EXPLORING THE ROLES OF MÜLLER GLIA AND ACTIVATED LEUKOCYTE CELL
ADHESION MOLECULE A IN ZEBRAFISH RETINAL REGENERATION
By KRISTIN M. ALLAN
Submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy
Department of Molecular Medicine
CASE WESTERN RESERVE UNIVERSITY
January 2021 CASE WESTERN RESERVE UNIVERSITY SCHOOL OF GRADUATE STUDIES
We hereby approve the thesis/dissertation of Kristin M. Allan Candidate for the degree of Doctor of Philosophy*
Bela Anand-Apte, MBBS, PhD, MBA Committee Chair
Alex Yuan, MD, PhD Thesis Advisor
Aleksandra Rachitskaya, MD Clinical Mentor
Tara DeSilva, PhD Committee Member
Takuya Sakaguchi, PhD Committee Member
Date of Defense December 10, 2020
*We also certify that written approval has been obtained for any proprietary materials contained therein
TABLE OF CONTENTS
LIST OF TABLES ...... v
LIST OF FIGURES ...... vi
ACKNOWLEDGEMENTS ...... ix
ABSTRACT ...... 1
CHAPTER 1: INTRODUCTION ...... 3
Anatomy and physiology of the retina ...... 3
The neural retina ...... 3
Retinal glia ...... 4
Retinal diseases and disorders ...... 8
Age-related macular degeneration ...... 8
Retinitis pigmentosa and other retinal dystrophies ...... 9
Diabetic retinopathy ...... 11
Regenerative medicine for treatment of retinal degeneration ...... 12
Retinal regeneration ...... 15
Zebrafish as a model system for human therapy and disease ...... 15
Regenerative potential of Müller glia ...... 16
Prominent regenerative mechanisms in the zebrafish retina ...... 18
CHAPTER 2: FLOW CYTOMETRIC CELL SORTING OF ZEBRAFISH MÜLLER GLIA ...... 23
Introduction ...... 23
Papain-based tissue dissociation ...... 24
i
Cells of the vertebrate retina ...... 24
The retinal extracellular matrix ...... 25
Materials and methods ...... 27
Fish and retina dissection ...... 27
Dissociation of adult zebrafish retinal tissue ...... 28
Sample preparation for flow cytometric cell sorting ...... 29
Instrumentation and gating strategy ...... 30
RT-PCR analysis ...... 30
Retina flat-mount ...... 30
Microscopy ...... 31
Results ...... 33
Müller glia in Tg(apoe:gfp) retinas ...... 33
Cell suspension counts and morphology ...... 33
Flow cytometric cell sorting of Müller glia ...... 34
Discussion ...... 41
Procedural considerations and troubleshooting ...... 43
Applications ...... 45
CHAPTER 3: THE ROLE OF ALCAMA IN ZEBRAFISH RETINAL REGENERATION ...... 48
Introduction ...... 48
Progenitor cell migration ...... 48
Activated leukocyte cell adhesion molecule A ...... 51
Alcama in retinal development and regeneration ...... 53
ii
Rationale, study goals, and scope ...... 55
Materials and methods ...... 64
Zebrafish ...... 64
Retinal injury – laser photocoagulation ...... 64
Morpholino treatments ...... 65
RNA extraction ...... 66
RT-PCR and qPCR ...... 67
EdU treatment and staining ...... 67
Immunohistochemistry – Zpr1 ...... 68
Immunohistochemistry – Alcama ...... 69
Microscopy ...... 69
Quantification of EdU lineage tracing assay ...... 70
SLO imaging and quantification ...... 71
Zpr1 immunofluorescence quantification ...... 72
Results ...... 77
Zebrafish Müller glia upregulate Alcama expression in response to injury ...... 77
Morpholino-mediated knockdown of Alcama ...... 78
Alcama inhibition affects progenitor cell migration during retinal regeneration .... 81
The regenerative process may be inhibited or delayed with Alcama knockdown ... 84
Alcama inhibition decreases cone regeneration ...... 85
Discussion ...... 99
Alcama expression in the zebrafish retina ...... 100
iii
Morpholino-mediated Alcama knockdown ...... 101
The functional role of Alcama in retinal regeneration ...... 104
CHAPTER 4: CONCLUSIONS AND FUTURE DIRECTIONS ...... 109
Conclusions ...... 109
Future directions ...... 112
Proposed mechanistic studies involving Alcama ...... 112
The extracellular matrix and regeneration ...... 119
The role of microglia in retinal regeneration ...... 121
Concluding remarks ...... 123
APPENDIX 1 ...... 127
Primary cell culture of zebrafish retinal cell suspensions ...... 127
Cell culture methods ...... 127
Immunocytochemistry methods ...... 128
Preliminary characterization of primary zebrafish retinal cell cultures ...... 132
Assessing the proliferative capacity of primary zebrafish retinal cell cultures ...... 132
Cell type-specific markers ...... 134
Applications and Future Directions ...... 139
Dissociation and primary culture of mouse Müller glia ...... 140
APPENDIX 2 ...... 143
LITERATURE CITED ...... 146
iv
LIST OF TABLES
Table 1: Typical flow cytometric cell sorting results for Tg(apoe:gfp) zebrafish retinal cell suspensions ...... 37
Table 2: Common pitfalls in the preparation and sorting of Müller glia from zebrafish retinal cell suspensions ...... 47
Table 3: Primer sequences for RT-PCR experiments ...... 143
Table 4: Antibodies used in immunohistochemistry or immunocytochemistry applications ...... 144
v
LIST OF FIGURES
Figure 1. Anatomy of the eye and retina ...... 7
Figure 2. Fundus photos of patients with various degenerative disorders that could someday benefit from regenerative therapy ...... 14
Figure 3. Comparison of human and zebrafish eyes ...... 20
Figure 4. Paradigm of zebrafish retinal regeneration after outer nuclear layer injury 21
Figure 5. Proposed models for various regenerative signaling mechanisms in zebrafish retinas ...... 22
Figure 6. Key steps and overall workflow for the dissection, dissociation, and preparation of a single cell suspension from zebrafish retinas of suitable quality for flow cytometric analysis ...... 32
Figure 7. Three-dimensional reconstruction images of flat-mounted retina from Tg(apoe:gfp) zebrafish ...... 38
Figure 8. Zebrafish retinal cell suspensions suitable for flow cytometric cell sorting . 39
Figure 9. Representative gating strategy for Tg(apoe:gfp) zebrafish retina cell suspensions ...... 40
Figure 10. Proliferating progenitor cells migrate from the inner to the outer nuclear layer as regeneration is occurring ...... 57
Figure 11. Various biochemical and physical signals integrated by the extracellular matrix ...... 58
Figure 12. Structural organization of activated leukocyte cell adhesion molecule ..... 59
Figure 13. Protein sequence alignment of zebrafish Alcama and human ALCAM ...... 60
Figure 14. Alcama inhibition in zebrafish embryos leads to a reduction in eye size and retinal cell loss ...... 61
vi
Figure 15. Retinal defects in ALCAM-knockout mice ...... 62
Figure 16. Neurogenic proliferating progenitor cell clusters surround an Alcama- positive Müller glia after injury in adult zebrafish retinas ...... 63
Figure 17. Targeted retinal injury using OCT-guided laser photocoagulation model .. 74
Figure 18. SLO imaging analysis quantification strategy ...... 75
Figure 19. Zpr1 (cone density) quantification strategy ...... 76
Figure 20. Alcama expression is specifically upregulated by Müller glia in response to injury in the zebrafish retina ...... 87
Figure 21. ALCAM expression in mouse retinas is restricted to the inner plexiform layer, ganglion cell layer, and choroid before and after injury ...... 88
Figure 22. Morpholino delivery and mechanism of action ...... 89
Figure 23. Validation of morpholino-mediated Alcama knockdown ...... 90
Figure 24. EdU lineage tracing assay outline and representative images ...... 92
Figure 25. Movement of EdU-positive cells within retinal lesions over time ...... 93
Figure 26. Total EdU-positive cell counts in lesions for EdU lineage tracing assays .... 94
Figure 27. SLO imaging analysis shows increased lesion sizes with Alcama morpholino treatment at some time points ...... 95
Figure 28. Representative images from SLO imaging analysis ...... 96
Figure 29. Immunohistochemistry of cone photoreceptors in control and Alcama morpholino-treated retinas at 14 days post lesion ...... 97
Figure 30. Comparison of in vivo imaging and immunohistochemistry as methods for assessing retinal regeneration ...... 98
Figure 31. Proposed model for Alcama-mediated mechanism of action ...... 125
vii
Figure 32. Microglia infiltrate the injury site after laser ablation injury ...... 126
Figure 33. Comparison of zebrafish whole retina primary culture grown on various substrates ...... 131
Figure 34. GFAP co-labels with ApoE:GFP expression to confirm Müller glia identity in culture ...... 135
Figure 35. Zebrafish whole retina primary cultures are Nestin-positive but PCNA- negative ...... 136
Figure 36. Primary retinal cultures incorporate BrdU when stimulated with IL-6 and at baseline ...... 137
Figure 37. Cell clusters around Müller glia primarily consist of amacrine/ganglion cells, bipolar cells, and cones ...... 138
Figure 38. SLO images of cone-labeled fish after laser injury ...... 145
viii
ACKNOWLEDGEMENTS
There are so many things I would like to say here that I am not quite sure where to start.
I think this is where I’m supposed to reflect fondly on the past several years and talk about
how much I am going to miss everything I’ve grown accustomed to during my time here.
While there is some truth to that statement, it does not fully reflect my outlook on
graduate school. To be frank, I did not enjoy it. I contemplated quitting more times than
I can count, struggled through each and every day, week, month, and year, but managed
to keep going until the very end. For this I am incredibly proud, something I do not allow
myself to say very often. Grad school expanded my scientific knowledge in immeasurable
ways, but it also taught me a lot about myself and helped shed light on my future goals.
More importantly, I am forever indebted to all of the amazing individuals who have been
there for me every step of the way, many of whom probably have no idea how much their
support has meant to me.
First I will of course thank my advisor, Alex Yuan, who took me in as his first
graduate student, an experience I think we both learned and grew from in a lot of ways,
both personally and professionally. I joined the Yuan lab seeking some extra clinical
exposure and a mentor that would be supportive of my career plans, and I was certainly
not disappointed. From the opportunities to be involved in clinical projects to the words
of wisdom and counsel any time I felt like giving up, I am beyond grateful. Next, thank you
to each and every one of my committee members for their continued support and
guidance: Bela Anand-Apte, Tara DeSilva, Takuya Sakaguchi, and Aleksandra Rachitskaya.
I always seemed to enter committee meetings feeling apprehensive and unsure of myself,
ix
but I always left them full of renewed energy, confidence, and optimism because of the
wonderful mentorship you all provided. An extra thank you to my Chair, Bela, for helping
me see that I could do this and encouraging me to keep going on several occasions when
I thought I was going to quit.
To the members of the Yuan lab, past and present, I can’t thank you all enough:
Rose DiCicco, Becky Schur, and Michael Ramos. I could not have gotten through these last four and a half years without you. Rose, you put your heart and soul into this lab and have always been willing to help with anything I needed, and I truly appreciate everything you have done. Becky, I know we didn’t get to work together for very long, but I really valued your advice and suggestions, and I am sure you will continue to be a great leader for the lab. Michael, you were always so helpful in the lab and really helped drive experiments forward in ways I couldn’t have done on my own, for which I am grateful, but even more so for the many “life talks” we had; I think we helped each other through a lot of tough times, and I wish you the best of luck in medical school and beyond.
I am so thankful to have found the Molecular Medicine Program and can’t imagine grad school without all of the wonderful people I have met and worked with because of it. My classmates: Noah Daniels, Emma Keller, Alyson Wolk, Megan Zangara, and John
Zhou. I remember meeting all of you at interview weekend and being so excited to see all of our names together on the incoming class list. You are all incredible individuals, and I can see that we have all grown so much. An extra thanks to Aly, my fellow Ophthalmic
Research student, for lending a helping hand around the department and always being available for much-needed coffee breaks; and to Noah, for our many post-S-cubed chats,
x which always got me through the extra hard weeks. I also want to thank Sarah Kostiha for being an amazing program manager, having a true open-door policy and offering a listening ear or shoulder to cry on any time, and for fostering my growth as a young professional.
Finally, and most importantly, my family: I know over the last several years, most of you have no idea what I do or what I’m talking about a lot of the time, but that never mattered. To Craig and Jenni, I’ve looked up to you for literally my entire life. You have both made sure I have a lot to live up to, and I know you will always be in my corner, cheering me on. I can’t wait to add another signature graduation picture to our collection!
