TUNEL assay was then performed and quantified
in comparison to control (4C-C’). Retinal tissue analysis for immune markers IBA1 (white) and
CD68 (green), with ratio of ramified morphology immune cells over total compared between
conditions (4D-D’). Total number of activated CD68+ immune cells was also compared between
treatment conditions (4D-D’’). Scale Bar: 20 μm (4-A). 30 μm (4B-D). Error bars indicate s.e.m.;
P-values from Student’s t-test. *p<0.05, **p<0.01, **p<0.001. DAPI (blue) marks nuclei in all
panels.

Microglia activation can be modulated by manipulating BMP/ TGFβ pathways
Lastly, we asked whether microglia activation can be biased by modulation of TGFβ and
BMP pathways. Recently, morphological changes in microglia have been shown to reflect activation
(Karperien et al., 2013). A change from a ramified non-activated morphology to an amoeboid shape
signifies activation, and the ratio of the two morphologies can be used as a measure of relative
activation. Retinal tissue was stained for the pan-microglia marker IBA1 and the inflammatory
activation marker CD68. We carried out fractional analysis comparing the ratio of ramified over total
(ramified and amoeboid microglia) between conditions (Fig. 2.4D) to determine the fraction of non-
activated microglia. Under PBS injected control conditions, all microglia have ramified morphology
(Fig. S3.5C) and NMDA damage leads to over sixty percent of microglia changing to amoeboid
morphology (Fig. 2.4D’). We found inhibition of BMP (Dorsomorphin treatment) led to further
reduction in the ramified fraction (less than twenty percent), indicating increased activation, whereas
supplementation of BMP4 or inhibition of TGFβ led to increases in ramified microglia, indicating
reduced activation (Fig. 2.4D’). Furthermore, the morphological changes in microglia closely
correlated with CD68 expression (Fig. 2.4D’’).

Discussion
Our studies identify the morphogen Decapentaplegic (Dpp) and its mammalian homologues
BMP/TGFβ as important regulators of retinal tissue survival post injury. Our findings suggest that in

44
the fly, Dpp is secreted from hemocytes that previous studies have shown to be drawn to the injury
site (Kelsey et al., 2012). Hemocytes have also been shown to secrete Dpp in the fly intestine and
embryo after injury (Ayyaz et al., 2015; Eleftherianos et. al., 2016). We find that the response to
these hemocyte derived ligands in the retina is dynamic: in elav+ cells, Smox nuclear translocation
is detected first after damage, while Mad phosphorylation occurs later and correlates with Dad::GFP
expression, consistent with Mad-mediated induction of Dad (Tsuneizumi et al., 1997; Inoue et
al.,1998; Hamaratoglu et. al., 2014). Mad phosphorylation also correlates with peak levels of
apoptosis, which are detected at 36 hours post UV, indicating that Mad activity is associated with
the apoptotic state. Since loss of hemocyte Dpp, or of retinal Sox and Sax all result in reduced tissue
loss, while loss of Tkv and Mad increased tissue loss, we propose that hemocyte-derived Dpp
induces retinal apoptosis by activating Sax/Smox signaling, while the later engagement of Tkv/Mad
signaling is required to downregulate the apoptotic response. The selective engagement of
Sax/Smox and Tkv/Mad signaling at different timepoints in the injury response is reminiscent of a
similar dynamic observed in intestinal stem cells after bacterial infection (Ayyaz et al., 2017; Cai et
al., 2019).
As Dad is an inhibitory Smad, we hypothesize that Dad induction by Mad is required as a
negative feedback signal to repress Smox signaling (S3.1-E). Dpp has been similarly shown to drive
a pro-apoptotic response under damage in other tissues such as ovarian somatic cells, leg disk and
retinal glia (Kang et. al., 2018; Manjón et. al., 2007; Velarde et. al., 2020). In ovarian cells,
overexpression of Dpp was shown to drive activation of the pro-apoptotic genes reaper (rpr) and
head involution defective (hid) (Kang et. al., 2018). Additionally, pSmox has been shown to induce
cell and stage specific apoptosis of larval neurons in other Drosophila models (Wang et. al., 2019).
Interestingly, knocking down Sax and Smox in the eye increased Dad-nGFP expression, while, as
expected, knocking down Tkv and Mad significantly reduced Dad-nGFP activity. Sax/Smox signaling
may thus partially repress Tkv/Mad signaling in the early phase of the injury response.

45
We also identified dynamic changes in the phosphorylation of the downstream effectors of
the mammalian homologues to Dpp, BMP & TGFβ, in our mouse retinal stress and injury models.
Under conditions of light-induced retinal stress based on our prior studies (Neves et al 2006), we
observed an initial TGFβ activation represented by an early peak of pSMAD2/3 at 6 hours post
exposure. This is followed by a peak in BMP (pSMAD1/5/8) activity 6 hours later. Intriguingly, we
observed an early Smox followed by a delayed Mad phosphorylation in UV-damaged flies as well,
suggestive of conserved arms of the pathway between the two species. These findings provide
further support to homology between pSmad2/3 / Smox, and Smad1/5/8 / Mad, also shown in
Drosophila heart injury model or other species such as Haemonchus contortus (Goldstein et al.,
2010; Li et al., 2020). We additionally observed that in our light damaged BALB/c mouse model
which bears a RPE65 SNP associated with increased susceptibility to light stress (Grimm et al.,
2000; Wenzel et al., 2003), pSmad 2/3 activity exists under control conditions suggesting this arm
of pathway is driven by retinal stress. Interestingly, there was no detection of Smad1/5/8 activity at
any time point following light exposure. These results suggest that the BMP arm is protective and
lack of activity of this arm of the pathway is associated with increased apoptosis.
Our data also demonstrate that knockdown of CD11b positive immune cells in the retina
significantly reduces the phosphorylated Smad 1/5/8 activity in Müller Glia during the light induced
damage response (Fig. S3.5F). This suggests that immune cells play a role in activation of the
mammalian homologue of Tkv/Mad pathway, BMP, which is not activated in their absence. Immune
cells may play a role in either directly secreting BMPs (Yousef et al., 2016) or by promoting BMP
induced Müller glial activation (Dharmarajan J. et al., 2017).
Supplementation of the mammalian homologues of Dpp, BMP4 and inhibition of TGFβ
through the ALK inhibitor SB431542, reduce apoptosis after light and NMDA induced injury.
Conversely, downregulation of BMP4 through the inhibitor Dorsomorphin increases both apoptosis
and total CD68+ cells, a known marker of immune cell mediated inflammation after injury (Chistiakov

46
et al., 2016). This suggests the parallel findings between Drosophila and mammalian homologues
are shared in both light induced and NMDA driven damage models. Previous work has found similar
findings in damaged retinal ganglion cells (Ueki and Reh, 2012; Ueki and Reh, 2013). Additionally,
NMDA damaged Müller glia from zebrafish, rodent and chick Müller glia derived progenitor cells
(MGPCs), showing phosphorylated Smad 1/5/8 promotes regeneration and phosphorylated Smad
2/3 inhibits it (Saika, 2005; Lenkowski et. al., 2013; Todd et. al., 2017).
The opposing effects of apoptotic driving TGFβ/Smad2/3 and apoptosis inhibiting
BMP4/Smad1/5/8 on retinal injury are also correlated with different downstream target genes.
BMP2/4 activity is closely associated with anti-apoptotic genes id1/2/3, sox9, ihx2, and wnt
10a/11/14/7b (Genander et. al., 2014). TGFβ is associated with TGFβ-inducible early response gene
1 (TIEG1), Bcl-2-interacting mediator of cell death (Bim), death-associated protein kinase (DAP-
kinase), TGF β-induced Gene Human Clone 3 (BIGH3) and repression of id2 (Jang et al., 2002;
Bender et al., 2004; Yu et al., 2008; Betts-Obregon et al., 2016). Additionally, its been shown that
TGFβ driven activation in injury can also drive delayed secondary necrosis in tissue, driving the
cellular membrane permeable to macromolecules, inducing delayed inflammation and leading to the
activation of cleaved caspase-1 (Rock and Kono, 2007; Appelbaum et. al., 2017). However, not all
previous literature on TGFβ defines it as pro-apoptotic within the retina. Recent work ablating TGFβ
signaling in retinal microglia using tamoxifen induced Tgfbr2
flox/flox
mice lead to increased retinal
degeneration and Müller cell gliosis (Ma et al., 2019). Also, AAV8-TGFβ1 was recently shown to
promote cone survival in rd1 mice (Wang et al., 2020). This protective response required Tgfbr1 and
Tgfbr2 activity or microglia which express them. This suggests that while TGFβ activity downstream
of Tgfbr2 on microglia drives secretion of pro-survival effectors, other cell types including Müller glia
may have different independent roles especially in acute damage.

47
These results highlight the critical role of Dpp/BMP/TGFβ in regulating retinal tissue repair.
Future studies targeting downstream aspects of the mammalian pathway may have therapeutic
implications to delay or repair the onset of inflammation and allay retinal damage.

Materials and Methods
Mice
All mice used in the described studies were housed and bred at the Association for
Assessment and Accreditation of Laboratory Animal Care International accredited vivarium of the
Buck Institute for Research on Aging, in a specific-pathogen-free facility, or in the UCSF Laboratory
Animal Resource Center, in individually ventilated cages on a standard 12:12 light cycle. All
procedures were approved by the Buck Institute Institutional Animal Care and Use Committee or
UCSF IACUC Animal Care Committee.
Intraocular Injections in Mice
For intravitreal injection, recombinant proteins or compounds in 1-μl volume were injected
into the right eye using a graduated pulled glass pipette and a wire plunger (Wiretrol II, 5-0000-2005,
Drummond Scientific Company) or directly by using a Hamilton 10uL pipette following isoflurane
anaesthesia.