To Mom and Dad, you have always supported me 110% in whatever I do, and grad school has been no different. You listen to my struggles and triumphs week after week, and I cannot tell you how much your love and support means to me. I hope I’ve made you proud, which is my goal in all aspects of life. And to Ryan, thank you does not even begin to cover it. You have been my rock through it all, encouraging me through each and every day, and making sure I end every day with a smile and feeling like I can take on the world.
I truly could not have done this without you.
xi
Exploring the Roles of Müller Glia and Activated Leukocyte Cell Adhesion Molecule A
in Zebrafish Retinal Regeneration
ABSTRACT By KRISTIN M. ALLAN
In contrast to humans, animals such as zebrafish are able to regenerate their retinas after injury. Retinal cells called Müller glia facilitate regeneration in fish, yet contribute to scar formation in humans. Zebrafish Müller glia respond to injury by dedifferentiating and undergoing an asymmetric cell division, giving rise to a Müller glia and a proliferating progenitor cell. Subsequent progenitor cell migration and differentiation restores proper retina structure and function. The mechanisms governing the migration and functional integration of these progenitor cells, however, are not yet understood. A cell-surface
adhesion molecule called Alcama, a novel marker of activated zebrafish Müller glia,
contributes to cell migration, ganglion cell axonal guidance, and retinal lamination during
development in both fish and mice. We hypothesize that Alcama, expressed in Müller glia
following injury, facilitates the migration of progenitor cells and is important for zebrafish retinal regeneration. Here, we describe how Müller glia were isolated from the heterogenic population of retinal cells in Tg(apoe:gfp) fish with fluorescence activated cell sorting (FACS). The GFP-high population of cells represent Müller glia, which only express Alcama after injury, a pattern that is unique to this population. We have previously shown that proliferating progenitor cells migrate from the inner to outer
1 nuclear layer of the retina following injury under normal conditions. To determine
Alcama’s role in regeneration, we knocked down its expression in the adult retina with in vivo electroporation of antisense morpholinos targeting Alcama. Using EdU lineage tracing, we have observed significant differences between control and Alcama morpholino-treated eyes in the proportion of EdU-positive cells in the inner and outer nuclear layers over time, suggesting that Alcama is playing a role in progenitor cell migration during regeneration. We have also seen a delay in overall regeneration after
Alcama morpholino treatment using in vivo SLO imaging to measure lesion area over time, as well as a reduction in regenerated cone cell density using immunohistochemistry. This work will contribute to a deeper understanding of mechanisms governing regeneration in zebrafish for future translation into human therapies to restore lost vision.
2
CHAPTER 1 INTRODUCTION
Anatomy and physiology of the retina
The neural retina
Our sense of sight is dependent on the ability of our eyes to capture and convert incoming
light signals to stimuli interpretable by the brain.1 The eye is a complex organ comprised of multiple tissue types that work together to produce functional vision (Fig. 1A). Most
directly responsible for this phenomenon is the retina, a highly complex and specialized
tissue covering the back of the eye, commonly regarded as an extension of the brain.2,3
Consequently, retinal cells are similar in some ways to their counterparts in the rest of
the central nervous system, but also exhibit many features specific to the retina.2,4,5
The retina is comprised of 18 transcriptionally unique cell types that play unique structural and functional roles, organized in layers of cell bodies and processes to facilitate the process of phototransduction6–9 (Fig. 1B). This process is initiated by the
posterior-most retinal layer of cells called photoreceptors, specifically rods and cones,
specialized neurons whose nuclei comprise the outer nuclear layer (ONL).10 Proteins called opsins contained within the photoreceptor outer segments undergo a light-induced
conformational change, which initiates a signaling cascade, culminating in polarization of
the photoreceptor to generate an electrical response that is propagated along complex networks of retinal cells.11–13 Moving anteriorly from the ONL is the outer plexiform layer
3
(OPL), a non-nuclear layer formed from the synapses between photoreceptors and other retinal neurons including bipolar and horizontal cells, whose nuclei reside in the inner
nuclear layer (INL); similarly, the inner plexiform layer (IPL) is formed from the synapses
between INL neurons and ganglion cells, whose nuclei comprise the ganglion cell layer
(GCL).14 Ganglion cell axon projections converge as the nerve fiber layer to form the optic
nerve, ultimately responsible for transmitting visual stimuli to the brain for
interpretation3,7,8,14 (Fig. 1B).
Retinal glia
In addition to the retinal neurons that are most directly responsible for facilitating the process of phototransduction, three major classes of glial cells also exist in the retina: microglia, astrocytes, and Müller glia.2 These cells not only play supportive roles to the
other cells, but also have their own important functions in maintaining retinal physiology.
Microglia are the resident immune cells of the central nervous system, including
the retina.15–17 In a healthy retina, microglia exhibit many branched and ramified
processes, dynamically surveying their surroundings and phagocytosing cellular debris when necessary.18,19 In this way, microglia are central to the process of synaptic pruning
in both the developing and mature brain, in which they facilitate the elimination of
unnecessary neuronal connections by engulfing cells and remodeling synapses, regulated by neuronal activity.20–22 In an unhealthy retina, microglia are activated and shift to an
enlarged, amoeboid morphology.19,21,23 Microglia may be activated in response to
disturbances in normal retinal homeostasis such as infection, injury, and other disease-
related damage, mediated by a variety of inflammatory responses.2,21,24,25 Note that there
4
is some disagreement in the field surrounding the distinction between microglia and
macrophages recruited in response to injury. Although it is generally accepted that both
activated microglia and monocyte-derived macrophages are involved in immune- modulated responses to damage, they share many of the same markers and morphology,
making it difficult to define roles that are unique to each.24–27 The chemokine receptors
CX3CR1 and CCR2, for example, are particularly controversial; CX3CR1-positive/CCR2- negative cells are reported by some to specifically identify microglia,26,28–30 but are found to be expressed at similar levels in both microglia and macrophages by others.24,31,32
In addition to microglia, two types of macroglia also reside in the retina, astrocytes
and Müller glia. Astrocytes are most well-studied in the brain but are present to limited
extents within the retina, primarily in association with vasculature and ganglion cell
axons.2,33 As the zebrafish retinal vasculature lies on top of the ganglion cell layer as
opposed to being incorporated throughout the retina itself in mammalian retinas, there
is a negligible presence of astrocytes within zebrafish retinas, primarily residing in the
nerve fiber layer and optic nerve.34,35 Astrocyte functions will therefore not be discussed at length here, as most are accomplished by Müller glia, discussed below. Incidentally, glial fibrillary acidic protein (GFAP) expression is often used in mammalian systems to identify astrocytes;33 in zebrafish retinas, however, GFAP is specific to Müller glia.34–37
Müller glia are the dominant glial cell of the retina, spanning its entire thickness
with extensive radial processes that interact with every cell type and consequently
provide metabolic and structural support to the entire retina.2,38,39 For example, Müller
glia extend their processes apically to interdigitate with the outer segments of
5
photoreceptors, supporting their fragile connecting cilia, while offering necessary
nutrients in exchange for disposal or recycling of metabolites, and are poised to receive
signals in times of stress.40,41 Müller glia processes also extend radially around cell bodies of photoreceptors, bipolar cells, and other retinal neurons42 (Fig. 1B). Müller cells extend
basally and project endfeet along the ganglion cell and nerve fiber layers, forming the
internal limiting membrane, separating the retina from the vitreous body.38 Similarly,
their processes envelop blood vessels not only for the purpose of nutrient exchange, but
also for establishing the blood-retina barrier.43–45 Müller glia are able to maintain their
physical and signaling contacts with the rest of the retina through a variety of ion
channels, receptors, transporters, and ligands distributed across their surfaces.42,43
Because of these intimate associations, Müller glia are regarded by some as the epicenter
of column-like functional units.46,47 Interestingly, this is mirrored in the process of regeneration, at the heart of which lies Müller glia (discussed in the proceeding sections).
6
A
B
Figure 1. Anatomy of the eye and retina. A) Diagram of the human eye. B) Schematic of the major cellular components and their organization in a mammalian retina. Layers: Ch, choroid; PE, (retinal) pigment epithelium; OS, outer segments; ONL, outer nuclear layer; OPL, outer plexiform layer; INL, inner nuclear layer; IPL, inner plexiform layer; GCL, ganglion cell layer; ON, optic nerve; NFL, nerve fiber layer. From Vecino, et al., 2016.2
7
Retinal diseases and disorders
Numerous genetic and environmental factors can affect the retina leading to impaired
vision and, in some cases, total blindness. A recent survey of U.S. adults indicates that loss
of eyesight is regarded as one of the most devastating conditions a person can experience,
alongside cancer and Alzheimer’s Disease.48 Loss of vision can result from a variety of diseases, inherited retinal dystrophies, and other disorders affecting the retina either directly or secondarily.49 While the pathophysiology varies widely between different
retinal conditions, the end result is loss of eyesight because of impaired cellular function
and an irreversible loss of cells through the process of retinal degeneration that will be
the focus of this section.50–52 Regenerative therapy serves as a valuable therapeutic tool
with the potential to treat multiple diseases.53 Just a few of the most prevalent
retinopathies are detailed below as examples of the many heterogeneous retinal
degenerative disorders that could potentially be treated by regenerative therapy (Fig. 2).
Age-related macular degeneration
Age-related macular degeneration (AMD) is the most prevalent retinal degeneration,
affecting over 10 million individuals in the United States and an estimated 170 million
worldwide.50,54,55 Because of the rapidly aging population, the incidence of AMD is likely
to increase dramatically in the coming years, with a projection of 288 million people
affected by the year 2040.56 AMD is a multifactorial retinopathy that primarily affects the
area of central vision called the macula. The progression of AMD is determined by a
variety of genetic and environmental factors, with aging being perhaps the most
8
significant risk factor.55,57 For example, diets rich in antioxidants may decrease the risk of
AMD onset, while smoking is reported to increase the risk substantially.57–60 There is also
significant evidence for at least 20 specific genetic variants linked to AMD risk. For
example, AMD susceptibility loci have been identified near genes involved in extracellular
matrix remodeling such as TIMP3 (tissue inhibitor of metalloproteinase 3) and ADAMTS9
(a disintegrin-like and metalloproteinase with thrombospondin type 1 motif 9), and
several complement associated genes including the first specific AMD susceptibility locus
to be identified, CFH (complement factor H), suggesting an immune component in AMD
progression.55,61–63
There are two subtypes of AMD, dry and wet. Dry AMD, accounting for up to 90%
of AMD cases, is characterized by the gradual loss of RPE with consequential loss of the
photoreceptors it supports and currently has no treatment64,65 (Fig. 2B). Patients are
encouraged to increase antioxidant intake in their diets, though it remains unclear
whether this strategy effectively slows the progression of the disease.65 Wet AMD has a
neovascular component, often resulting in more rapid vision loss due to leakage,
hemorrhage, and/or edema from the new discordant vessels.66,67 The few therapies
available for wet AMD, such as anti-VEGF (vascular endothelial growth factor) injections,
only help slow the progression of the disease and are inconsistently or minimally
effective, especially long-term.68,69
Retinitis pigmentosa and other retinal dystrophies
In contrast to AMD, which primarily affects tissue underlying the retina with
consequential photoreceptor cell death, the inherited condition of retinitis pigmentosa
9
directly impacts the photoreceptors themselves70 (Fig. 2C). Retinitis pigmentosa (RP) is a
progressive disease resulting in the gradual loss of photoreceptors, and affects
approximately 1 in 4000 individuals.71,72 Photoreceptor degeneration is initially restricted to rods, impairing peripheral and night vision, eventually spreading centrally and also damaging cones, potentially leading to total blindness.71 The diagnosis, management, and
therapeutic strategies for RP are particularly challenging: the inheritance pattern may be
autosomal-dominant, autosomal-recessive, X-linked, or may be sporadic.70,72 Onset of
disease can occur at virtually any age depending on severity of symptoms and time of
presentation, though is most often diagnosed between early and mid-adulthood.71,73,74
To date, causative mutations in more than 100 genes and loci have been identified.75
While RP is the most common, many other forms of inherited retinal dystrophies
also lead to devastating vision loss through their own mechanisms, the heterogeneity of
which is highlighted by the identification of over 250 causative genes.75–77 For example,
Stargardt Disease is a progressive, inherited macular degeneration with onset usually
occurring in childhood and is most commonly caused by mutations in the ABCA4 gene,
though its clinical presentation varies widely and does not necessarily correlate with the
particular mutation.78,79 Conversely, genotype/phenotype correlations do tend to exist
for Leber Congenital Amarosis (LCA), which usually presents in infancy, and may result in a slow, RPE dysfunction-dependent retinal degeneration, or more severe photoreceptor defect depending on the causative mutation.80 To date, LCA has over 20 known causative
gene mutations, most of which are autosomal-recessive including RPE65, for which exists
the first FDA-approved gene therapy for an inherited disease.75,80–82 While gene therapy
10
is a promising path forward in some cases, the immense heterogeneity of inherited retinal dystrophies demonstrates a need for additional or complementary therapies.
Diabetic retinopathy
Although not an inherent retinal disease, diabetic retinopathy is a complication experienced by about one-third of the 422 million patients with the systemic disease diabetes mellitus, making it the leading cause of vision loss in working age adults.70,83–85
The retinal vasculature responds to diabetic hyperglycemia initially with increased
permeability and vessel occlusions, eventually progressing to include neovascularization,
hemorrhage, and even retinal detachment due to excessive fluid build-up or fibrosis86,87
(Fig. 2D). Because these pathologic responses cause significant metabolic changes,
neuronal cell death and retinal degeneration can eventually result, further contributing
to the metabolic disruption.88–90 Current therapies primarily treat the vascular
dysfunction central to the condition;70 future treatments could conceivably consist of
regenerative therapy to restore lost cells in conjunction with vascular therapy for
maintenance. Retinal fibrosis and scarring can also occur in response to these changes as
part of the tissue’s attempt to protect itself.87 Scar formation can be intentionally induced
using laser photocoagulation, a common treatment of diabetic retinopathy in addition to
anti-VEGF therapy, to prevent the spread of vision-harming effects.86 Ironically, while scar
formation is inherently neuroprotective either as a natural response or induced, it is
similar in many ways to the process of regeneration, yet scar formation also impedes
regeneration in the mammalian retina.91,92
11
Regenerative medicine for treatment of retinal degeneration
Perhaps one of the largest barriers in treating retinal degeneration lies in the differences
in pathology and progression of so many heterogeneic diseases. Because many of these
ultimately result in the loss of retinal cells, regenerative therapy offers a viable treatment
option for many of these conditions.70,93 Several strategies have emerged as promising
avenues to explore for generating retinal neurons.94 Using a variety of cell culture conditions and differentiation protocols, many groups have reported generation of retinal precursors and mature retinal cells from embryonic stem cells (ESCs) and induced pluripotent stem cells (iPSCs).95–97 Similarly, retinal organoid systems are also being
developed in three-dimensional cell culture environments utilizing iPSCs.98–100 These
studies may be the first steps in establishing feasibility for transplantation of retinal
grafts.101,102 Many of these protocols, however, are extremely time consuming; they may
also be better served as tools for modeling disease or pharmacological screening rather
than for direct use in patients for the foreseeable future.103,104
One of the major challenges with transplantation of progenitor cells and
differentiated neurons or grafts is their integration with native retinal circuitry, and it
remains uncertain as to if or how this effectively occurs.105–107 A less common approach
is direct stimulation or reprogramming of resident cells, particularly Müller glia, in vivo
through genetic or pharmacologic manipulation in mammalian systems to give rise to
functional neurons that do appear to integrate with the existing circuitry, but this effect
is quite limited.108,109 Further exploration of mechanisms that inherently promote
functional regeneration in animals such as zebrafish can advance all of these strategies by
12
improving the formation of retinal progenitor cells and integration of the new neurons.
The limited demonstration and study of how transplanted cells and newly generated retinal neurons are able to integrate into existing circuitry is a primary reason for our study of how progenitor cells migrate during zebrafish retinal regeneration in Chapter 3.
13
A C
B D
Figure 2. Fundus photos of patients with various degenerative disorders that could someday benefit from regenerative therapy. A) Normal retina; healthy vasculature, clear macular and optic disc. B) Age-related macular degeneration (dry); advanced deterioration of the RPE and photoreceptors around the macula. C) Retinitis pigmentosa; characteristic bone-spicules of RPE visible from peripheral photoreceptor cell death. D) Diabetic retinopathy; hemorrhage, micro aneurysms, and ‘cotton wool spots’ thought to represent damage to the nerve fiber layer.
14
Retinal regeneration
While humans and other mammals experience retinal degeneration in numerous ways,
they cannot replace lost or damaged cells, resulting in a permanent loss of vision.110 Some
animals such as zebrafish, however, have the inherent ability to regenerate their retinas
and restore functional vision.36,111–114 The current treatment options for most retinal
conditions are extremely limited, or nonexistent; we therefore use zebrafish as a
comparative model to fully understand how the regenerative process occurs in order to
someday translate that knowledge into therapy for human use.
Zebrafish as a model system for human therapy and disease
Human and zebrafish retinas appear to be very different in terms of overall size and shape
because they are adapted to such different environments (Fig. 3). The fish lens, for
example, is round and proportionally much larger than the ellipsoid human lens, giving it
a higher refractive index.115,116 Similarly, the fish iris is stationary and does not change size to adjust the amount of light passing to the retina as it does in humans; the fish compensates with a highly reflective iris and movement of RPE into the outer segments to shield them from phototoxicity under photopic conditions. Unlike most other animals, the human retina has a macula, the cone-dense area of central vision. Another difference
worth noting is in the vasculature: while human retinal vasculature is found within the
retina itself, the retinal vasculature in zebrafish lies on top of the tissue.34,35
Although some clear differences exist, the same major structures are present in
both zebrafish and human eyes. Importantly, the cell types and organization of the retina
15
are highly conserved (Fig. 3).117 Zebrafish are over 70% homologous with humans, and
84% of known disease-causing genes in humans have an orthologous gene in zebrafish.118
This structural and genetic similarity, in addition to their inherent regenerative capacity,
make the zebrafish an ideal vertebrate model system for studying human eye disease and
regenerative therapy.
Regenerative potential of Müller glia
The heart of regenerative potential in both zebrafish and human retinas lies with their
Müller glia. Although human retinas do not regenerate in response to damage, the Müller
glia respond in some ways that parallel the regenerative response seen in zebrafish.114
Müller glia in humans and other mammals exhibit a gliotic response to damage and form a scar. This occurs through the process of reactive gliosis, where the Müller glia continually proliferate but hypertrophy and become fibrotic.91,119 Increased expression of
intermediate filament proteins such as GFAP, vimentin, and nestin, is a major component
of this process, increasing the physical rigidity of the Müller cells.91,120,121 Scar formation
is paradoxically protective and detrimental to the retina: reactive glia limit the extent of
damage to the tissue by secreting neurotrophic factors, but prolonged reactivity further damages the retina and the scar itself impairs vision and prevents any functional regeneration from occurring.91,122 Understanding how to shift Müller glia response to damage from reactive gliosis to functional regeneration will be a key component for the development of successful regenerative therapies.
Müller glia comprise a functional stem cell niche within the adult zebrafish retina.36 The proliferative response of Müller glia seen in reactive gliosis is mirrored in the
16
regenerative process of zebrafish Müller glia; however, their Müller glia first
dedifferentiate and undergo only a single, asymmetric cell division, giving rise to a Müller
glia and a progenitor cell, which continues to proliferate.111,113,123 It is this re-entry into
the cell cycle and generation of multipotent progenitor cells that is the defining aspect
between gliosis and scar formation in mammals but successful regeneration in fish.114,124
The progenitor cells eventually differentiate to replace the cells lost to damage, the
identity of which depends on the form of injury used to target certain parts of the retina
and activate the Müller glia (Fig. 4).113,123,125
Common injury paradigms used to study subsequent regeneration of the zebrafish
retina include: puncture injury involving a needle poke through the entire retina, resulting
in progenitor cell differentiation into all retinal cell types;125,126 oubain treatment
primarily damages the inner retina, and the regenerated cells therefore include ganglion,
amacrine, and bipolar cells;127–129 and outer retina damage is achieved by targeting the
photoreceptors such as with phototoxicity through intense light exposure or selective
photoreceptor ablation with laser injury.130–132 All of the resulting regenerated cone cells arise from Müller glia-derived progenitor cells that also give rise to some new rod cells; however, rods will also be generated from rod progenitor cells, which may be derived
from multipotent progenitors but also persist in the ONL and contribute to the continued
growth of the retina throughout the zebrafish lifespan.130,133–135 We primarily use the laser
injury paradigm and will therefore be focusing on the mechanisms involved in
regeneration of ablated photoreceptors.
17
Prominent regenerative mechanisms in the zebrafish retina
Significant progress has been made in the field to identify numerous mechanisms,
signaling molecules, and cellular processes that play key roles in zebrafish retinal
regeneration (Fig. 5).136 Activation of Müller glia is initiated by signals given off from
injured or dying neurons,114,137 immune cells responding to the damage,138,139 and from
the Müller glia themselves through positive feedback mechanisms.140–142 Identifying what these signals are is often the first step in determining the downstream mechanisms at
play throughout the regenerative process. For instance, increased HB-EGF (heparin
binding epidermal growth factor) expression is detected almost immediately after retinal
injury and has been shown to be necessary and sufficient for Müller glia stimulation and
subsequent regeneration (Fig. 5B).142 Furthermore, HB-EGF stimulation induced Müller glia activation in damaged avian and mouse retinas.143 Downstream, HB-EGF stimulates
multiple key signaling cascades including MAPK, Jak/Stat, Notch, and Wnt, which are also
activated by other injury-induced signals.142–144 CNTF (ciliary neurotrophic factor), an IL-6 family member, stimulates both MAPK and Jak/STAT signaling.110,145,146 Multiple Wnt
ligands are upregulated in response to injury, and Wnt signaling is necessary to maintain
the proliferation of Müller glia-derived progenitor cells.147,148
Jak/STAT signaling is one of the most prominent pathways activated in Müller glia in the initiation of the regenerative process. Stimulation of Jak/STAT3 through treatment with IL-6 family members has been shown to be necessary and sufficient to induce a regenerative response in Müller glia of uninjured zebrafish retinas (Fig. 5A).149 In response
to injury in a murine system, Jak/STAT signaling is increased,150 yet does not lead to
18
regeneration but to gliosis,110,146,151 demonstrating that there are many aspects of
regeneration to be considered in conjunction to fully understand the process. Similarly,
increased Notch signaling in fish has an inhibitory effect on progenitor cell formation and is suppressed following retinal injury; however in mammals, increased Notch signaling is required for Müller glia dedifferentiation, yet must be inhibited to promote subsequent cell cycle exit and differentiation (Fig. 5C).114,141,152
Many of these and other key regenerative pathways converge on the same
transcription factor that may be a cornerstone for regenerative therapy: Ascl1a (Achaete-
scute homolog 1a)153 (Fig. 5). In fact, forced expression of Ascl1 in a mouse model of
retinal damage is sufficient to induce Müller glia-derived progenitor cells that give rise to
functional neurons to some extent.108 In a follow-up study, the same group showed that
inhibiting Jak/STAT signaling actually enhanced the extent to which their model of injury
and Ascl1 expression can produce functional neurons in a mammalian system.109 This
highlights the dynamic nature of the regenerative process and the balance of processes
such as proliferation that must be found to produce a functional regenerative response
in mammals. An understanding of how regeneration is occurring as a complete picture,
as opposed to identifying individual therapeutic targets, is therefore an essential step to
translation.
19
Homo sapiens Danio rerio
Figure 3. Comparison of human and zebrafish eyes. Left: human (Homo sapiens) eye representations. Right: zebrafish (Danio rerio) eye representations. Top: external appearances. Middle: cross-sectional diagrams of major eye structures, from Chhetri et al., 2014.115 Bottom: histological retina sections, from Richardson et al., 2017.117 NFL, nerve fiber layer; GCL, ganglion cell layer; IPL, inner plexiform layer; INL, inner nuclear layer; OPL, outer plexiform layer; ONL, outer nuclear layer; IS, inner segments; OS, outer segments; PR, photoreceptors; RPE, retinal pigment epithelium.
20
Figure 4. Paradigm of zebrafish retinal regeneration after outer nuclear layer injury. In response to outer nuclear layer injury, such as from light-induced photoreceptor cell death, the Muller glia in fish respond by dedifferentiating (green) and undergo a single asymmetric cell division to produce a Muller glia and a progenitor cell (pink). The progenitor cells proliferate, and then migrate along the Muller glia, eventually differentiating into photoreceptors to reconstitute the retina. Adapted from Nagashima et al., 2013.123
21
A B
C
Figure 5. Proposed models for various regenerative signaling mechanisms in zebrafish retinas. A) Injury stimulates production of signaling molecules such as IL-6 and leptin, which converge on the Jak/Stat pathway to drive HB-EGF and Ascl1a expression to promote regeneration. Adapted from Zhao et al., 2014. B) Injury stimulates HB-EGF expression, driving Ascl1a activity to ultimately drive regenerative pathways and feedback mechanisms. Adapted from Wan et al., 2012. C) Injury signals such as TNFα stimulate MAPK and other pathways, and lift the inhibitory effect of Notch signaling, to induce Müller glia dedifferentiation and subsequent regeneration. From Wan and Goldman, 2016.136 These models highlight the extremely complex interplay of numerous signaling mechanisms involved in zebrafish retinal regeneration.