Light and NMDA damage in mice
C57BL/6, BALB/c, or BRE-eGFP 4x backcrossed with C57BL/6 background were dark
adapted for 18 hours, then intravitreally injected of control PBS or 25 mM NMDA, and recombinant

48
protein of either; mBMP4, Dorsomorphin (BMP Inhibitor), or SB431542 (TGFβ Inhibitor). Animals
were then exposed to either 5000-20,000 lux for 1-1.5 hours, or 25 mM NMDA. In case of light
damage, mice were allowed to recover from anesthesia, returned to their cages, and housed in
darkness until analysis. Retinal tissue was collected under control state, 0, 6, 12, 24 or 48 hours
post exposure.
UV damage in Drosophila pupae retina and larvae
Pupae retinas were exposed to 17.5 microJoules of UV-C radiation 24 hours post puparium
formation and either collected 5 days post adulthood or 24, 27, 30, 36, and 48 hours post exposure.
Histological analysis, imaging, and quantification methods
Retinal sections and macrophages were analyzed by immunohistochemistry (IHC) and
other histological methods and imaged using a LSM 700 confocal laser-scanning microscope,
images were processed sequentially on separate channels. All images were used for
quantification purposes and processed with Adobe Photoshop software, ImageJ, Imaris 9.5.1,
ZEN 3.2 and LAS X software. Eyes were fixed with 4% formaldehyde, PBS washed and placed
in progressively increasing concentrations of PBS/Sucrose solution (5, 10, 15, 20%), mounted in
O.C.T.(TissueTek) compound in -80 degrees C overnight, sectioned into 10 micron tissue on
slide, prepared as (Lamba et al., 2010) and analyzed with the following antibodies: Elav-9F8A9
(1:200, rat, DSHB), pMad-EP823Y (1:300, rabbit, abcam), DCP1-Asp216 (1:100, rabbit, Cell
Signaling), pSmad 1/5/9 (1:300, rabbit, Cell Signaling), pSmad 2/3 (1:300, rabbit, Cell Signaling),
GFP (1:1000, rabbit, GeneTex), CRALBP (1:200, mouse, Santa Cruz), TUNEL, IBA1 (1:200,
rabbit, Abcam), and CD68 (1:100, rat, BioLegend).

49
Statistical analysis
All counts are presented as average and standard error of mean (SEM). Statistical analysis
was carried out using Microsoft Excel or GraphPad Prism 8.0.1, and Student’s t test or two-way
analysis of variance (ANOVA) was used to determine statistical significance, assuming normal
distribution and equal variance.

50
Chapter III: Conclusion and Final Thoughts
Hemocyte derived Dpp controls the Drosophila damaged retinal tissue response
I have demonstrated through listed data that hemocyte derived Dpp controls the
Drosophila damaged retinal tissue response. Due to 95% of hemocytes in Drosophila being
plasmatocytes, plasmatocytes are likely a significant source of Dpp expression under eye
damage. While hml(Δ)-Gal4 used in this work functionally targets plasmatocytes for gene
knockdown, to fully confirm this statement, one could also use the Peroxidasin-GAL4 (Pxn-GAL4)
driver with the UAS-Dpp-RNAi/GAL4 system as another well characterized plasmatocyte driver
line to better segregate Dpp loss of function between plasmatocyte and hemocyte drivers (Shin
et al., 2020). Ultimately, the finding of immune cells expressing Dpp growth factor to promote
retinal tissue resilience and recovery demonstrates the crucial importance of a sensitive and
robust immune system that is constantly surveilling the retinal environment for stressors. Doing
so and acting promptly to resolve them is a crucial part of retinal homeostasis.

Under UV radiation damage, Sax/Smox inhibit apoptosis and Mad/Tkv promote damage
Another important finding of this research is the bimodal Dpp downstream element driven
response under direct eye UV irradiation using both whole eye and photoreceptor only
knockdown. Our data suggests that photoreceptors are a significant receptor source of the Dpp-
Mad/Smox paradigm, as we see similar results under both whole eye and photoreceptor only
knockdown. To better clarify this result, we could further examine the retinal injury under loss of
function RNAi condition using the UAS-Dpp-RNAi/GAL4 system through drivers dedicated to
each primary cell type that makes up the ommatidium unit eye. Our data suggests that the
photoreceptor is a vital part of the eye in both flies and mice, and the immune cell retina damage

51
response paradigm may take its role into account. Further examination of this question could
result in useful diagnostic tools in other homologous animal models and in people.

BMP4 activation/TGFβ inhibition protect against damage induced retinal apoptosis
An additional discovery from our work was identification, in multiple age, sex, and species
of mouse retina, of the correlation between injury induced apoptosis and mouse BMP-4 activation
and inhibition during the retinal damage response. We have found that timing is an important
factor in resolving retinal injury. After retinal injury and ligand activated response, Sax/pSmox and
approximate homologues TGFβR2/pSmad2/3 activate first in damaged tissue, and correlate with
an increase in retinal apoptosis. This is followed sometime later by Tkv/pMad and its homologues
BMPR2/pSmad 1/5/8 driving an increase in photoreceptor and retinal survival post injury. I further
speculate that timed inhibition and supplementation of TGFβ superfamily complexes into the eye
through non-invasive means such as eye drops could be a potential future remedy to acute
injuries from the sun and sudden light exposure or other forms of acute injury. I hope that such a
world is not far off.

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Figure 3.1: Diagram of Dpp and TGFβ/BMP signaling act as central controllers of
photoreceptor death and survival after acute damage within the retina. Based on our data,
under immune cell secreted Dpp/TGFβ/BMP ligand activation, Sax/pSmox ~ TGFβR2/pSmad2/3
activate first and correlate with an increase in retinal apoptosis, followed by Tkv/pMad ~
BMPR2/pSmad 1/5/8 driving an increase in retinal survival post injury.

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Supplemental Information

54

Supplementary Figure 3.1: Representative images and quantitation of control pMad+
Elav+ cells. Images under all timepoints of control pMad+ Elav+ cells and non-UV WT genotype
is compared (S1-A). Control nSmox+ Elav+ cells under all timepoints and non-UV WT genotype
is compared (S1-B). GMR::Gal4;Hml::DppRNAi progeny total DCP1+ Elav+ cells is compared to
control no UV animals (S1-C). Sep::Gal4;mCherry-RNAi control progeny is compared vs no UV
and with GMR::Gal4 control progeny (S1-D). GMR::Gal4;UAS::Dad-RNAi is compared with
control UV progeny (S1-E). Scale Bar: 20 μm. Error bars indicate s.e.m.; P-values from Student’s
t-test. *p<0.05, **p<0.01, **p<0.001.

55

56
Supplementary Figure 3.2: Quantitation of Individual Drosophila eyes. Representative
images and quantitation of all experiments were collected below. Control UV WT progeny pMad+
Elav+ cells compared with non UV progeny (S2-A). UV exposed control nSmox+ Elav+ cells is
compared with non UV progeny (S2-B). Total WT DCP1+ Elav+ cells quantitation compared with
no UV (S2-C). Total Dad-nGFP+ Elav+ cells compared between UV and no UV WT progeny (S2-
C’). Individual quantitation of total GMR::G4;Hml::DppRNAi DCP1+ Elav+ cells and Dad-nGFP+
Elav+ cells compared with WT UV (S2-D). Error bars indicate s.e.m.; P-values from Student’s t-
test. *p<0.05, **p<0.01, **p<0.001.

57

Supplementary Figure 3.3: Quantitation of Individual Drosophila eyes continued.
Representative images and quantitation of all experiments were collected below. UV exposed
GMR::G4;SaxRNAi individual progeny DCP1+ Elav+ cells and Dad-nGFP+ Elav+ cells are
compared to UV WT (S3-A). UV exposed GMR::G4;SmoxRNAi individual progeny DCP1+ Elav+
cells and Dad-nGFP+ Elav+ cells are compared to UV WT (S3-B). UV exposed GMR::G4;TkvRNAi
individual progeny DCP1+ Elav+ and Dad-nGFP+ Elav+ cells are compared to UV WT (S3-C). UV
exposed GMR::G4;MadRNAi individual progeny DCP1+ Elav+ and Dad-nGFP+ Elav+ cells are
compared to UV WT (S3-D). Scale Bar: 20 μm. Error bars indicate s.e.m.; P-values from Student’s
t-test. *p<0.05, **p<0.01, **p<0.001.

58

Supplementary Figure 3.4: Comparing UV exposed Sep::G4;Smox and MadRNAi
individual progeny DCP1+ Elav+ cells to UV Sep WT. Error bars indicate s.e.m.; P-values from
Student’s t-test. *p<0.05, **p<0.01, **p<0.001.

59

Supplementary Figure 3.5: Representative images of pSmad 1/5/9 (BMP) and pSmad 2/3
(TGFB) animals pSmad 1/5/9 (BMP) and pSmad 2/3 (TGFB) control images of C57 animals
(S5-A). TUNEL stain in red of BALB/c animals post light exposure, with DAPI in blue (S5-B). PBS
WT Control unexposed retina with IBA1 in white and CD68 in green (S5-C). pSmad 1/5/9
representative images of BALB/c animals post light exposure at 0, 24 and 36 hours (S5-E).
Representative images of DT injected CD11b::DTR mice with pSmad 1/5/9 (BMP) in red, and
CRALBP in white (S5-E). Scale Bar: 30 μm.