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CHAPTER 2 FLOW CYTOMETRIC CELL SORTING OF ZEBRAFISH MÜLLER GLIA
This chapter is adapted from the article originally published in Cytometry Part A: Preparing a single cell suspension from zebrafish retinal tissue for flow cytometric cell sorting of Müller glia.154
Introduction
Flow cytometry is a technique to analyze individual cells based on visual light scattering
and fluorescent signals labeling cell markers to establish unique molecular signatures.155
While flow cytometry is traditionally used for analytical purposes, specialized instrumentation has expanded the field to include cell sorting, allowing for the physical separation and collection of cell populations of interest for downstream applications.156
Flow cytometric cell sorting first requires a sample suitable for running on a flow
cytometer, which entails preparing a cell suspension from solid tissue if analysis of
primary cells is desired. A clean and reliable cell suspension for flow cytometric analysis
must avoid excessive cell death, damage, and aggregation.
Although many previous publications indicate using cell sorting as part of their
experimental workflow, we have found no reports offering the necessary detail to
generate a quality cell suspension from zebrafish retinas with sufficient
reproducibility.140,157–162 This chapter details the preparation of a single cell suspension from zebrafish retinal tissue, optimized for the subsequent flow cytometric cell sorting of
23
Müller glia. Using cell suspensions from Tg(apoe:gfp) transgenic fish retinas, we can isolate and collect Müller glia expressing GFP under control of the ApoE promoter, specific to Müller glia in the zebrafish retina.36 Because of their inherent ability to give rise to a functional stem cell niche, zebrafish Müller glia are of particular interest for regenerative medicine and cell therapy research.111,123,135,163–165
Papain-based tissue dissociation
A major determinant of cell preparation quality is the method of tissue dissociation, which should be optimized for the application of interest.154,166 We use a papain-based dissociation of adult zebrafish retinas to obtain a single cell suspension suitable for subsequent flow cytometric cell sorting. Papain is a cysteine protease that digests many components of the retinal extracellular matrix more effectively and gently than other commonly used enzymes like dispase, trypsin, or collagenase.157,158,167,168 Several vertebrate models have been used to demonstrate that papain is the least damaging and most effective proteolytic enzyme used to dissociate cells from a variety of delicate neural tissues, including the retina.167–172
Cells of the vertebrate retina
Eight major classes of neural and glial cells comprise the highly specialized and complex tissue that is the vertebrate retina. Each cell type plays their own structural and functional roles within the retina, reflected in their unique phenotypic and morphologic variations.6,7
Retinal cells are arranged in distinct laminations of cell bodies and processes that work together in complex networks to facilitate the retina’s primary function of
24
phototransduction.8 Histologically, the retina is organized into three nuclear layers,
containing cell bodies, intercalated by two plexiform layers comprised of cell
processes.14,173 (Fig. 1B, Fig. 3).
The major retinal glial cells are Müller glia, which provide structural metabolic, and functional support to virtually every cell in the retina.38,39 In addition to helping
maintain retinal homeostasis, Müller glia also have inherent regenerative potential in the
vertebrate retina. While Müller glia respond to damage in mammals by contributing to
scar formation, they give rise to a functional stem cell niche that regenerates the retina
in animals such as zebrafish.123,165,174,175 This makes Müller glia of particular interest in the
progression of regenerative therapies and underscores the importance of developing
tools to study them, such as a reliable and reproducible method for isolation. Although
the overall size and shape of ocular structures varies between humans and fish because they are adapted to such different environments, the same major cell types are present in both, and their organization is highly conserved.176,177 The zebrafish retina is therefore
a useful model with which to study retinal cell biology and regeneration with the added
benefit of translational feasibility.
The retinal extracellular matrix
Just as diverse and complex as the cells they support are the non-cellular constituents of
the retina, making up its extracellular matrix. Extracellular matrix generally consists of
various proteins, carbohydrates, glycoproteins, proteoglycans, fibers, and other factors
such as enzymes and signaling molecules secreted by the surrounding cells.178,179 The
composition of extracellular matrix in different parts of the retina and its surrounding
25
tissue reflects the heterogeneity of the retina and other ocular tissues. The most
prominent extracellular matrix components in the adult retina include laminins,
collagens, tenascin-C and –R, and chondroitin sulfate proteoglycans.180–184 Fibronectin is
a glycoprotein commonly found in the extracellular matrices of a variety of tissues, but it is not a significant component of the adult retina.178 Not only does the extracellular matrix
bring structural integrity to the retina, but it also plays an active role in a variety of
signaling processes on a molecular level including axonal guidance, angiogenesis, cell
polarization, and retinogenesis.185–187 Because it is directly involved in such dynamic and developmentally important processes, the composition of the extracellular matrix therefore changes during development. Laminin, collagen IV, and fibronectin, for example, are transiently expressed at certain stages of retinal development.180
Effectively disrupting interactions between cells and their surrounding tissue is an
essential part of obtaining a retinal cell suspension suitable for flow cytometry. Obtaining
intact Müller glia is particularly challenging because of their large size and extensive
processes. The dissociation must therefore be thorough enough to disrupt Müller glia
interactions with surrounding cells and extracellular matrix, but gentle enough to
preserve Müller glia integrity. Here, we detail a dissociation procedure of adult zebrafish
retinas that routinely results in a clean cell suspension with over 90% viability while
keeping Müller glia intact for subsequent flow cytometric cell sorting.
26
Materials and methods
Fish and retina dissection
Before dissection, wild-type or Tg(apoe:gfp) zebrafish 6-18 months of age were dark
adapted overnight. This causes the retinal pigmented epithelium (RPE) to move away
from the photoreceptor outer segments, which promotes a clean retina dissection with
minimal RPE contamination.184,188 Fish were euthanized using the IACUC-approved
procedure of placing them one at a time in a divided tank, opposite the side containing
ice, until unresponsive to tail pinch. Eyes were enucleated immediately after sacrifice with
a blunt, curved edge eye dressing forceps (Miltex 18-781, or similar). To enucleate, eyes
were rotated until the curved part of the forceps is under the eye and resting in the eye
cavity. The forceps were then closed to pinch the optic nerve and pulled up gently to
remove the eye, which was placed in dissection media (EBSS + 1% penicillin/streptomycin)
and held on ice.
Eyes were removed from the dissection medium and held on a sterile petri dish
with Dumont #1 forceps, then punctured at the limbus with a 23-30 ga needle. Starting
at this puncture, VANNAS microscissors were used to dissect away the iris and lens from
the posterior chamber, removed from the eye cup with forceps. To remove the retina
from the eye cup, Dumont #1 forceps were used to hold the sclera while the retina was
peeled away from the eye cup with Ekhardt or similar retina forceps (20-25 ga). If the
optic nerve was still intact, it was clipped with microscissors to fully separate the retina
from the eye cup. Retinas were then minced with the dissecting scissors with at least 12-
15 perpendicular cuts to make approximately equal sized pieces. The minced tissue was
27
transferred to a 15 mL conical tube, containing a small amount of dissecting medium, in
a minimal volume (<50 μL) of additional dissecting medium with a p1000 pipette, held on
ice. After all retinas were collected, additional dissecting medium was used to bring each
sample up to the appropriate volume(s) (see below).
Dissociation of adult zebrafish retinal tissue
Retinal tissue was dissociated using the Worthington Papain Dissociation System. A vial
of papain (Worthington, #LK003176) was reconstituted in EBSS that was equilibrated with
95% O2 : 5% CO2 by passing through a 0.22 μm filter over the surface of the liquid, then incubated at 37 °C for 10 minutes or until fully dissolved to activate the enzyme. The
volume of all tissue samples to be dissociated plus the volume of EBSS used to
reconstitute the papain should be equal to 5 mL, for a final working solution of 20
units/mL papain in 1 mM L-cysteine with 0.5 mM EDTA. After reconstituting a vial of
DNase I (Worthington, #LK003170) in 500 μL equilibrated EBSS, 250 μL was added to the
activated papain, giving a working solution of 0.005% DNase I. This papain/DNase solution
was added to the tissue samples in volumes proportional to the number of retinas
contained within each sample.
The tissue was then triturated by pipetting up and down about 10 times, gently
but thoroughly with a p1000 pipette. The samples were then equilibrated with 95% O2 :
5% CO2 as described above before incubating at 37 °C with 160 rpm agitation in a shaking
incubator for 10-12 minutes. Trituration was repeated before returning to the incubator
for an additional 10-12 minutes. Samples were then centrifuged at 850 x g for 5 minutes
at room temperature. The supernatants were pipetted off and pellets suspended in
28 inhibitor solution (0.95 mg/ml ovomucoid inhibitor [Worthington, #LK003182] and
0.00475% DNase I in equilibrated EBSS), maintaining the appropriate volume ratio previously established. The samples were centrifuged again at 850 x g for 5 minutes at room temperature, supernatants removed, and the pellets were suspended in 550 μL sterile, room temperature sorting buffer (1% FBS, 1 mM EDTA, 25 mM HEPES, pH 7.0-7.4 in 1x PBS). This cell suspension was passed through a 30 μm filter into a clean tube; the original tube was rinsed with an additional 550 μL sorting buffer and also passed through the filter for a final 1 mL cell suspension, taking volume lost through the filter into account
(Fig. 6).
Sample preparation for flow cytometric cell sorting
Cell suspensions were mixed well, and a dilution was loaded into a hemocytometer
(Hausser Scientific, Bright-Line, Improved Neubauer), viewed with a light microscope at
10x magnification to count the four corner quadrants. The concentrations of cell suspensions were calculated by averaging the corner quadrant counts, multiplying by the dilution factor, and multiplying by the volume of a small square (e.g. 104, hemocytometer- specific). Since the cell suspensions are 1 mL, the concentration is also the total number of cells obtained from the dissociation. Additional sorting buffer was added so that the final cell suspension to be sorted was at 4-4.5million cells/mL.
For compensation controls, DRAQ7 DNA dye was diluted 1:100 (3 μM final concentration) in an aliquot of WT cells for a DRAQ7-only control; the remaining WT cells served as the unstained control; and an aliquot of Tg(apoe:gfp) cells was set aside for the
GFP-only control. DRAQ7 was added 1:100 (3 μM final concentration) to the remaining
29
Tg(apoe:gfp) cell suspension, which is the sample to be sorted, labeled with both GFP and
DRAQ7.
Instrumentation and gating strategy
The BD FACSAriaII was used with a 100 μm nozzle at 20 psi for sorting cell suspensions.
The blue laser (488 nm) was used to excite the native GFP signal in Tg(apoe:gfp) cell
suspensions, detected by the 515/20 filter (505 LP dichroic). The red laser (640 nm) was
used to excite the DRAQ7 dye, detected by the 730/45 filter (690LP dichroic). The gating
strategy outlined in Figure 9 was used to separate and collect live, single cells in the
desired populations.
RT-PCR analysis
GFP-negative, GFPlow, and GFPhigh cell populations were sorted directly into TRIzol LS
(Invitrogen) to isolate total RNA according to the manufacturer’s protocol. Oligo d(T)
primers, random hexamers, and MultiScribe Reverse Transcriptase (TaqMan Reverse
Transcription Reagents, Invitrogen) were used to generate cDNA from isolated RNA. PCR
reactions used EmeraldAmp GT PCR MasterMix (Takara Bio) and gene-specific primers
(Table 3).
Retina flat-mount
Retinas for flat-mount were dissected from dark-adapted Tg(apoe:gfp) fish as described
previously, but were kept intact and transferred to a clean glass slide. A 15 ° angled blade
was used to make radial cuts in four quadrants so that the retina laid flat before mounting
with Vectashield.
30
Microscopy
The Leica TCS SP8 confocal scanning microscope (Leica Microsystems Europe) was used to image retina flat-mounts. The 488 nm laser, HyD detector, and HC PL APO CS2 40x/1.30 oil objective were used to obtain three-dimensional z-stacks, imaged through the entire thickness of the retina and processed with the 3D Viewer in the Leica Application Suite X
Software. Cell suspensions prepared for sorting were imaged using the Zeiss Axio Imager
Z1 upright fluorescence microscope (Carl Zeiss Microscopy, LLC). Small drops (10-20 μL) of cell suspensions were placed on a glass slide and covered with a glass coverslip. Trypan
Blue was first added in a 1:1 ratio to a separate aliquot to assess viability before sorting.
The plan‐NEOFLUAR 20x/0.5 objective was used with EGFP (apoe:GFP), Cy5 (DRAQ7), and
DIC channels (brightfield) to obtain images, processed using ZEN Pro 2.5 imaging software.
31
Figure 6. Key steps and overall workflow for the dissection, dissociation, and preparation of a single cell suspension from zebrafish retinas of suitable quality for flow cytometric analysis. Refer to the text for a detailed description of all procedures.
32
Results
Müller glia in Tg(apoe:gfp) retinas
In Tg(apoe:gfp) zebrafish retinas, Müller glia specifically express GFP under control of the
ApoE promoter.36 This is particularly apparent by three-dimensional reconstruction of
Tg(apoe:gfp) flat-mount retinas, where the GFP-positive cells are undeniably Müller glia, as no other retina cell type exhibits their unique morphology (Fig. 7). Müller cell processes
are extensive and span nearly the entire thickness of the retina. These observations not
only confirm the identify of GFP-positive cells as Müller glia in Tg(apoe:gfp) retinas, but
also underscore the importance of thorough and gentle separation of these cells from the
rest of the retina.
Cell suspension counts and morphology
Cell suspensions generated from 6-18 month old zebrafish retinas with the technique
detailed in this chapter routinely yield an average of about 800,000 cells/retina. Younger
or smaller retinas will yield lower counts, while older or larger retinas tend to contain
more cells. After the final cell suspension is obtained, a trypan blue exclusion assay is
recommended to preliminarily assess cell viability before proceeding to cell sorting. This
can easily be done by diluting the cell suspension used to count 1:1 by volume with a 0.4%
trypan blue solution. The dye will only be taken up by dead or dying cells, staining them
blue, and viable cells will remain impermeable, excluding the dye.189 An initial estimate of cell viability is therefore obtained by counting the blue and non-blue cells. Cell suspensions with less than 70% viability or excessive amounts of cell debris and
33 aggregates are not recommended for cell sorting or other flow cytometry applications.
Our cell preparation procedure routinely results in 90-99% overall cell viability (Table 1).
This is illustrated in Figure 8A, where only 1 cell out of 13 is stained with trypan blue, to give an initial estimate of about 92% live cells. This is in agreement with the proportion of cells stained with DRAQ7, a fluorescent dye that functions similarly to trypan blue, as viewed under the microscope (Fig. 8C) or analyzed by flow cytometry (Fig. 9). The latter shows 93% live cells, reflecting the number of DRAQ7-negative cells at the moment they pass through the flow cytometer.
With our cell preparation technique, which uses a combination of enzymatic
(papain) and mechanical (trituration) disruption of retinal tissue, we consistently produce a cell suspension free of clumps with minimal debris, indicative of a clean and thorough dissociation (Fig. 8). Importantly, this procedure is gentle enough to preserve Müller glia morphology. When cell suspensions sit on a slide for several minutes, the Müller glia elongate and extend their processes, demonstrating that even after dissociation, they remain intact (Fig. 8B) and reminiscent of their characteristic appearance in vivo (Fig. 7).
These images further highlight the necessity of using a large nozzle (100 μm) and corresponding low pressure (20 psi) when sorting Müller glia to minimize the possibility of their processes being sheared as they are sorted.
Flow cytometric cell sorting of Müller glia
The gating strategy we have employed for isolating Müller glia from a Tg(apoe:gfp) zebrafish retinal cell suspension is outlined in Figure 9. Cells are first selected based on forward and side scatter, excluding any contaminating debris, which exhibit higher side-
34
to-forward scatter ratios than intact cells (Fig. 9A). Aggregate correction is then used to
eliminate clumps of multiple cells based on side scatter width vs. height (Fig. 9B) and on
forward scatter width vs. height (Fig. 9C). Live cells are then identified as DRAQ7-negative,
excluding any that are dead or dying and have consequently taken up DRAQ7 dye (Fig.
9D). The single, live cells are finally split into three distinct populations based on their
native GFP signal: GFP-negative, GFP-low, and GFP-high (Fig. 9F). A wild-type, unstained
cell suspension is always included as a control and should contain 100% GFP-negative cells
(Fig. E) to establish the GFP-negative gate. In Tg(apoe:gfp) cell suspensions, 77-83% of all
sorted cells are GFP-negative, while 12-17% are GFP-low and 2-4% are GFP-high (Table 1).
We acknowledge that the cut-off between the GFP-low and GFP-high cells is somewhat
arbitrary, reflective of the necessary subjectivity in distinguishing between two
populations in flow cytometry; however, this cut-off remains consistent between
experiments.
We have confirmed that the GFP-high population contains Müller glia by RT-PCR
analysis of glial fibrillary acidic protein (gfap) expression (Fig. 9G). Gfap is an intermediate
filament protein and a classical marker of Müller glia.36,124,132,190 Gfap is expressed only by the GFP-high population, absent in both the GFP-negative and GFP-low populations (Fig.
9G). This expression pattern, along with the characteristic morphology exhibited by the
GFP-positive cells in Tg(apoe:gfp) retinas (Fig. 7), has led us to conclude that cells of the
GFP-high population are Müller glia. Furthermore, we have observed that while the small amount of GFP signal in GFP-low cells is detectable by the BD FACSAriaII, it is not visible when imaged by confocal microscopy. We are hypothesizing that the GFP-low population
35
is likely comprised of other retinal cells that have taken up small amounts of GFP protein
exuded by the GFP-expressing Müller glia by means of intercellular transport, such as in
extracellular vesicles. Making the appropriate distinction between the GFP-low and GFP- high populations and subsequent confirmation of Müller glia identify is therefore essential before selecting cells with which to proceed in downstream applications like
RNA-sequencing.
36
Table 1. Typical flow cytometric cell sorting results for Tg(apoe:gfp) zebrafish retinal cell suspensions
Cell viability GFPhigh GFPlow GFPnegative 90-99% 2-4% 12-17% 77-83%
37
A
B C
Figure 7. Three-dimensional reconstruction images of flat-mounted retina from Tg(apoe:gfp) zebrafish. A) 3D reconstruction of Tg(apoe:gfp) retina (300 x 300 x 100 μm). B, C) Zoomed-in and rotated perspectives of (A). The GFP-positive, elongated processes spanning nearly the entire thickness of the retina exemplify Müller glia morphology, indicating the specificity of the ApoE promoter for Müller glia in the zebrafish retina. All images are oriented with the internal limiting membrane (Müller glia-vitreous junction) at the top and external limiting membrane (Müller glia-photoreceptor junction) at the bottom of the images.
38
Phase Contra A B C
st/Merge
apoe:gfp
DRAQ7
Figure 8. Zebrafish retinal cell suspensions suitable for flow cytometric cell sorting. A) Trypan blue staining, showing one dead trypan blue-positive cell (arrow). B) Example Müller glia (GFP+) that has extended its processes after sitting under a coverslip for several minutes. C) Representative fluorescence of cell suspension samples going through the flow cytometer: three GFP+ Müller glia and two dead, DRAQ7+ cells. Scale bars are 50 μm.
39
G
Figure 9. Representative gating strategy for Tg(apoe:gfp) zebrafish retina cell suspensions. A) Cells are separated out from debris based on forward and side scattering. B) Aggregate correction on side scatter. C) Aggregate correction on forward scatter. D) Viable cells selected by exclusion of DRAQ7 dye, which fluoresces at a wavelength of 700 nm. E) Unstained control. F) Single, live cells clustering into three populations based on GFP expression. G) RT-PCR analysis of gfap expression alongside rpl13a (ribosomal protein) housekeeping control gene in each cell population, indicating that the GFP-high population contains Müller glia.
40
Discussion
This protocol uses a combination of enzymatic and mechanical methods to dissociate retinal tissue, digesting the extracellular matrix to leave the intact cellular components in suspension. The dissociation is carried out so that interactions between cells are disrupted enough to minimize cell clumps and aggregates, but without over-dissociating or damaging the cells themselves. This is particularly challenging in the retina, where cellular interactions are varied and complex. Photoreceptors, ganglion cells, and other retinal neurons, for example, propagate messages of visual stimuli between one another via synapses, connected by gap junctions that are comprised of connexin proteins in vertebrates.191–195 Anchoring junctions, including adherens junctions and desmosomes,
also play essential roles in upholding the structural integrity and corresponding
functionality of the retina.196–198 Perhaps the most intricate connections are those of the
Müller glia themselves; they form contacts with every major retinal cell type, as their
processes extend through nearly the entire thickness of the retina.43,199 A notable
example is the intimate association of Müller glia microvilli with photoreceptor outer
segments, important both structurally and functionally for a variety of processes,
including phototransduction and maintenance of photoreceptor metabolism.40
Furthermore, Müller glia processes form extensive contacts with other glial cells, retinal
neurons, and blood vessels to provide necessary metabolic support and signaling
molecules throughout the retina.43,200
All of these interactions must be completely but gently disrupted, executed in our hands with the Worthington Papain Dissociation System to generate a single cell
41
suspension suitable for flow cytometric cell sorting. Papain is known for its ability to
efficiently and gently dissociate delicate tissues, particularly effective on neuronal
tissue.168–171 We have used a single kit to dissociate as many as 30 adult zebrafish retinas,
though the manufacturer indicates a single kit can dissociate up to 0.4 cm3 of total
tissue.172 Multiple kits can be combined to accommodate larger tissue quantities and,
conversely, a single kit may be used to dissociate multiple smaller tissue samples. In the
latter case, it is important to establish appropriate volume ratios of dissociation reagents
throughout the entire process to maintain the necessary concentrations of enzymes and
buffers, detailed in this chapter’s Materials and Methods.
We use an exclusively papain-based dissociation, which contrasts with most
previously published methods. Other protocols often employ harsher enzymes such as
dispase,159 accutase,140 or trypsin,157,158,160 either alone or in conjunction with papain. A recent single-cell analysis of Müller glia from zebrafish and other species offers the only known report of a retinal dissociation using only papain; however, it details neither the dissociation protocol nor the cell sorting results,161 reflecting a common lack of important
procedural details in the literature as a whole. Many reports that do offer a description
of their protocol are aiming to sort cells other than Müller glia such as
microglia/macrophages157 and photoreceptors157,158,160 from zebrafish retinas. Any
studies that are sorting Müller glia from zebrafish retinas utilize the Tg(gfap:gfp)
transgenic line.140,159,161,162 To our knowledge, our use of the Tg(apoe:gfp) fish for flow cytometric cell sorting of zebrafish Müller glia is therefore the first to be reported.154
Furthermore, most of these studies do not give details such as cell viability; one group,
42
however, does report overall viability of about 95%, comparable to our own.160 More
importantly, previous studies sorting Müller glia make no mention of a distinction
between GFP-high and GFP-low cell populations.140,159,161,162 Because Müller glia provide
metabolic support to all other cells in the retina,38,39,43,200 it is not unexpected that some
GFP expressed by Müller glia might be given off and taken up by other cells, regardless of
the Müller glia-specific promoter that controls GFP expression. This caveat should therefore be carefully considered when analyzing sorting data from previous and future studies. The consideration is especially important in those involving large-scale
characterizations such as single cell RNA-sequencing, as the results could be easily skewed
by including GFP-low population cells mistaken for Müller glia.
Procedural considerations and troubleshooting
When preparing a cell suspension from zebrafish retinas that is suitable for flow cytometric cell sorting of Müller glia, several common problems may occur. Table 2
summarizes key steps in this process with their corresponding pitfalls and how to avoid
or overcome them. It is important to find a balance of complete but gentle dissociation in order to separate as many cells as possible from their surrounding tissue without compromising their integrity. Mincing the dissected retinal tissue before proceeding with
dissociation, for example, improves the dissociation efficiency by increasing the surface
area of tissue accessible to the digestive enzymes.166 Similarly, multiple short incubations
in papain alternated with gentle trituration through an appropriately sized pipette tip (i.e.
p1000) effectively achieves this balance of a complete-but-mild dissociation. This is in
contrast to dissociation procedures involving one continuous incubation in harsher
43
enzymes,158–160 and rigorous trituration with narrower instruments such as a needle or fire-polished pipettes,140,160 methods which may result in more dead and damaged cells.
In addition, equilibration is not a practice reported by other retina dissociation
procedures, despite being important for maintaining physiological pH throughout the
dissociation to prevent aggregation and improve viability of dissociated cells. Foregoing
equilibration steps may therefore compromise the efficiency of dissociation and quality
of resulting cell suspensions. Conditions of centrifugation are also important to consider
during the later steps of dissociation. Samples are spun at 850 x g, which may be
considered high when compared to typical speeds for live cell centrifugation; however,
zebrafish retinal cells are particularly buoyant, and we have found that lower speeds
result in decreased yields from loss of cells in the supernatants. Centrifuging at room
temperature helps to prevent clumping when suspending in the appropriate buffers,
which are also used at room temperature. Finally, the use of a viability dye like DRAQ7 in the final sample to be sorted is essential and should not be excluded. The addition of such a dye allows for more accurate quantitation than trypan blue and, importantly, ensures no dead or dying (i.e. DRAQ7-positive) cells are included in the collected populations at the time of sorting. Appropriate control samples must also be included to accurately establish gates for each unique experiment. Primary cell samples are recommended for this purpose as opposed to fluorescent beads to take any auto-fluorescence from primary cells into account.