60

References

Achberger, K., Probst, C., Haderspeck, J., Bolz, S., Rogal, J., Chuchuy, J., Nikolova, M.,
Cora, V., Antkowiak, L., Haq, W., Shen, N., Schenke-Layland, K., Ueffing, M., Liebau, S., &
Loskill, P. (2019). Merging organoid and organ-on-a-chip technology to generate complex
multi-layer tissue models in a human retina-on-a-chip platform. eLife, 8.
https://doi.org/10.7554/elife.46188
Agaisse, Hervé, et al. “Signaling Role of Hemocytes in Drosophila JAK/STAT-Dependent
Response to Septic Injury.” Developmental Cell, vol. 5, no. 3, 2003, pp. 441–50. Crossref,
doi:10.1016/s1534-5807(03)00244-2.
Akhtar ‐Schäfer, Isha, et al. “Modulation of Three Key Innate Immune Pathways for the Most
Common Retinal Degenerative Diseases.” EMBO Molecular Medicine, vol. 10, no. 10, 2018.
Crossref, doi:10.15252/emmm.201708259.
Akhtar ‐Schäfer, I., Wang, L., Krohne, T. U., Xu, H., & Langmann, T. (2018). Modulation of
three key innate immune pathways for the most common retinal degenerative diseases.
EMBO Molecular Medicine, 10(10). https://doi.org/10.15252/emmm.201708259
Appelbaum, Tatyana, et al. “Strong Upregulation of Inflammatory Genes Accompanies
Photoreceptor Demise in Canine Models of Retinal Degeneration.” PLOS ONE, edited by
Thomas Langmann, vol. 12, no. 5, 2017, p. e0177224. Crossref,
doi:10.1371/journal.pone.0177224.

61
Arias, Carolina, et al. “Dpp-Expressing and Non-Expressing Cells: Two Different
Populations of Growing Cells in Drosophila.” PLOS ONE, edited by Eduardo Moreno, vol.
10, no. 3, 2015, p. e0121457. Crossref, doi:10.1371/journal.pone.0121457.
Ashery-Padan, R., & Gruss, P. (2001). Pax6 lights-up the way for eye development. Current
Opinion in Cell Biology, 13(6), 706-714. https://doi.org/10.1016/s0955-0674(00)00274-x
Ayyaz, Arshad, et al. “Haemocytes Control Stem Cell Activity in the Drosophila Intestine.”
Nature Cell Biology, vol. 17, no. 6, 2015, pp. 736–48. Crossref, doi:10.1038/ncb3174.
Baden, T., Euler, T., & Berens, P. (2019). Understanding the retinal basis of vision across
species. Nature Reviews Neuroscience, 21(1), 5-20. https://doi.org/10.1038/s41583-019-
0242-1
Balenci, L., Wonders, C., Coles, B. L., Clarke, L., & Van der Kooy, D. (2013). Bone
morphogenetic proteins and secreted frizzled related protein 2 maintain the quiescence of
adult mammalian retinal stem cells. STEM CELLS, 31(10), 2218-2230.
https://doi.org/10.1002/stem.1470
Barbot, A. (2017). From retina to cortex. Journal of Vision, 17(15), 14.
https://doi.org/10.1167/17.15.14
Behar-Cohen, F., Baillet, De Ayguavives, Krutmann, J., Pena Garcia, Reme, C., Wolffsohn,
& Ortega García, P. (2013). Ultraviolet damage to the eye revisited: Eye-sun protection
factor (E-spf®), a new ultraviolet protection label for eyewear. Clinical Ophthalmology, 87.
https://doi.org/10.2147/opth.s46189

62
Bender, Herdis, et al. “TIEG1 Facilitates Transforming Growth Factor-?-Mediated Apoptosis
in the Oligodendroglial Cell Line OLI-Neu.” Journal of Neuroscience Research, vol. 75, no.
3, 2004, pp. 344–52. Crossref, doi:10.1002/jnr.10856.
Betts-Obregon, B. S., et al. “TGFβ Induces BIGH3 Expression and Human Retinal Pericyte
Apoptosis: A Novel Pathway of Diabetic Retinopathy.” Eye, vol. 30, no. 12, 2016, pp. 1639–
47. Crossref, doi:10.1038/eye.2016.179.
Bhatia, B., Jayaram, H., Singhal, S., Jones, M. F., & Limb, G. A. (2011). Differences between
the neurogenic and proliferative abilities of Müller glia with stem cell characteristics and the
ciliary epithelium from the adult human eye. Experimental Eye Research, 93(6), 852-861.
https://doi.org/10.1016/j.exer.2011.09.015
Blaivas, M. (2003). The injured eye. Ultrasound in Medicine & Biology, 29(5), S6.
https://doi.org/10.1016/s0301-5629(03)00091-7
Blake, J. A., Baldarelli, R., Kadin, J. A., Richardson, J. E., & Bult, C. J. (2021). Mouse
Genome Database (MGD): Knowledgebase for mouse–human comparative biology.
Nucleic Acids Research, 49(1), D981–D987. https://doi.org/10.1093/nar/gkaa1083
Bloom, S. M., & Singal, I. P. (2011). The outer bruch membrane layer. Retina, 31(2), 316-
323. https://doi.org/10.1097/iae.0b013e3181ed8c9a
Bousquet, E., Zhao, M., Thillaye-Goldenberg, B., Lorena, V., Castaneda, B., Naud, M. C.,
Bergin, C., Besson-Lescure, B., Behar-Cohen, F., & De Kozak, Y. (2015). Choroidal mast
cells in retinal pathology. The American Journal of Pathology, 185(8), 2083-2095.
https://doi.org/10.1016/j.ajpath.2015.04.002

63
Bryda, E. C. (2013). The Mighty Mouse: The Impact of Rodents on Advances in Biomedical
Research. The Journal of the Missouri State Medical Association, 110(3), 207-211.
https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3987984/
Busbee, B. G., Brown, G. C., & Brown, M. M. (2003). Cost-effectiveness of ocular
interventions. Current Opinion in Ophthalmology, 14(3), 132-138.
https://doi.org/10.1097/00055735-200306000-00004
Catalani, E., Silvestri, F., Bongiorni, S., Taddei, A. R., Fanelli, G., Rinalducci, S., De Palma,
C., Perrotta, C., Prantera, G., & Cervia, D. (2021). Retinal damage in a new model of
hyperglycemia induced by high-sucrose diets. Pharmacological Research, 166, 105488.
https://doi.org/10.1016/j.phrs.2021.105488
Chakrabarti, Sveta, et al. “Remote Control of Intestinal Stem Cell Activity by Haemocytes in
Drosophila.” PLOS Genetics, edited by Utpal Banerjee, vol. 12, no. 5, 2016, p. e1006089.
Crossref, doi:10.1371/journal.pgen.1006089.
Charlton-Perkins, M. A., Sendler, E. D., Buschbeck, E. K., & Cook, T. A. (2017).
Multifunctional glial support by semper cells in the drosophila retina. PLOS Genetics, 13(5),
e1006782. https://doi.org/10.1371/journal.pgen.1006782
Chaturvedi, R., Reddig, K., & Li, H. (2014). Long-distance mechanism of neurotransmitter
recycling mediated by glial network facilitates visual function in drosophila. Proceedings of
the National Academy of Sciences, 111(7), 2812-2817.
https://doi.org/10.1073/pnas.1323714111

64
Chen, L., Liu, M., Luan, Y., Liu, Y., Zhang, Z., Ma, B., Liu, X., & Liu, Y. (2018). BMP ‐6
protects retinal pigment epithelial cells from oxidative stress ‐induced injury by inhibiting the
MAPK signaling pathways. International Journal of Molecular Medicine.
https://doi.org/10.3892/ijmm.2018.3675
Chen, S. P., Bhattacharya, J., & Pershing, S. (2017). Association of vision loss with
cognition in older adults. JAMA Ophthalmology, 135(9), 963.
https://doi.org/10.1001/jamaophthalmol.2017.2838
Cheng, S., Cipi, J., Ma, S., Hafler, B. P., Kanadia, R. N., Brush, R. S., Agbaga, M., & Punzo,
C. (2020). Altered photoreceptor metabolism in mouse causes late stage age-related
macular degeneration-like pathologies. Proceedings of the National Academy of Sciences,
117(23), 13094-13104. https://doi.org/10.1073/pnas.2000339117
Chinnery, H. R., McMenamin, P. G., & Dando, S. J. (2017). Macrophage physiology in the
eye. Pflügers Archiv - European Journal of Physiology, 469(1), 501-515.
https://doi.org/10.1007/s00424-017-1947-5
Chistiakov, Dimitry A., et al. “CD68/Macrosialin: Not Just a Histochemical Marker.”
Laboratory Investigation, vol. 97, no. 1, 2016, pp. 4–13. Crossref,
doi:10.1038/labinvest.2016.116.
Christensen, Ian, et al. “The Susceptibility of Retinal Ganglion Cells to Glutamatergic
Excitotoxicity Is Type-Specific.” Frontiers in Neuroscience, vol. 13, 2019. Crossref,
doi:10.3389/fnins.2019.00219.

65
Christensen, I., Lu, B., Yang, N., Huang, K., Wang, P., & Tian, N. (2019). The susceptibility
of retinal ganglion cells to Glutamatergic Excitotoxicity is type-specific. Frontiers in
Neuroscience, 13. https://doi.org/10.3389/fnins.2019.00219
Close, J. L. “Retinal Neurons Regulate Proliferation of Postnatal Progenitors and Müller Glia
in the Rat Retina via TGF Signaling.” Development, vol. 132, no. 13, 2005, pp. 3015–26.
Crossref, doi:10.1242/dev.01882.
Cuenca N, Fernández-Sánchez L, Campello L, Maneu V, De la Villa P, Lax P, Pinilla I (2014)
Cellular responses following retinal injuries and therapeutic approaches for
neurodegenerative diseases. Prog Retin Eye Res 43:17–75

De Mayo, B. (2014). Vision and the perception of light and color. The Everyday Physics of
Hearing and Vision. https://doi.org/10.1088/978-1-6270-5675-5ch6
Denton, Donna, et al. “Crosstalk between Dpp and Tor Signaling Coordinates Autophagy-
Dependent Midgut Degradation.” Cell Death & Disease, vol. 10, no. 2, 2019. Crossref,
doi:10.1038/s41419-019-1368-9.
Dey, Nidhi Sharma, et al. “Dpp Dependent Hematopoietic Stem Cells Give Rise to Hh
Dependent Blood Progenitors in Larval Lymph Gland of Drosophila.” ELife, vol. 5, 2016.
Crossref, doi:10.7554/elife.18295.
Dharmarajan, S., Fisk, D. L., Sorenson, C. M., Sheibani, N., & Belecky-Adams, T. L. (2017).
Microglia activation is essential for BMP7-mediated retinal reactive gliosis. Journal of
Neuroinflammation, 14(1). https://doi.org/10.1186/s12974-017-0855-0

66
Dzik, Jolanta Maria. “Evolutionary Roots of Arginase Expression and Regulation.” Frontiers
in Immunology, vol. 5, 2014. Crossref, doi:10.3389/fimmu.2014.00544.
Eleftherianos, Ioannis, et al. “TGF-β Signaling Regulates Resistance to Parasitic Nematode
Infection in Drosophila Melanogaster.” Immunobiology, vol. 221, no. 12, 2016, pp. 1362–68.
Crossref, doi:10.1016/j.imbio.2016.07.011.
Eme-Scolan, E., & Dando, S. J. (2020). Tools and approaches for studying Microglia in vivo.
Frontiers in Immunology, 11. https://doi.org/10.3389/fimmu.2020.583647
Eusebio, N., Tavares, L., & Pereira, P. S. (2018). CtBP represses DPP-dependent mad
activation during drosophila eye development. Developmental Biology, 442(1), 188-198.
https://doi.org/10.1016/j.ydbio.2018.07.018
Evangelho, K., Mogilevskaya, M., Losada-Barragan, M., & Vargas-Sanchez, J. K. (2017).
Pathophysiology of primary open-angle glaucoma from a neuroinflammatory and
neurotoxicity perspective: A review of the literature. International Ophthalmology, 39(1),
259-271. https://doi.org/10.1007/s10792-017-0795-9
Fatima, S. (2021). Bone morphogenetic protein (BMP) signaling in early embryonic
development and cancer-an overview. Biomedical Journal of Scientific & Technical
Research, 34(4). https://doi.org/10.26717/bjstr.2021.34.005592
Forrester, J. V., Kuffova, L., & Delibegovic, M. (2020). The role of inflammation in diabetic
retinopathy. Frontiers in Immunology, 11. https://doi.org/10.3389/fimmu.2020.583687

67
Franze, K., Grosche, J., Skatchkov, S. N., Schinkinger, S., Foja, C., Schild, D., Uckermann,
O., Travis, K., Reichenbach, A., & Guck, J. (2007). Müller cells are living optical fibers in the
vertebrate retina. Proceedings of the National Academy of Sciences, 104(20), 8287-8292.
https://doi.org/10.1073/pnas.0611180104
Frieler, Ryan A., and Richard M. Mortensen. “Immune Cell and Other Noncardiomyocyte
Regulation of Cardiac Hypertrophy and Remodeling.” Circulation, vol. 131, no. 11, 2015,
pp. 1019–30. Crossref, doi:10.1161/circulationaha.114.008788.
Genander, Maria, et al. “BMP Signaling and Its PSMAD1/5 Target Genes Differentially
Regulate Hair Follicle Stem Cell Lineages.” Cell Stem Cell, vol. 15, no. 5, 2014, pp. 619–
33. Crossref, doi:10.1016/j.stem.2014.09.009.
Gervais, Louis, and Allison J. Bardin. “Tissue Homeostasis and Aging: New Insight from the
Fly Intestine.” Current Opinion in Cell Biology, vol. 48, 2017, pp. 97–105. Crossref,
doi:10.1016/j.ceb.2017.06.005.
Giannelli, S. G., Demontis, G. C., Pertile, G., Rama, P., & Broccoli, V. (2011). Adult human
Müller glia cells are a highly efficient source of rod photoreceptors. STEM CELLS, 29(2),
344-356. https://doi.org/10.1002/stem.579
Glickman, R. D. (2011). Ultraviolet Phototoxicity to the retina. Eye & Contact Lens: Science
& Clinical Practice, 37(4), 196-205. https://doi.org/10.1097/icl.0b013e31821e45a9

68
Gold, Katrina S., and Katja Brückner. “Macrophages and Cellular Immunity in Drosophila
Melanogaster.” Seminars in Immunology, vol. 27, no. 6, 2015, pp. 357–68. Crossref,
doi:10.1016/j.smim.2016.03.010.
Gold, K. S., & Brückner, K. (2016). Macrophages and cellular immunity in Drosophila
melanogaster. Semin Immunol, 27(6), 357-368. https://doi.org/
10.1016/j.smim.2016.03.010
Goldman, D. (2014). Müller glial cell reprogramming and retina regeneration. Nature
Reviews Neuroscience, 15(7), 431-442. https://doi.org/10.1038/nrn3723
Goldstein, Jeffery A., Sasha Bogdanovich, et al. “Excess SMAD Signaling Contributes to
Heart and Muscle Dysfunction in Muscular Dystrophy.” Human Molecular Genetics, vol. 23,
no. 25, 2014, pp. 6722–31. Crossref, doi:10.1093/hmg/ddu390.
Goldstein, Jeffery A., Sean M. Kelly, et al. “SMAD Signaling Drives Heart and Muscle
Dysfunction in a Drosophila Model of Muscular Dystrophy.” Human Molecular Genetics, vol.
20, no. 5, 2010, pp. 894–904. Crossref, doi:10.1093/hmg/ddq528.
Gong, T., Liu, L., Jiang, W., & Zhou, R. (2019). DAMP-sensing receptors in sterile
inflammation and inflammatory diseases. Nature Reviews Immunology, 20(2), 95-112.
https://doi.org/10.1038/s41577-019-0215-7
Grimm, Christian, and Charlotte E. Remé. “Light Damage as a Model of Retinal
Degeneration.” Methods in Molecular Biology, 2012, pp. 87–97. Crossref, doi:10.1007/978-
1-62703-080-9_6.

69
Groeger, G., Doonan, F., Cotter, T. G., & Donovan, M. (2012). Reactive oxygen species
regulate Prosurvival ERK1/2 signaling and bFGF expression in gliosis within the retina.
Investigative Opthalmology & Visual Science, 53(10), 6645. https://doi.org/10.1167/iovs.12-
10525
Guo, S., Jiang, X., Mao, B., & Li, Q. (2019). The design, analysis and application of mouse
clinical trials in oncology drug development. BMC Cancer, 19(1).
https://doi.org/10.1186/s12885-019-5907-7
Hamaratoglu, Fisun, et al. “Dpp/BMP Signaling in Flies: From Molecules to Biology.”
Seminars in Cell & Developmental Biology, vol. 32, 2014, pp. 128–36. Crossref,
doi:10.1016/j.semcdb.2014.04.036.
Haynes, T., Gutierrez, C., Aycinena, J., Tsonis, P. A., & Del Rio-Tsonis, K. (2007). BMP
signaling mediates stem/progenitor cell-induced retina regeneration. Proceedings of the
National Academy of Sciences, 104(51), 20380-20385.
https://doi.org/10.1073/pnas.0708202104
Heavner, W., & Pevny, L. (2012). Eye development and Retinogenesis. Cold Spring Harbor
Perspectives in Biology, 4(12), a008391-a008391.
https://doi.org/10.1101/cshperspect.a008391
Helmbacher, F. (2020). Fat1 regulates astrocyte maturation and angiogenesis in the retina.
https://doi.org/10.1101/2020.04.23.057141

70
Hickman, D., Johnson, J., Vemulapalli, T., Crisler, J., & Shepherd, R. (2017). Commonly
used animal models. Principles of Animal Research, 117-175. https://doi.org/10.1016/b978-
0-12-802151-4.00007-4