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Applications
Once sorted, zebrafish Müller glia can be used for a variety of downstream applications, and the collection buffer should be chosen accordingly. Cells to be collected for molecular analysis of proteins or nucleic acids, for example, may be sorted directly into the appropriate lysis buffer at the temperature indicated by a particular protocol. Or, if analysis on live cells is desired, complete cell culture media should be used to collect populations of interest at room temperature. In either case, cells may be collected in
additional sorting buffer, centrifuged to pellet, and diluted to the desired concentration
in applicable buffer; however, we have found this to be difficult, as the cells do not pellet
well after sorting due to residual charges they may carry from the sorting process.
Furthermore, pellets are often not visible unless the concentration is at least 500,000
cells/mL, which is difficult to achieve when collecting the relatively rare population of
Müller glia unless there are a large number of retinas to use as starting material. The
methods to obtain a single cell suspension from zebrafish retinas as described in this
chapter, with subsequent cell sorting of Müller glia, serve as valuable tools for zebrafish
cell biology and regenerative ophthalmic medicine. While this protocol is specifically
optimized to collect Müller glia from Tg(apoe:gfp) fish retinas, the overall process is
generally applicable to the collection of other retinal cell types, preparation of cell
suspensions from similarly delicate or heterogenic neural tissues, and to additional flow
cytometric techniques to offer novel cell-specific insights.
While the cell preparation detailed in this chapter is optimized for the flow
cytometric cell sorting of Müller glia, the dissociation procedure is also applicable for cell
45 culture purposes, as many of the same principles for producing a cell suspension for flow cytometry are relevant for culturing primary cells. Using this dissociation procedure, we have developed a primary cell culture of whole retinal cell suspensions from zebrafish using this dissociation procedure; we have also adapted the protocol slightly for primary cultures of mouse Müller glia (Appendix 1).
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Table 2. Common pitfalls in the preparation and sorting of Müller glia from zebrafish retinal cell suspensions KEY STEP DO DON’T Retina dissection - Use scissors to mince retinas - Place whole retinas in collection thoroughly before transferring to tube collection tube in dissection medium
Papain dissociation - Be sure to add DNase to both the - Invert the tube at any time, as papain solution and inhibitor solution this will cause tissue to stick to to prevent clumping of exposed DNA sides of tube, decreasing from dying cells dissociation efficiency and -Use EBSS with a pH indicator viability -Skip the equilibration steps Mechanical dissociation - Triturate the dissociating tissue 5-10 - Use a pipette other than p1000 times before, during, and after size to triturate the tissue: a incubation in papain at 10 minute smaller size will be too harsh intervals while a larger size will result in an - Pipette up and down slowly and ineffective dissociation gently - Introduce bubbles into solution
Centrifugation - Spin samples at 850 x g (use RCF - Spin at less than 800 x g, units) resulting in loss of cells in the supernatant - Spin at greater than 900 x g, resulting in an overly compact cell pellet and increase damage to cells
Final cell suspension - Warm sorting buffer to room - Use cold buffer in final cell preparation temperature before using in final suspension, which will result in suspension of cell pellet clumping - Pass final cell suspension through a 30 μm gravity filter
Preparing samples for - Always prepare an unstained control - Prepare samples for sorting at flow cytometry sample greater than 5 million cells/mL, - Prepare single color control samples which may cause clumping, each time a new cell reduction in efficiency, and/or type/marker/fluorophore decreased yield combination is utilized
Cell sorting - Exclude dead cells and debris - Use a nozzle size less than 100 - Collect cells in appropriate buffer, μm, as this may cause clogging or depending on downstream exclusion of the large Müller glia applications cells
47
CHAPTER 3 THE ROLE OF ALCAMA IN ZEBRAFISH RETINAL REGENERATION
This chapter is adapted from a manuscript submitted (December 2020) for publication in Experimental Eye Research: Alcama facilitates the migration of progenitor cells in zebrafish retinal regeneration.
Introduction
The human retina is susceptible to degeneration from a variety of genetically and environmentally driven conditions. The resulting cell loss and damage is irreversible and leads to permanent vision loss. Animals such as zebrafish have the inherent ability to regenerate their retinas, restoring lost cells and functional vision. The process of regeneration is extremely complex and must be understood in its entirety before that
knowledge can be translated into viable regenerative therapies. While there has been
substantial progress in the field, especially in the initiation of the regenerative process,
some aspects remain largely unclear. We are particularly interested in gaining a better
understanding of how the migration of new cells occurs and contributes to the overall
process of regeneration.
Progenitor cell migration
In zebrafish, Müller glia respond to injury by dedifferentiating, re-entering the cell cycle,
and giving rise to a Müller glia and a progenitor cell, which proliferates. These early steps,
occurring within the first 72 hours of injury response, have been extensively
studied.123,132,201 Understanding how Müller glia give rise to progenitor cells, rather than
48 initiating the gliotic response that occurs in mammals, is a primary focus in the field and is the first step toward making regenerative therapy a reality. As this part of the regenerative process becomes clearer, we also need to understand how the newly generated cells migrate along the Müller glia, essential for their differentiation and integration into existing retinal architecture. Without this, restoration of functional vision cannot occur.
It is generally accepted that cellular migration occurs to some extent during retinal regeneration, but the ways in which it proceeds remain unclear.36,201,202 Time-course studies have shown clusters of progenitor cells surrounding Müller glia around 72 hours after injury, at which point progenitor cell proliferation is most robust.123,132,201 There is also evidence to suggest that some of the observed migration is due to interkinetic nuclear migration (IKNM) of the Müller glia nuclei within the cell moving apically toward the outer limiting membrane where Müller cells meet the photoreceptor outer segments.123,124 Most of these studies utilize a light-injury paradigm occurring over several hours, making the Müller glia response somewhat asynchronous and difficult to interpret. We therefore sought to better understand how progenitor cell migration is occurring with our laser injury method, which administers instantaneous damage to a small area of the retina where subsequent regeneration can be followed131 (Fig. 10). After injury, BrdU was injected IP at 3 days post lesion (dpl) to label the progenitor cells when their proliferation is most robust.132,201 These retinas were collected at various time points post-injury and stained for BrdU (Fig. 10A). By quantifying the number of BrdU-positive cells in each retinal layer at each time point, we can see that the progenitor cells migrate
49
out of the inner nuclear layer and into the outer nuclear layer over time (Fig. 10B-C). The
large majority of cells incorporating BrdU at 3 dpl reside in the outer nuclear layer by 14
dpl. These results demonstrate that the movement of BrdU-positive nuclei represent the
migration of progenitor cells rather than IKNM, which would have shown an initial
movement of the labeled nuclei toward the ONL, eventually returning to the INL. This also
supports the finding that IKNM likely occurs between one and two days post injury to
initiate the asymmetric division of Müller glia.123,202 Our results then suggest that the
proliferation of the resulting progenitors indeed occurs in the INL with subsequent cellular
migration. Now, we aim to elucidate some of the mechanisms underlying this process.
The extracellular matrix, cytoskeleton, and cell adhesion-dependent signaling are
interrelated phenomena that play major roles in facilitating the migration of neural
progenitor cells.201,203 The extracellular matrix (ECM) maintains the necessary microenvironment for an adult stem cell niche by regulating the exchange of signals required for multipotency and differentiation and by physically sculpting the spaces in which cellular morphogenesis can or cannot occur.204,205 In the central nervous system
specifically, the ECM promotes axonal guidance and synapse formation during development, but inhibits regeneration in mammals by contributing to glial scar formation.203,206 The ECM acts as a scaffold through which cells can move, facilitated by
interactions between specific cell adhesion molecules, mechanosensitive channels, or
other signaling pathways that enact the necessary changes on gene expression and
cytoskeleton to move the cell as the surroundings dictate (Fig. 11).204,207 ECM-cell interactions may directly contact or indirectly stimulate cytoskeletal rearrangement and
50
contraction through Rho/ROCK-mediated signaling, implicated in both progenitor cell and
IKNM in the retina and elsewhere.202,208–210 Integrins are perhaps the most well-studied
cellular ECM substrates, required for cell migration, proliferation, and patterning during
nervous system development.211,212
The adult stem cell niche is further comprised of the surrounding cells, adhesion molecules, and other contact-dependent signaling mechanisms, functioning in cell-cell interactions that are also required for progenitor cell migration.213 Cell adhesion
molecules integrate the interactions between cells, ECM, intracellular signaling mechanisms, and cytoskeleton dyamics.214 Cadherins, for example, bind calcium ions to
stabilize homophilic interactions between cells, initiating cytoplasmic signaling with β-
catenin, Rac1, and Rho/ROCK kinases, ultimately leading to cytoskeletal changes required
for cell migration of individual cells or collectively migrating clusters.215–217
Complementary to their role in facilitating migration, cadherins, integrins, and other cell
adhesion molecules activate cyclins/CDKs, MAPK, and Wnt signaling, playing active roles in maintaining the balance between proliferation and differentiation that is essential to neural progenitor cell function.213,214,218–220 Specifically, we are interested in
understanding the mechanisms that facilitate the migration and functional integration of
progenitor cells in the zebrafish retina. Here, we focus on a particular adhesion molecule,
Alcama, for its potential role in these processes during retinal regeneration.
Activated leukocyte cell adhesion molecule A
Alcama, or activated leukocyte cell adhesion molecule A, is a transmembrane
glycoprotein immunoglobulin superfamily member in zebrafish, orthologous to human
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ALCAM, and known by several aliases including DM-GRASP, CD166, Neurolin, BEN, and
SC1.221–223 Beginning at its amino terminus, Alcama consists of five extracellular immunoglobulin (Ig) domains, two variable (V) domains and three constant (C) domains
adjacent to the cell membrane, followed by a transmembrane domain, and a short
cytoplasmic carboxy-terminal tail (Fig. 12).221,224,225 The extracellular Ig domains,
comprising the majority of the protein, mediates both homophilic ALCAM-ALCAM
interactions and heterophilic interactions with CD6, a well-studied interacting partner;
interactions with other adhesion molecules such as NgCAM/L1CAM or E-cadherin have
also been implied.221,226–228 These interactions may be cis-acting oligomerization or
clustering of adhesion molecules on the surface of a single cell mediated by the (C)
domains, or trans-acting between other cells and mediated by the N-terminal (V)
domains, both of which ultimately facilitate connections between cells (Fig. 12).221,225
ALCAM expression in mammalian systems has been found in numerous cell and
tissue types, such as the central nervous system, epithelial cells, endothelial cells
(including those of retinal vasculature), and many stem cell populations such as cancer and hematopoietic cells.221,226 Consequently, ALCAM functions in a variety of processes,
ranging from T-cell activation and proliferation to axonal guidance, cell migration, and differentiation in both developing and adult organisms.221,224,226,229 As the extracellular Ig
domains mediate the interactions between cells, the highly conserved cytoplasmic tail
facilitates the appropriate intracellular signaling cascade(s) to initiate the corresponding cellular changes.226 ALCAM is not directly bound to the cytoskeleton but is indirectly
connected to it via the scaffolding proteins ezrin and syntenin-1, also aided by the
52
tetraspanin CD9 (Fig. 12A-C).221,230,231 Ubiquitination of the C-terminal tail has been
described as a key determinant of ALCAM endocytosis, which is an important factor in
regulation of function.221,232
Beyond these specific interactions, however, little is known about the mechanistic
underpinnings of the various functions facilitated by ALCAM, and much less is known
about zebrafish Alcama. Consequently, we must infer that most of what is known about
the function of this molecule from mammalian studies can be applied to some extent to
zebrafish Alcama, which is 38% identical and 34% similar to human ALCAM (Fig. 13),
consistent with the overall homology between humans and zebrafish of about 70%.118
Importantly, the functional Ig domains and cytoplasmic tail are conserved, demonstrating
structural similarity that often translates to corresponding function (Fig. 12D, Fig. 13).
Because we will be looking at Alcama in the context of zebrafish retinal regeneration,
however, for which humans do not have the capacity, most of the known background
information merely serves as a foundation for further research, and it will be necessary
to understand Alcama structure, function, and mechanisms of action during the
regenerative process in the zebrafish retina.
Alcama in retinal development and regeneration
In zebrafish, Alcama has almost exclusively been studied in a developmental context,
especially for its role in neurogenesis in the central nervous system, retina, and peripheral
motor neurons.223,233,234 In the retina specifically, Alcama has been shown to play
important developmental roles. When Alcama is inhibited in zebrafish embryos, for example, the overall size of the eye is significantly decreased and retinas are visibly
53
deformed (Fig. 14).223 Reduction in the number of cells in both the inner nuclear and
ganglion cell layers are primarily responsible for the smaller eye size. Retinal ganglion cells
are particularly affected when Alcama is inhibited during development, which prevents
their differentiation.223,233 This is perhaps due in part to Alcama’s role in axonal guidance.