Hu R, Kagele DA, Huffaker TB, Runtsch MC, Alexander M, Liu J, Bake E, Su W, Williams
MA, Rao DS et al (2014) miR-155 promotes T follicular helper cell accumulation during
chronic, low-grade infammation. Immunity 41:605–619
Hudry, E., & Vandenberghe, L. H. (2019). Therapeutic AAV gene transfer to the nervous
system: A clinical reality. Neuron, 102(1), 263. https://doi.org/10.1016/j.neuron.2019.03.020
Inoue, Hirofumi, et al. “Interplay of Signal Mediators of Decapentaplegic (Dpp): Molecular
Characterization of Mothers against Dpp, Medea, and Daughters against Dpp.” Molecular
Biology of the Cell, edited by Carl-Henrik Heldin, vol. 9, no. 8, 1998, pp. 2145–56. Crossref,
doi:10.1091/mbc.9.8.2145.
Israel, A. (2009). The IKK complex, a central regulator of NF- B activation. Cold Spring
Harbor Perspectives in Biology, 2(3), a000158-a000158.
https://doi.org/10.1101/cshperspect.a000158
Jassim, O. W. “Dmp53 Protects the Drosophila Retina during a Developmentally Regulated
DNA Damage Response.” The EMBO Journal, vol. 22, no. 20, 2003, pp. 5622–32. Crossref,
doi:10.1093/emboj/cdg543.
Jassim, O. W. (2003). Dmp53 protects the drosophila retina during a developmentally
regulated DNA damage response. The EMBO Journal, 22(20), 5622-5632.
https://doi.org/10.1093/emboj/cdg543

71
Javier, Anna L., et al. “Bmp Indicator Mice Reveal Dynamic Regulation of Transcriptional
Response.” PLoS ONE, edited by Michael Schubert, vol. 7, no. 9, 2012, p. e42566.
Crossref, doi:10.1371/journal.pone.0042566.
Jeibmann, A., & Paulus, W. (2009). Drosophila melanogaster as a model organism of brain
diseases. International Journal of Molecular Sciences, 10(2), 407-440.
https://doi.org/10.3390/ijms10020407
Ji, Tuo, et al. “199 Regulation of BMP-4 Expression by Inflammatory Cytokines.”
Gastroenterology, vol. 146, no. 5, 2014, p. S-52. Crossref, doi:10.1016/s0016-
5085(14)60182-7.
Joly, S., Pernet, V., Samardzija, M., & Grimm, C. (2011). Pax6-positive Müller glia cells
express cell cycle markers but do not proliferate after photoreceptor injury in the mouse
retina. Glia, 59(7), 1033-1046. https://doi.org/10.1002/glia.21174
Jorstad, N. L., Wilken, M. S., Grimes, W. N., Wohl, S. G., VandenBosch, L. S., Yoshimatsu,
T., Wong, R. O., Rieke, F., & Reh, T. A. (2017). Stimulation of functional neuronal
regeneration from Müller glia in adult mice. Nature, 548(7665), 103-107.
https://doi.org/10.1038/nature23283
Kamiya, Yuto, et al. “Specificity of the Inhibitory Effects of Dad on TGF-β Family Type I
Receptors, Thickveins, Saxophone, and Baboon InDrosophila.” FEBS Letters, vol. 582, no.
17, 2008, pp. 2496–500. Crossref, doi:10.1016/j.febslet.2008.05.052.

72
Kamiya, Y., Miyazono, K., & Miyazawa, K. (2008). Specificity of the inhibitory effects of dad
on TGF-β family type I receptors, Thickveins, saxophone, and baboon inDrosophila. FEBS
Letters, 582(17), 2496-2500. https://doi.org/10.1016/j.febslet.2008.05.052
Kang, Igojo, et al. “Identification of Target Genes Regulated by the Drosophila Histone
Methyltransferase Eggless Reveals a Role of Decapentaplegic in Apoptotic Signaling.”
Scientific Reports, vol. 8, no. 1, 2018. Crossref, doi:10.1038/s41598-018-25483-9.
Kang, M., & Ryoo, H. D. (2009). Suppression of retinal degeneration in drosophila by
stimulation of ER-associated degradation. Proceedings of the National Academy of
Sciences, 106(40), 17043-17048. https://doi.org/10.1073/pnas.0905566106
Karl, M. O., et al. “Stimulation of Neural Regeneration in the Mouse Retina.” Proceedings of
the National Academy of Sciences, vol. 105, no. 49, 2008, pp. 19508–13. Crossref,
doi:10.1073/pnas.0807453105.

Karlstetter M, Scholz R, Rutar M, Wong WT, Provis JM, Langmann T (2015) Retinal
microglia: just bystander or target for therapy? Prog Retin Eye Res 45:30–57
Karperien, Audrey, et al. “Quantitating the Subtleties of Microglial Morphology with Fractal
Analysis.” Frontiers in Cellular Neuroscience, vol. 7, 2013. Crossref,
doi:10.3389/fncel.2013.00003.
Kashima, Risa, and Akiko Hata. “The Role of TGF-β Superfamily Signaling in Neurological
Disorders.” Acta Biochimica et Biophysica Sinica, vol. 50, no. 1, 2017, pp. 106–20. Crossref,
doi:10.1093/abbs/gmx124.

73
Katz, Ben. “Drosophila Photoreceptors and Signaling Mechanisms.” Frontiers, 2009,
www.frontiersin.org/articles/10.3389/neuro.03.002.2009/full.
Kelsey, E. M., et al. “Schnurri Regulates Hemocyte Function to Promote Tissue Recovery
after DNA Damage.” Journal of Cell Science, vol. 125, no. 6, 2012, pp. 1393–400. Crossref,
doi:10.1242/jcs.095323.
Kolster, H. (2012). Comparative mapping of visual areas in the human and macaque
occipital cortex. Journal of Vision, 12(9), 1378-1378. https://doi.org/10.1167/12.9.1378
Kondoh, H. (2002). Development of the eye. Mouse Development, 519-538.
https://doi.org/10.1016/b978-012597951-1/50025-1

Kramer J, Chirco KR, Lamba DA. Immunological Considerations for Retinal Stem Cell
Therapy. Adv Exp Med Biol. 2019;1186:99-119. doi: 10.1007/978-3-030-28471-8_4. PMID:
31654387.
Krebs, M. C., Collin, G. B., Hicks, W. L., Yu, M., & Shi, L. Y. (2017). Mouse models of human
ocular disease for translational research. Plos One, 12(8), e0183837.
https://doi.org/10.1371/journal.pone.0183837
Kuczma, M., & Kraj, P. (2015). Bone morphogenetic protein signaling regulates
development and activation of CD4+ T cells. Bone Morphogenic Protein, 171-193.
https://doi.org/10.1016/bs.vh.2015.05.001
Kuehn, Sandra, et al. “Concentration-Dependent Inner Retina Layer Damage and Optic
Nerve Degeneration in a NMDA Model.” Journal of Molecular Neuroscience, vol. 63, no. 3–
4, 2017, pp. 283–99. Crossref, doi:10.1007/s12031-017-0978-x.

74
Kumar, J. P. (2013). Building an Ommatidium One Cell at a Time. Developmental Dynamics,
241(1), 136-149. https://doi.org/10.1002/dvdy.23707
Lamba, Deepak, et al. “Neural Regeneration and Cell Replacement: A View from the Eye.”
Cell Stem Cell, vol. 2, no. 6, 2008, pp. 538–49. Crossref, doi:10.1016/j.stem.2008.05.002.
Lambuk, Lidawani, et al. “Antiapoptotic Effect of Taurine against NMDA-Induced Retinal
Excitotoxicity in Rats.” NeuroToxicology, vol. 70, 2019, pp. 62–71. Crossref,
doi:10.1016/j.neuro.2018.10.009.
Land, M. F., & Nilsson, D. (2012). Animal eyes.
https://doi.org/10.1093/acprof:oso/9780199581139.001.0001
Lebestky, T. (2000). Specification of drosophila hematopoietic lineage by conserved
transcription factors. Science, 288(5463), 146-149.
https://doi.org/10.1126/science.288.5463.146
Lee, J. E., Liang, K. J., Fariss, R. N., & Wong, W. T. (2008). Ex vivo dynamic imaging of
retinal Microglia using time-lapse confocal microscopy. Investigative Ophthalmology &
Visual Science, 49(9), 4169-4176. https://doi.org/10.1167/iovs.08-2076
Lenkowski, Jenny R., et al. “Retinal Regeneration in Adult Zebrafish Requires Regulation of
TGFβ Signaling.” Glia, vol. 61, no. 10, 2013, pp. 1687–97. Crossref, doi:10.1002/glia.22549.

75
Lenkowski, J. R., Qin, Z., Sifuentes, C. J., Thummel, R., Soto, C. M., Moens, C. B., &
Raymond, P. A. (2013). Retinal regeneration in adult zebrafish requires regulation of TGFβ
signaling. Glia, 61(10), 1687-1697. https://doi.org/10.1002/glia.22549
Li, Chun-Yan, et al. “TGFβ Receptor Saxophone Non-Autonomously Regulates Germline
Proliferation in a Smox/DSmad2-Dependent Manner in Drosophila Testis.” Developmental
Biology, vol. 309, no. 1, 2007, pp. 70–77. Crossref, doi:10.1016/j.ydbio.2007.06.019.
Li, Fang-Fang, et al. “Identification and Characterization of an R-Smad Homologue (Hco-
DAF-8) from Haemonchus Contortus.” Parasites & Vectors, vol. 13, no. 1, 2020. Crossref,
doi:10.1186/s13071-020-04034-0.
Li, C., Guo, Z., & Wang, Z. (2007). TGFβ receptor saxophone non-autonomously regulates
germline proliferation in a Smox/dsmad2-dependent Manner in drosophila testis.
Developmental Biology, 309(1), 70-77. https://doi.org/10.1016/j.ydbio.2007.06.019
Liou, R. H., Edwards, T. L., Martin, K. R., & Wong, R. C. (2020). Neuronal reprogramming
for tissue repair and Neuroregeneration. International Journal of Molecular Sciences,
21(12), 4273. https://doi.org/10.3390/ijms21124273
Liu, R., & Desai, L. P. (2015). Reciprocal regulation of TGF-β and reactive oxygen species:
A perverse cycle for fibrosis. Redox Biology, 6, 565-577.
https://doi.org/10.1016/j.redox.2015.09.009
Liu, R., & Gaston Pravia, K. (2010). Oxidative stress and glutathione in TGF-β-mediated
fibrogenesis. Free Radical Biology and Medicine, 48(1), 1-15.
https://doi.org/10.1016/j.freeradbiomed.2009.09.026

76
London A, Benhar I, Schwartz M. The retina as a window to the brain-from eye research to
CNS disorders. Nat Rev Neurol. 2013 Jan;9(1):44-53. doi: 10.1038/nrneurol.2012.227.
Epub 2012 Nov 20. PMID: 23165340.

Luo, Xi, et al. “Foxo and Fos Regulate the Decision between Cell Death and Survival in
Response to UV Irradiation.” The EMBO Journal, vol. 26, no. 2, 2006, pp. 380–90. Crossref,
doi:10.1038/sj.emboj.7601484.
Ma, Wenxin, et al. “Absence of TGFβ Signaling in Retinal Microglia Induces Retinal
Degeneration and Exacerbates Choroidal Neovascularization.” ELife, vol. 8, 2019. Crossref,
doi:10.7554/elife.42049.
Maekawa, Shigeto, et al. “The Neuroprotective Effect of Hesperidin in NMDA-Induced
Retinal Injury Acts by Suppressing Oxidative Stress and Excessive Calpain Activation.”
Scientific Reports, vol. 7, no. 1, 2017. Crossref, doi:10.1038/s41598-017-06969-4.
Malzer, Elke, et al. “The Integrated Stress Response Regulates BMP Signalling through
Effects on Translation.” BMC Biology, vol. 16, no. 1, 2018. Crossref, doi:10.1186/s12915-
018-0503-x.
Manjón, Cristina, et al. “Sharp Boundaries of Dpp Signalling Trigger Local Cell Death
Required for Drosophila Leg Morphogenesis.” Nature Cell Biology, vol. 9, no. 1, 2006, pp.
57–63. Crossref, doi:10.1038/ncb1518.

Martinez, F. O., & Gordon, S. (2014). The M1 and M2 paradigm of macrophage activation:
time for reassessment. F1000prime reports, 6, 13. https://doi.org/10.12703/P6-13

77
McPherson, Scott W., et al. “The Retinal Environment Induces Microglia-like Properties in
Recruited Myeloid Cells.” Journal of Neuroinflammation, vol. 16, no. 1, 2019. Crossref,
doi:10.1186/s12974-019-1546-9.
Minhas, Gillipsie, et al. “Cellular Stress Response and Immune Signaling in Retinal
Ischemia–Reperfusion Injury.” Frontiers in Immunology, vol. 7, 2016. Crossref,
doi:10.3389/fimmu.2016.00444.
Mirzoyan, Z., Sollazzo, M., Allocca, M., Valenza, A. M., Grifoni, D., & Bellosta, P. (2019).
Drosophila melanogaster: A model organism to study cancer. Frontiers in Genetics, 10.
https://doi.org/10.3389/fgene.2019.00051
Moon, L. D., & Fawcett, J. W. (2001). Reduction in CNS scar formation without concomitant
increase in axon regeneration following treatment of adult rat brain with a combination of
antibodies to TGFβ1and β2. European Journal of Neuroscience, 14(10), 1667-1677.
https://doi.org/10.1046/j.0953-816x.2001.01795.x
Moraczewski, A. L. (2007). History and examination of the injured eye. Ocular Trauma, 19-
38. https://doi.org/10.1016/b978-1-4160-2559-7.50009-x
Moses, K., and G. M. Rubin. “Glass Encodes a Site-Specific DNA-Binding Protein That Is
Regulated in Response to Positional Signals in the Developing Drosophila Eye.” Genes &
Development, vol. 5, no. 4, 1991, pp. 583–93. Crossref, doi:10.1101/gad.5.4.583.
Nagashima, M., & Hitchcock, P. F. (2021). Inflammation regulates the multi-step process of
retinal regeneration in Zebrafish. Cells, 10(4), 783. https://doi.org/10.3390/cells10040783

78
Neal, Scott J., et al. “Drosophila ML-DmD17-C3 Cells Respond Robustly to Dpp and Exhibit
Complex Transcriptional Feedback on BMP Signaling Components.” BMC Developmental
Biology, vol. 19, no. 1, 2019. Crossref, doi:10.1186/s12861-019-0181-0.
Neuzillet, C., De Gramont, A., Tijeras-Raballand, A., De Mestier, L., Cros, J., Faivre, S., &
Raymond, E. (2013). Perspectives of TGF-β inhibition in pancreatic and hepatocellular
carcinomas. Oncotarget, 5(1), 78-94. https://doi.org/10.18632/oncotarget.1569
Neves, Joana, et al. “Immune Modulation by MANF Promotes Tissue Repair and
Regenerative Success in the Retina.” Science, vol. 353, no. 6294, 2016, p. aaf3646.
Crossref, doi:10.1126/science.aaf3646.
Ninov, Nikolay, et al. “Dpp Signaling Directs Cell Motility and Invasiveness during Epithelial
Morphogenesis.” Current Biology, vol. 20, no. 6, 2010, pp. 513–20. Crossref,
doi:10.1016/j.cub.2010.01.063.
Niwa, Masayuki, et al. “Retinal Cell Degeneration in Animal Models.” International Journal
of Molecular Sciences, vol. 17, no. 1, 2016, p. 110. Crossref, doi:10.3390/ijms17010110.
Okunuki, Yoko, Ryo Mukai, Elizabeth A. Pearsall, et al. “Microglia Inhibit Photoreceptor Cell
Death and Regulate Immune Cell Infiltration in Response to Retinal Detachment.”
Proceedings of the National Academy of Sciences, vol. 115, no. 27, 2018, pp. E6264–73.
Crossref, doi:10.1073/pnas.1719601115.
Okunuki, Yoko, Ryo Mukai, Takeshi Nakao, et al. “Retinal Microglia Initiate
Neuroinflammation in Ocular Autoimmunity.” Proceedings of the National Academy of
Sciences, vol. 116, no. 20, 2019, pp. 9989–98. Crossref, doi:10.1073/pnas.1820387116.

79
Ortín-Martínez, A., Nadal-Nicolás, F. M., Jiménez-López, M., Alburquerque-Béjar, J. J.,
Nieto-López, L., García-Ayuso, D., Villegas-Pérez, M. P., Vidal-Sanz, M., & Agudo-Barriuso,
M. (2014). Number and distribution of mouse retinal cone photoreceptors: Differences
between an albino (Swiss) and a pigmented (C57/BL6) strain. PLoS ONE, 9(7), e102392.
https://doi.org/10.1371/journal.pone.0102392
Ozakman, Yaprak, and Ioannis Eleftherianos. “TGF-β Signaling Interferes With the
Drosophila Innate Immune and Metabolic Response to Parasitic Nematode Infection.”
Frontiers in Physiology, vol. 10, 2019. Crossref, doi:10.3389/fphys.2019.00716.
Plemel, J. R., Stratton, J. A., Michaels, N. J., Rawji, K. S., Zhang, E., Sinha, S., Baaklini, C.
S., Dong, Y., Ho, M., Thorburn, K., Friedman, T. N., Jawad, S., Silva, C., Caprariello, A. V.,
Hoghooghi, V., Yue, J., Jaffer, A., Lee, K., Kerr, B. J., … Yong, V. W. (2020). Microglia
response following acute demyelination is heterogeneous and limits infiltrating macrophage
dispersion. Science Advances, 6(3), eaay6324. https://doi.org/10.1126/sciadv.aay6324
Ranaei Pirmardan, Ehsan, et al. “In Vivo Evaluation of PAX6 Overexpression and NMDA
Cytotoxicity to Stimulate Proliferation in the Mouse Retina.” Scientific Reports, vol. 8, no. 1,
2018. Crossref, doi:10.1038/s41598-018-35884-5.
Rathnasamy, G., Foulds, W. S., Ling, E., & Kaur, C. (2019). Retinal microglia – A key player
in healthy and diseased retina. Progress in Neurobiology, 173, 18-40.
https://doi.org/10.1016/j.pneurobio.2018.05.006

80
Ray, C. T., Horvat, M., Croce, R., Christopher Mason, R., & Wolf, S. L. (2008). The impact
of vision loss on postural stability and balance strategies in individuals with profound vision
loss. Gait & Posture, 28(1), 58-61. https://doi.org/10.1016/j.gaitpost.2007.09.010
Reichenbach, A., & Bringmann, A. (2013). New functions of Müller cells. Glia, 61(5), 651-
678. https://doi.org/10.1002/glia.22477
Rock, Kenneth L., and Hajime Kono. “The Inflammatory Response to Cell Death.” Annual
Review of Pathology: Mechanisms of Disease, vol. 3, no. 1, 2008, pp. 99–126. Crossref,
doi:10.1146/annurev.pathmechdis.3.121806.151456.
Rübsam, A., Parikh, S., & Fort, P. E. (2018). Role of Inflammation in Diabetic Retinopathy.
International journal of molecular sciences, 19(4), 942.
https://doi.org/10.3390/ijms19040942

Safi, S., & Ahmadieh, H. (2017). Burden of vision loss in the Eastern Mediterranean Region,
1990–2015: Findings from the global burden of disease 2015 study. International Journal of
Public Health, 63(S1), 199-210. https://doi.org/10.1007/s00038-017-1000-7
Saglam, A., Kim, S., Ahn, K., Oh, I., & Lee, K. (2014). BMP2 shows neurotrophic effects
including neuroprotection against neurodegeneration. NeuroReport, 25(8), 549-555.
https://doi.org/10.1097/wnr.0000000000000130
Saika, Shizuya. “TGFβ Pathobiology in the Eye.” Laboratory Investigation, vol. 86, no. 2,
2005, pp. 106–15. Crossref, doi:10.1038/labinvest.3700375.