Alcama knockdown in zebrafish embryos has been shown to disrupt the guidance of
motor neuron axons during development.234 This effect is phenocopied in ALCAM
knockout mice, which additionally exhibit defects in retinal ganglion cell axon
fasciculation, appearing wide and spread out compared to clear, tight bundles seen in
their wild-type counterparts (Fig. 15A).235 These mice also exhibit severe retinal
dysplasias, with disruptions to the normal laminations seen in healthy retinas (Fig. 15B), suggesting a role for ALCAM in cellular organization and patterning.
There is only one study to our knowledge that has recognized a potential role for
Alcama in the context of adult zebrafish retinal regeneration, where it is identified as a novel marker for dedifferentiated Müller glia.123 Here, Alcama expression is observed in
newly formed Müller glia in the ciliary marginal zone (CMZ), an adult stem cell niche that
contributes to the continuous growth of the fish retina throughout its lifespan, as well as
Müller glia in the INL responding to damage.123,133 After light injury, Müller glia extend
Alcama-positive processes through the thickness of the retina, and each one is
surrounded by a neurogenic cluster of proliferating progenitor cells, which would
eventually migrate to the outer nuclear layer (Fig. 16).123 They also demonstrated that
another adhesion molecule, N-cadherin, facilitates neurogenic cluster formation and
54 migration of progenitors to contribute to retinal neuron regeneration. N-cadherin may therefore be interesting to explore as a putative Alcama interaction partner.
Rationale, study goals, and scope
Much of the focus in retinal regeneration research has been on the ways in which Müller glia are activated and divide to regenerate the retina in animals such as zebrafish, instead of contributing to scar formation as seen in humans and other mammals. Inadequate attention has been given to studying another key aspect of regeneration, the migration of progenitors. Adhesion molecules act as bridges between the extracellular matrix and intracellular cytoskeleton and signaling cascades to facilitate the movement of cells in various biological processes. We are interested in a specific adhesion molecule, Alcama, for its potential role in zebrafish retinal regeneration. While there is substantial evidence to suggest that Alcama plays a role in cell migration and guidance, nearly all of this information has been gathered from a developmental context. And although there are often parallels between development and regeneration, only one study, to our knowledge, has suggested a role for Alcama during zebrafish retinal regeneration.123 This is only demonstrated by showing Alcama expression at a single time point (3 dpl), however, and evidence for its involvement in regeneration is merely circumstantial (Fig.
16). We are aiming to further characterize the expression pattern of Alcama before and during regeneration and, importantly, demonstrate a functional role for this particular adhesion molecule.
Our overall hypothesis is that Alcama facilitates the migration of progenitor cells and is therefore important for zebrafish retinal regeneration. We also hope to provide a
55 solid foundation on which to base future experiments aimed at elucidating specific mechanisms underlying Alcama’s role in zebrafish retinal regeneration. Furthermore, we hope that this study can serve as a proof of concept for exploring the potential roles of other adhesion molecules, as well as additional factors that may contribute to the migration of progenitor cells. This will aid in translation of regenerative therapies by contributing to the overall understanding of the regenerative process in zebrafish retinas, but may also serve as a tool for the application of future therapies, such as creating a scaffold or other means of guiding stem cells to their appropriate locations.
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A
B 3 dpl 4 dpl 7 dpl 14 dpl
C
Figure 10. Proliferating progenitor cells migrate from the inner to the outer nuclear layer as regeneration is occurring. A) Schematic of experimental design: day 0, each fish retina injured with three distinct laser lesions using our OCT-guided laser injury system; day 3, all fish receive intraperitoneal injections of BrdU (0.25 μmol/g body weight, two injections, two hours apart) to label progenitor cells when their proliferation is most robust; eyes collected and retinal sections stained for BrdU 4 hours later for the 3 day time point, or on days 4, 7, and 14. B) Representative images for each time point; all sections stained for BrdU (green) and counterstained with propidium iodide. C) Number of BrdU-positive cells counted in each retinal layer at the center-most 3-5 sections of lesions for indicated time points and plotted in terms of percentage of total BrdU-positive cells. INL, inner nuclear layer; ONL, outer nuclear layer; IPL, inner plexiform layer; NFL/GCL, nerve fiber layer/ganglion cell layer, quantified together. Data points represent averages of ‘n’ laser lesions quantified with standard error indicated by error bars. 3 days post lesion (dpl), n = 3; 4 dpl, n = 7; 7 dpl, n = 5; 14 dpl, n = 4. Two-tailed t-tests comparing INL to ONL are significant at all time points (p < 0.05), with the exception of 4dpl. Graph and calculations performed using GraphPad Prism 8.
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*
Figure 11. Various biochemical and physical signals integrated by the extracellular matrix. Schematic representation of the numerous roles played by the ECM. Most of these processes are facilitated by membrane proteins, including various adhesion molecules. Asterisks represent the ways in which Alcama itself could conceivably be interacting with the ECM and/or signaling intracellularly (color coded for the diagrammed processes). Adapted from Muncie and Weaver, 2018.204
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A
B
C
D
Figure 12. Structural organization of activated leukocyte cell adhesion molecule. A) Illustration of two ALCAM molecules, depicted as a homodimer. B) Key of ALCAM interaction molecules illustrated in A and C. C) Example ALCAM-ALCAM interactions; cis-ALCAM contacts are mediated by the C domains, while trans- acting contacts are mediated by the V domains; intracellular interactions with the cytoskeleton are supported by scaffold proteins ezrin and syntenin-1 and tetraspanin CD9. Adapted from von Lersner, et al., 2019.221 D) Schematic representation and comparison of ALCAM/Alcama protein domains between humans and zebrafish. Domain information obtained from UniProtKB for zebrafish Alcama (Q90460) and human ALCAM (Q13740). Blue rectangles represent V-type Ig domains; orange rectangles, C-type Ig domains; green hexagons, transmembrane domains; pink ovals, cytoplasmic C-terminal tail. Numbers above (zebrafish Alcama, Z) or below (human ALCAM, H) indicate the amino acid positions for the corresponding domains. Not to scale.
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Figure 13. Protein sequence alignment of zebrafish Alcama and human ALCAM. Protein sequences from zebrafish (Danio rerio) Alcama (NP_571075.1) and human (Homo sapiens) ALCAM (NP_001618.2) were aligned using the UniProt Align Tool CLUSTALO Program, demonstrating 37% sequence identity (216 amino acids) and 33% similarity (188 amino acids). Asterisks, highlighted in yellow, indicate identical amino acids; two dots, highlighted in green, indicate amino acids with highly similar properties; single dots, highlighted in blue, indicate amino acids with weakly similar properties.
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A B
Figure 14. Alcama inhibition in zebrafish embryos leads to reduction in eye size and retinal cell loss. Embryos were injected with either anti-Alcama/Neurolin-a morpholinos (MO-Na) to inhibit Alcama expression, or buffer control (PR) and analyzed in various ways and time points as indicated. A) Quantification of eye diameter relative to body size. Significant reduction in eye size is observed at 3 and 6 days post fertilization (dpf) when treated with 1 or 5 ng MO-Na compared to control. B) Cell numbers in both the retinal ganglion cell layer (RGCL) and inner nuclear layer (INL) are decreased by approximately 50% in embryos treated with 2.5 ng MO-Na compared to WT control. Error bars indicate standard deviations (A, B). C-F) Methylene blue-stained histological sections of eyes from embryos treated with buffer control (C, E) or Alcama morpholino (D, F) at 2 dpf (C, D) or 4 dpf (E, F) demonstrate the reduction in overall eye size and cell number with Alcama inhibition. F) Scale bar = 50 μm (C-F). Arrows indicate apoptotic cells. From Diekmann, et al., 2009.223
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A +/+ ALCAM ALCAM -/-
B -/- ALCAM +/+ ALCAM
Figure 15. Retinal defects in ALCAM-knockout mice. Comparison of retinas from WT and ALCAM (BEN) knockout mice. A) Retina flat-mounts from WT (ALCAM+/+, left) or knockout (ALCAM-/-, middle) mice, stained for neurofilaments to highlight retinal ganglion cell axons with quantification of axon fascicle widths (right) demonstrating significantly wider bundles in ALCAM-/- mice. B) Hemotoxylin and eosin-stained histological sections from WT and ALCAM-/- mouse retinas, demonstrating severe retinal dysplasias without ALCAM expression. Scale bar = 50 μm. Adapted from Weiner, et al., 2004.235
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A B
C
Figure 16. Neurogenic proliferating progenitor cell clusters surround an Alcama-positive Müller glia after injury in adult zebrafish retinas. Retina injured using intense light exposure, and sections stained at 3 days post lesion (dpl) in all images. A) Retinas from Tg(gfap:nGFP) fish, whose Müller glia nuclei express GFP, also stained for Alcama in white/magenta. Scale bar = 20 μm. B) Retinas from Tg(gfap:GFP) fish, whose Müller glia express cytoplasmic GFP, also stained for Alcama in white/magenta. Scale bar = 20 μm. C) Retinas from Tg(gfap:nGFP) fish, whose Müller glia nuclei express GFP, also stained for proliferation marker PCNA (red) and progenitor cell marker Rx1 (white/magenta), showing that clusters of proliferating progenitor cells surround Müller glia after injury, which are shown to express Alcama at the same 3 dpl time point in A and B. Scale bar = 5 μm. Adapted from Nagashima, et al., 2013.123
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Materials and methods
Zebrafish
Adult zebrafish are raised and maintained by husbandry staff at the Lerner Research
Institute (Cleveland, OH) on an Aquatic Habitats recirculating water system (Pentair;
Apopka, FL) in a 14:10 hr light:dark cycle at 28.5 °C. All procedures conform to the ARVO
statement for the Use of Animals in Ophthalmic and Vision Research. Protocols are
approved by the Cleveland Clinic Institutional Animal Care and Use Committee (Cleveland,
OH). Before enucleation, wild-type or Tg(apoe:gfp) zebrafish 6-18 months of age are dark
adapted overnight and euthanized by placing one at a time in a divided tank, opposite the
side containing ice, until unresponsive to tail pinch. For procedures requiring anesthesia,
fish are placed in 0.16 mg/mL tricaine in system (fish) water until unresponsive to external
stimuli.
Retinal injury – laser photocoagulation
Full details of the method summarized here can be found in our previous publication.131
Anesthetized fish are placed in a custom rubber holder and fitted with a
polymethylacrylate (PMMA) contact lens (Ø = 5.2 mm, r = 2.70 mm, center thickness = 0.4
mm; Cantor & Nissell, Ltd., Northamptonshire, UK) using Systane Ultra hydrating tears
(Alcon Laboratories, Fort Worth, TX). This ensures optimal imaging of the fish eye, which
is adapted to aquatic environments. Laser photocoagulation is carried out with an OCT
imaging system with a 7 μm axial resolution (SDOIS 840HR; Bioptigen, Durham, NC, USA)
fitted with a 532 nm diode laser (Oculight GL; Iridex, Mountain View, CA) using wide-field
64
objectives (≥50° field of view). The laser is incorporated into the OCT imaging path with a
custom beam combining module, which consists of a dichroic beamsplitter (685-nm long-
pass; Semrock, Inc., Rochester, NY), relay optics (Thorlabs, Newton, NJ), and a modified
200 μm multimode optical fiber (Fig. 17). An adjustable aperture set to 7.3 mm was used
to deliver a 300 ms pulse of 42-47 mW output power for each lesion. Laser spots were
targeted with the C-scan image and rotating the fish accordingly on a 5-axis positioning
stage (Model AIM-RAS; Bioptigen, Inc.). When histological sections are to be obtained,
three laser lesions are placed superior to the optic nerve, as seen in Figure 17. When
retinas are to be collected for FACS or expression analysis, as many lesions as allowed by
the physical and optical restraints of the system and time of anesthesia are placed
superior to the optic nerve; typically, 40-49 lesions.
Morpholino treatments
Custom-designed lissamine-tagged morpholinos (Gene Tools, LLC) were received as a lyophilized powder and reconstituted in sterile, deionized water to a 3 mM working solution. The anti-Alcama morpholino (5’-TGTGTTTAAGCTATGCTTACTGTGA-3’) is designed to target and bind a splice site acceptor sequence between exon 5 and intron 5, causing introduction of a premature stop codon and nonsense-mediated decay, ultimately resulting in the degradation of Alcama mRNA. The scrambled control morpholino (5’-CCTCTTACCTCAGTTACAATTTATA-3’) is comprised of a random sequence, not specific to any part of the zebrafish genome/transcriptome. Morpholino delivery is adapted from a previously described protocol (Fig. 22).236 Briefly, an anesthetized fish is placed on paper towels soaked in system water under a dissecting microscope at 4-5x
65
magnification for the duration of the procedure. The outer cornea is first removed using
one pair of Dumont #1 forceps (or similar) to stabilize the eye and another to pull off the
outer cornea. The tip of a 15-degree angle stab blade (LaserEdge Plus, PL7515) is used to make an incision in the superior-temporal region of the iris. A pulled 1 mm capillary
needle is then used to draw up 0.5 μL of 3 mM morpholino solution by capillary action.