81
Sakami, S., Imanishi, Y., & Palczewski, K. (2018). Müller glia phagocytose dead
photoreceptor cells in a mouse model of retinal degenerative disease. The FASEB Journal,
33(3), 3680-3692. https://doi.org/10.1096/fj.201801662r
Sanes, Joshua R., and S. Lawrence Zipursky. “Design Principles of Insect and Vertebrate
Visual Systems.” Neuron, vol. 66, no. 1, 2010, pp. 15–36. Crossref,
doi:10.1016/j.neuron.2010.01.018.
Sanjabi, Shomyseh, et al. “Regulation of the Immune Response by TGF-β: From
Conception to Autoimmunity and Infection.” Cold Spring Harbor Perspectives in Biology,
vol. 9, no. 6, 2017, p. a022236. Crossref, doi:10.1101/cshperspect.a022236.
Sells, S., & Fixott, R. S. (1957). Evaluation of research on effects of visual training on visual
functions*. American Journal of Ophthalmology, 44(2), 230-236.
https://doi.org/10.1016/0002-9394(57)90012-0
Sheth, B. R., & Young, R. (2016). Two visual pathways in primates based on sampling of
space: Exploitation and exploration of visual information. Frontiers in Integrative
Neuroscience, 10. https://doi.org/10.3389/fnint.2016.00037

Shin, Mingyu, et al. “Subpopulation of Macrophage-Like Plasmatocytes Attenuates
Systemic Growth via JAK/STAT in the Drosophila Fat Body.” Frontiers in Immunology, vol.
11, Jan. 2020, p. 63. DOI.org (Crossref), doi:10.3389/fimmu.2020.00063.
Silies, M., Yuva-Aydemir, Y., Rut Franzdottir, S., & Klämbt, C. (2010). The eye imaginal disc
as a model to study the coordination of neuronal and glial development. Fly, 4(1), 71-79.
https://doi.org/10.4161/fly.4.1.11312

82
Song Z., Coca D., Billings S., Postma M., Hardie R.C., Juusola M. (2009) Biophysical
Modeling of a Drosophila Photoreceptor. In: Leung C.S., Lee M., Chan J.H. (eds) Neural
Information Processing. ICONIP 2009. Lecture Notes in Computer Science, vol 5863.
Springer, Berlin, Heidelberg. https://doi.org/10.1007/978-3-642-10677-4_7
Spokony, R. “Spokony Insertions.” Personal Communication to FlyBase, 2013,
scholar.google.com/scholar_lookup?title=Spokony+insertions&author=R+Spokony&author
=K+White&publication_year=2013&.
Sprecher, S. G., & Desplan, C. (2008). Development of the drosophila melanogaster eye:
From precursor specification to terminal differentiation. Animal Models in Eye Research, 27-
47. https://doi.org/10.1016/b978-0-12-374169-1.00004-7
Stenkamp, D. L. (2015). Development of the vertebrate eye and retina. Progress in
Molecular Biology and Translational Science, 397-414.
https://doi.org/10.1016/bs.pmbts.2015.06.006
Stieglitz, T. (2020). Of man and mice: Translational research in Neurotechnology. Neuron,
105(1), 12-15. https://doi.org/10.1016/j.neuron.2019.11.030
Stofanko, M., Kwon, S. Y., & Badenhorst, P. (2010). Lineage tracing of Lamellocytes
demonstrates drosophila macrophage plasticity. PLoS ONE, 5(11), e14051.
https://doi.org/10.1371/journal.pone.0014051

83
Sugimoto, H., Grahovac, G., Zeisberg, M., & Kalluri, R. (2007). Renal fibrosis and
Glomerulosclerosis in a new mouse model of diabetic nephropathy and its regression by
bone Morphogenic protein-7 and advanced Glycation end product inhibitors. Diabetes,
56(7), 1825-1833. https://doi.org/10.2337/db06-1226
Suh, Greg S. B., et al. “Drosophila JAB1/CSN5 Acts in Photoreceptor Cells to Induce Glial
Cells.” Neuron, vol. 33, no. 1, 2002, pp. 35–46. Crossref, doi:10.1016/s0896-
6273(01)00576-1.

Suh HS, Zhao ML, Derico L, Choi N, Lee SC. Insulin-like growth factor 1 and 2 (IGF1,
IGF2) expression in human microglia: differential regulation by inflammatory
mediators. J Neuroinflammation. 2013 Mar 12;10:37. doi: 10.1186/1742-2094-10-
37. PMID: 23497056; PMCID: PMC3607995.
Talati, Megha, et al. “BMP Pathway Regulation of and by Macrophages.” PLoS ONE, edited
by Ari Waisman, vol. 9, no. 4, 2014, p. e94119. Crossref,
doi:10.1371/journal.pone.0094119.
Tappeiner, Christoph, et al. “Inhibition of the TGFβ Pathway Enhances Retinal
Regeneration in Adult Zebrafish.” PLOS ONE, edited by Ryan Thummel, vol. 11, no. 11,
2016, p. e0167073. Crossref, doi:10.1371/journal.pone.0167073.
Teraoka, M. E., Paschaki, M., Muta, Y., & Ladher, R. K. (2009). Rostral paraxial mesoderm
regulates refinement of the eye field through the bone morphogenetic protein (BMP)
pathway. Developmental Biology, 330(2), 389-398.
https://doi.org/10.1016/j.ydbio.2009.04.008

84
Thompson, Adam, et al. “Activation of the BMP4/Smad1 Pathway Promotes Retinal
Ganglion Cell Survival and Axon Regeneration.” Investigative Opthalmology & Visual
Science, vol. 60, no. 5, 2019, p. 1748. Crossref, doi:10.1167/iovs.18-26449.
Tian, Aiguo, et al. “Injury-Stimulated and Self-Restrained BMP Signaling Dynamically
Regulates Stem Cell Pool Size during Drosophila Midgut Regeneration.” Proceedings of the
National Academy of Sciences, vol. 114, no. 13, 2017, pp. E2699–708. Crossref,
doi:10.1073/pnas.1617790114.
Todd, Levi, et al. “BMP- and TGFβ-Signaling Regulate the Formation of Müller Glia-Derived
Progenitor Cells in the Avian Retina.” Glia, vol. 65, no. 10, 2017, pp. 1640–55. Crossref,
doi:10.1002/glia.23185.
Tsuneizumi, Kazuhide, et al. “Daughters against Dpp Modulates Dpp Organizing Activity in
Drosophila Wing Development.” Nature, vol. 389, no. 6651, 1997, pp. 627–31. Crossref,
doi:10.1038/39362.

Ueki, Y., & Reh, T. A. (2013). EGF stimulates Müller glial proliferation via a BMP-dependent
mechanism. Glia, 61(5), 778-789. https://doi.org/10.1002/glia.22472
Upadhyay, Ambuj, et al. “TGF-β Family Signaling in Drosophila.” Cold Spring Harbor
Perspectives in Biology, vol. 9, no. 9, 2017, p. a022152. Crossref,
doi:10.1101/cshperspect.a022152.

85
Urbina, P., & Singla, D. K. (2014). BMP-7 attenuates adverse cardiac remodeling mediated
through M2 macrophages in prediabetic cardiomyopathy. American Journal of Physiology-
Heart and Circulatory Physiology, 307(5), H762-H772.
https://doi.org/10.1152/ajpheart.00367.2014
Van Norman, G. A. (2019). Phase II trials in drug development and adaptive trial design.
JACC: Basic to Translational Science, 4(3), 428-437.
https://doi.org/10.1016/j.jacbts.2019.02.005

VanHook, A., & Letsou, A. (2007). Head involution inDrosophila: Genetic and
morphogenetic connections to dorsal closure. Developmental Dynamics, 237(1), 28-38.
https://doi.org/10.1002/dvdy.21405
Velarde, Sergio, et al. “Dpp and Hedgehog Promote the Glial Response to Neuronal
Damage in the Developing Drosophila Visual System.” BioRxiv, 2020. bioRxiv,
doi:10.1101/2020.02.24.960625.
Velarde, S. B., Quevedo, A., Estella, C., & Baonza, A. (2020). DPP and hedgehog promote
the glial response to neuronal damage in the developing drosophila visual system.
https://doi.org/10.1101/2020.02.24.960625
Vlisidou, Isabella, and Will Wood. “Drosophilablood Cells and Their Role in Immune
Responses.” FEBS Journal, vol. 282, no. 8, 2015, pp. 1368–82. Crossref,
doi:10.1111/febs.13235.
Volland, S., Esteve-Rudd, J., Hoo, J., Yee, C., & Williams, D. S. (2015). A comparison of
some organizational characteristics of the mouse central retina and the human macula.
PLOS ONE, 10(4), e0125631. https://doi.org/10.1371/journal.pone.0125631

86
Wang, Zixing, et al. “Two-Factor Specification of Apoptosis: TGF-β Signaling Acts
Cooperatively with Ecdysone Signaling to Induce Cell- and Stage-Specific Apoptosis of
Larval Neurons during Metamorphosis in Drosophila Melanogaster.” Apoptosis, vol. 24, no.
11–12, 2019, pp. 972–89. Crossref, doi:10.1007/s10495-019-01574-4.
Warr, C., Shaw, K., Azim, A., Piper, M., & Parsons, L. (2018). Using mouse and drosophila
models to investigate the mechanistic links between diet, obesity, type II diabetes, and
cancer. International Journal of Molecular Sciences, 19(12), 4110.
https://doi.org/10.3390/ijms19124110
Wenzel, Andreas, et al. “The Genetic ModifierRpe65Leu450: Effect on Light Damage
Susceptibility in c-Fos-Deficient Mice.” Investigative Opthalmology & Visual Science, vol.
44, no. 6, 2003, p. 2798. Crossref, doi:10.1167/iovs.02-1134.
White, D. Alan, et al. “Increased Sensitivity to Light-Induced Damage in a Mouse Model of
Autosomal Dominant Retinal Disease.” Investigative Opthalmology & Visual Science, vol.
48, no. 5, 2007, p. 1942. Crossref, doi:10.1167/iovs.06-1131.
Wordinger, R. J., & Clark, A. F. (2007). Bone morphogenetic proteins and their receptors in
the eye. Experimental Biology and Medicine, 232(8), 979-992. https://doi.org/10.3181/0510-
mr-345
Xu, D., et al. “The Effector Caspases DrICE and Dcp-1 Have Partially Overlapping Functions
in the Apoptotic Pathway in Drosophila.” Cell Death & Differentiation, vol. 13, no. 10, 2006,
pp. 1697–706. Crossref, doi:10.1038/sj.cdd.4401920.

87
Yao, L. C. “Schnurri Transcription Factors from Drosophila and Vertebrates Can Mediate
Bmp Signaling through a Phylogenetically Conserved Mechanism.” Development, vol. 133,
no. 20, 2006, pp. 4025–34. Crossref, doi:10.1242/dev.02561.
Young, Richard W. “Cell Proliferation during Postnatal Development of the Retina in the
Mouse.” Developmental Brain Research, vol. 21, no. 2, 1985, pp. 229–39. Crossref,
doi:10.1016/0165-3806(85)90211-1.
Zhang, Jianye, et al. “Molecular Pharmacodynamics of Emixustat in Protection against
Retinal Degeneration.” Journal of Clinical Investigation, vol. 125, no. 7, 2015, pp. 2781–94.
Crossref, doi:10.1172/jci80950.
Zhou, T., Huang, Z., Sun, X., Zhu, X., Zhou, L., Li, M., Cheng, B., Liu, X., & He, C. (2017).
Microglia polarization with M1/M2 phenotype changes in rd1 mouse model of retinal
degeneration. Frontiers in Neuroanatomy, 11. https://doi.org/10.3389/fnana.2017.00077

Abstract (if available)

Abstract Visual perception through the eye is crucial for most animals’ survival in the wild. This remains true among humans, who devote two-thirds of all electrical activity and fifty percent of all neural tissue in the brain to vision (Sells & Fixott, 1957; Kolster, 2012; Sheth et al., 2016). ❧ Within the eye, the retina is the main light sensing tissue. It is responsible for translating an organisms’ perception of the world into neural impulses in the brain. The retina consists of several basic structures; photoreceptor cells, support cells, pigmented epithelium, and resident immune cells, all of which work together to sustain the organ (Barbot et al., 2017; Baden et al., 2019). Any injury that damages the individual structures of the retina must be handled rapidly to maintain its function. ❧ Retinal homeostasis depends on elaborate management of cell survival and apoptosis in response to injury. The result of this outcome is crucial for the organism’s long-term survival as most retinal cells are irreplaceable. However, the signaling mechanisms that control these processes remain poorly understood. Decapentaplegic (Dpp), the first identified secreted morphogen, is required for tissue formation, growth, scope, and maturation in the Drosophila eye. Due to previous findings (Kelsey et al., 2012; Ayyaz et al., 2015; Neves et al., 2016), we hypothesized that retinal morphogenic factors involved in growth and maturation such as Dpp may also play a role in tissue repair and recovery. ❧ We first chose to resolve the question of whether Drosophila immune cells, ninety five percent of which are hemocytes, drove expression of Dpp under retinal injury. To accomplish this, we used a previously characterized fly line overexpressing a constitutively active form of the Jun N-terminal Kinase Kinase (JNKK) Hemipterous (Hep) under the control of the photoreceptor and cone cell driver Sep-Gal4 (Therrien et al., 1999; Jasper et al., 2002; Kanda et al., 2004; Luo et al., 2007; Kelsey et al., 2012). When Sep-HepACT was crossed to flies expressing Sax-RNAi or Smox-RNAi under the control of UAS, a significant increase in surviving photoreceptors compared to control flies was observed. Over-expression of the negative feedback inhibitor of Dpp signaling, Daughters against Dpp (Dad; Tsuneizumi et al., 1997; Kamiya et al., 2008), also promoted survival. Conversely, we found that knocking down Tkv and Mad resulted in a significant decrease in surviving photoreceptors. When Sep-HepACT was crossed to flies expressing Hemolectin-Dppᴿᴺᴬⁱ apoptosis was also significantly reduced. ❧ To answer the question of how eye injury drives Dpp expression, we used a Drosophila model for irradiation-induced damage, collecting glass multimer reporter driver (GMR-Gal4) (Moses and Ruben, 1991) to express RNAi targeting Dpp pathway components in all postmitotic cells of the retina. We find that the BMP orthologue Decapentaplegic (Dpp) is released by immune cells after irradiation and stimulates apoptosis shortly after damage through the type I receptor Thickveins (Tkv) and the signal transducer Mothers against Dpp (Mad). At a later stage, the alternative type I receptor Saxophone, and its target Smad on X (Smox) promote tissue repair and photoreceptor survival. ❧ In mammals, Dpp homologues include various members of the bone morphogenic protein (BMP) and transforming growth factor beta (TGFβ) family. Their downstream targets include Smad 1/5/8 (activated by BMP) and Smad 2/3 (activated by TGFβ). BMP 2/4 have been shown to act in an anti-inflammatory manner, biasing macrophages to their anti-inflammatory M2 subtype (Talati et al., 2014; Dzik, et al., 2014; Ji et al., 2014; Shinohara et. al., 2010). These proteins also stimulate Müller glia proliferation, promote survival of retinal ganglion cells after damage, and decrease microglial activation in vivo (Ueki et. al., 2012; Ueki et. al., 2013; Mitchell et. al., 2013; Thompson et al., 2019). However, the timing and mechanism(s) by which BMP/TGFβ signaling modulates repair in the mammalian retina is not fully understood. It is also unclear how these two homologous pathways interact in this tissue. ❧ To determine if the dual role of BMP signaling is conserved in the mammalian retina, we induced separately either light damage or excitotoxicity in C57BL/6J mice. We found BMP downstream signaling is activated in retinal support cells, the Müller glia. We then coupled this treatment with supplementation of BMP4 or inhibition of BMP using small molecules. We found that under both injury models, BMP4 promotes tissue survival under damage, while inhibition of BMP negatively affects tissue survival after light damage. ❧ Lastly, we sought to determine how innate immune cells of the eye were affected by the BMP/TGF? pathways. We accomplished this by analyzing changes in morphology which have been shown to reflect activation (Karperien et al., 2013). We found that, in comparison to control, with nearly all immune cells displaying highly ramified morphology, inhibition of BMP led to significant reduction in ramified morphology, while supplementation led to a notable increase. ❧ In conclusion, our data identify novel evolutionarily conserved mechanisms by which retinal homeostasis is maintained. My work is first known to show that Dpp and TGFβ/BMP signaling act as central controllers of photoreceptor death and survival after acute damage within the retina.

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Creator Kramer, Joshua Phillip (author)

Core Title Mediating the homeostatic response during light & cytotoxic induced retinal damage

School Leonard Davis School of Gerontology

Degree Doctor of Philosophy

Degree Program Biology of Aging

Degree Conferral Date 2021-08

Publication Date 08/05/2021

Defense Date 08/05/2021

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

Tag BMP,bone morphogenic protein,cytotoxicity,Drosophila,homeostasis,Light,mouse,OAI-PMH Harvest,retina,TGFβ,transforming growth factor beta

Format application/pdf (imt)

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Lithgow, Gordon (committee chair), Davis, Kelvin (committee member), Jasper, Heinrich (committee member), Lamba, Deepak (committee member), Tower, John (committee member)

Creator Email jpkramer@usc.edu,jpkramer1@gmail.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-oUC15710809

Unique identifier UC15710809

Legacy Identifier etd-KramerJosh-10007

Document Type Dissertation

Format application/pdf (imt)

Rights Kramer, Joshua Phillip

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

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