The needle is inserted into the incision and injected intravitreally using a microinjector
with an injection time of 0.9 ms (Nikon PLI-188, Garden City, NY). The eye is gently rotated
within the socket with platinum-plated electroporation tweezers (CUY650P3), with the
negative electrode oriented behind the eye without touching it directly; the positive
electrode touches the nasal side of the globe to maintain the necessary rotation during
electroporation. A 75 V pulse is then delivered with a 50 ms duration (BTX ECM 830
Square Wave Electroporation System, Harvard Apparatus; connection cables for
electroporation tweezers: C115SCB-2). The fish is then returned to a tank of clean system
water to recover. This process takes approximately 5-7 minutes per fish and is completed
within 30 minutes of injury.
RNA extraction
For whole retina analysis, retinas are dissected out as described in Chapter 2, then
transferred whole to a nuclease-free Eppendorf tube with retina forceps, held on dry ice,
and stored at -80 °C. For RNA extraction, tubes are removed from freezer and cold TRIzol
reagent (Invitrogen) is added immediately before homogenizing with motorized handheld
homogenizer to lyse tissue, and the manufacturer’s protocol is followed to obtain RNA
with A260/280 around 1.8 and A260/230 around 2. For FAC-sorted Müller glia, retinas are
66
dissected, dissociated, and prepared for flow cytometric cell sorting as described in
Chapter 2 from Tg(apoe:gfp) fish, and RNA is extracted using the RNeasy Micro Kit
(Qiagen). Müller glia are sorted directly into Buffer RLT at 4 °C, mixing the contents
periodically throughout the sorting, making sure the volume of the sorted cells does not
exceed the volume of lysis buffer. RNA extraction then proceeds according to the
manufacturer’s protocol, with final RIN values of at least 7.
RT-PCR and qPCR
Total RNA was extracted as described above. Oligo(dT) and random hexamers were used with SuperScript IV Reverse Transcriptase (Invitrogen) to generate cDNA. Subsequent PCR reactions (RT-PCR) were performed with EmeraldAmp GT PCR Master Mix (Takara Bio
Inc.) and gene-specific primers (Table 3). Quantitative PCR (qPCR) was performed in triplicate with TaqMan Fast Advanced Master Mix (Applied Biosystems) using TaqMan
Gene Expression Assays (Applied Biosystems) for Alcama (Dr03073952_m1, FAM-MGB
Dye) or GAPDH (Dr03436842_m1, FAM-MGB Dye) on a QuantStudio 3 Real-Time PCR
System (Applied Biosystems). Relative quantification of Alcama mRNA was calculated
using the ΔΔCT method, normalized to GAPDH levels and compared to a single sample, performed with QuantStudio Design and Analysis Software.
EdU treatment and staining
Three days after retinal injury, 20 μL of 20 mM EdU solution was injected intraperitoneally
with a 33 ga needle at 3 days post lesion. Eyes were collected four hours later for a 3-day
time point, or at the indicated time point after dark adaptation for at least two hours.
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Enucleated eyes were placed in optimal cutting temperature media, flash frozen in a 2- methylbutane bath over liquid nitrogen, and stored at -80 °C before cryosectioning into
10 μm sections using CFSA 0.5x slides (Leica) and adhesive tape to transfer (Leica,
#39475214). After sectioning, slides were stored at -20 °C.
Retinal sections were brought to room temperature and air-dried fully, then fixed in 4% paraformaldehyde in 1x PBS for 1 minute. Sections were washed in 1x PBS three times, five minutes each, before incubating in a humidified chamber for one hour in blocking solution (20% normal goat serum, 0.5% Triton x-100 in 1x PBS). Tissue was then incubated for 20 minutes at room temperature in EdU labeling solution – the following solutions were mixed to prepare 1 mL: 760 μL 1x PBS, pH 7.4; 100 μL 1 M Tris, pH 8.5; 40
μL 100 mM copper (III) sulfate; 100 μL 0.5 M L-ascorbate; 2.4 μL 488-labeled azide (Click-
It EdU Imaging Kit, Invitrogen). Samples were washed three times, at least 5 minutes each, in 1x PBS and incubated in DAPI solution for 10 minutes (Thermo Scientfic, 1 mg/mL; diluted 1:1000 in 1x PBS). The slides were washed briefly in 1x PBS before mounting with
Fluoromount-G Mounting Medium (SouthernBiotech).
Immunohistochemistry – Zpr1
Retinal sections were obtained, stored, and fixed as described above. The fixed retinal
sections were washed three times, five minutes each, in 1x PBS and incubated in blocking
solution (5% normal goat serum, 0.1% Tween-20, 0.1% DMSO in 1x PBS) for one hour at
room temperature in a humidified chamber. Sections were incubated in Zpr1 primary
antibody (Table 4), diluted in blocking solution, overnight at 4 °C in a humidified chamber.
Slides were then washed three times, five minutes each, at room temperature in 1x PBSTD
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(0.1% Tween-20, 0.1% DMSO in 1x PBS). Secondary antibody (goat anti-mouse, AF488;
Table 4) was diluted in block and incubated on sections for one hour at room temperature
in a humidified chamber. Slides were washed three times, five minutes each, in 1x PBSTD,
counterstained with DAPI, and mounted as described above.
Immunohistochemistry – Alcama
Retinal sections were obtained, stored, and fixed as described above. The fixed retinal sections were washed three times, 10 minutes each, in 1x PBS and incubated in blocking solution (20% normal goat serum, 0.5% Tween-20 in 1x PBS) for one hour at room
temperature in a humidified chamber. Primary antibody (Alcama, Table 4) was diluted in diluting solution (1% normal goat serum, 0.5% Tween-20 in 1x PBS) and incubated on sections overnight at 4 °C in a humidified chamber. Slides were washed three times, 10 minutes each, in washing solution (1% normal goat serum in 1x PBS) and incubated in secondary antibody (goat anti-rabbit, AF488; Table), diluted in diluting solution, for one hour at room temperature in a humidified chamber. Sections were then washed three times, 10 minutes each, in washing solution, counterstained with DAPI, and mounted as described above.
Microscopy
Immunofluorescence imaging was carried out with a Leica TCS SP8 confocal scanning microscope (Leica Microsystems Europe). The 405 nm, 488 nm, and 552 nm lasers were
used with HyD and PMT detectors, and HC PL APO CS2 40x/1.30 oil objective to obtain
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three-dimensional z-stacks, imaged through the entire thickness of retinal sections and
processed with the 3D Viewer in the Leica Application Suite X Software.
Quantification of EdU lineage tracing assay
EdU-positive cells were counted manually from images of retinal section z-stacks (10 μm)
for each of five retinal layer groupings: NFL/GCL, IPL, INL, ONL/OPL, OSZ/RPE. If a cell
spanned two layers, it was counted in the layer that contained the majority of the cell.
This was repeated for every section encompassing an entire lesion. For each layer, EdU-
positive cell counts were averaged across all sections (mean of 16 sections/lesion) for a
single lesion to get one ‘n’ value. Thus, every time point and treatment condition (Alcama
or control morpholino) encompasses measurements taken from an ‘n’ of 14-18 lesions,
obtained from at least 6 different fish. ONL/INL ratios were obtained by dividing the
average ONL cell count obtained for each lesion by the average INL cell count for the same
lesion; this was repeated for each lesion as an ONL/INL ‘n’ value and plotted by time point.
Total EdU-positive cell counts were obtained by dividing all EdU-positive cells counted in
all layers of a lesion by the total number of sections (i.e. images) quantified for that lesion.
These data were plotted by time point and treatment group.
GraphPad Prism 8 was used for statistical analyses and associated figures. Multiple
t-tests with Holm-Sidak correction (without assuming equal standard deviations) were
used to determine statistically significant differences between the two treatment groups
at each time point. The alpha level was set as 0.05, and a statistically significant adjusted p-value was therefore defined as p < 0.05.
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SLO imaging and quantification
Anesthetized fish were placed in a custom rubber holder and fitted with a polymethylacrylate (PMMA) contact lens (Ø = 5.2 mm, r = 2.70 mm, center thickness = 0.4 mm; Cantor & Nissell, Ltd., Northamptonshire, UK) using Systane Ultra hydrating tears
(Alcon Laboratories, Fort Worth, TX). IR-dark field cSLO images were obtained with an
HRA2 system (Heidelberg Engineering, Carlsbad, CA) equipped with wide-field objective
(≥50° field of view). SLO images were obtained in this way at baseline immediately after injury; fish were then treated with Alcama or control morpholino within 30 minutes of injury and imaged every other day for 14 days.
Lesion areas were quantified with Image J software. Lesions were outlined with the freehand tool and area quantities obtained using the measurement function (Fig. 18).
The line drawing tool was used to draw a straight line between the centers of the temporal lesion (TL) and nasal lesion (NL), left and right of the central lesion (CL), respectively. The ends of the line were aligned with the labels of each lesion, which automatically appear at the centroid of the drawn lesion areas. This drawn measurement,
TL-NL, was used to scale the time points back to the day 0 image for a given eye: the TL-
NL measurement for time point ‘x’ was divided by the day 0 TL-NL measurement for the same eye to get the TL-NL ratio. This ratio was squared to translate a linear measurement to the two-dimensional measurement of area to get the scaling factor. This scaling factor was multiplied by the raw area measurement to get the final scaled lesion area measurements used for statistical analysis. Scaling in this way is necessary to take into account any variance in the zoom between pictures (Fig. 18).
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A total of 23 lesions in 8 fish for the Alcama group and 33 lesions in 11 fish were
quantified and scaled as described, analyzed blindly by three different individuals.
GraphPad Prism 8 was used for statistical analyses and associated figures. Multiple t-tests
with Holm-Sidak correction (without assuming equal standard deviations) were used to
determine statistically significant differences between the two treatment groups at each time point. The alpha level was set as 0.05, and a statistically significant adjusted p-value was therefore defined as p < 0.05.
Zpr1 immunofluorescence quantification
Images of Zpr1-stained sections were used to quantify relative cone density using Image
J software. Integrated density (ID) and area (A) measurements were obtained for both lesioned areas and non-lesioned areas; background mean fluorescence measurements
were also obtained above and below the stained region of interest and an average
calculated to obtain an average background fluorescence (ABF) measurement for each
image (Fig. 19). As a readout for cone density within and outside a lesion, corrected total
fluorescence (CTF) was calculated for both areas, subtracting background fluorescence:
lesion CTF = IDlesion – (Alesion × ABF); non-lesion CTF = IDnon-lesion – (Anon-lesion × ABF). The CTF
ratio was then calculated for an individual image: CTF ratio = Lesion CTF ÷ Non-Lesion CTF.
These measurements were averaged for sections spanning an entire lesion, about 10-12
sections per lesion, to obtain an individual data point indicating the proportion of cones
that were regenerated relative to the baseline cone density in the region of interest (Fig.
19).
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A total of 14 lesions in 6 fish for the control morpholino-treated group and 15 lesions in 6 fish for the Alcama morpholino-treated group were analyzed in this manner.
GraphPad Prism 8 was used for statistical analyses and associated figures. An independent t-test (two-tailed, equal variance) was used to compare morpholino treatment groups. The alpha level was set as 0.05, and a statistically significant adjusted p-value was therefore defined as p < 0.05.
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A
C B
E D
Figure 17. Targeted retinal injury using OCT-guided laser photocoagulation model. A) Setup for optical coherence tomography (OCT) guided laser photocoagulation. Ap, adjustable aperture; DM, dichroic mirror; fc, collection; fcoll, collimating; foph, ophthalmic; fr, relay; fs, scanning lenses; Gx and Gy, two axis scanning mirrors. B) Anesthetized fish in holder prior to OCT-guided laser ablation. C) Above view with arrow pointing to contact lens, which keeps the eye wet and allows for appropriate light refraction. D) OCT image of laser lesions 1 day post lesion. Arrowhead points to center of three lesions, where the photoreceptor layer has been ablated. E) Confocal scanning laser ophthalmoscopy (cSLO) images of three laser lesions made superior to the optic nerve, 1 day post-lesion. Adapted from DiCicco, et al., 2014.131
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0d 14d
Scaling lesion areas: • Day 0 TL–NL / Day # TL–NL = TL-NL Ratio • (TL-NL Ratio)2 = Scaling factor • Raw area measurement * scaling factor = scaled lesion area
Figure 18. SLO imaging analysis quantification strategy. Scanning laser ophthalmoscopy (SLO) images were used to track lesion areas in the same fish over time. Example images are shown for baseline (0d) and any other day # (14d) to demonstrate the measurement of lesion areas and calculation of scaled lesion areas. Lesion traces (yellow) were made around the outermost edge of a lesion, discernible as different from the surrounding area. Temporal lesion to nasal lesion lengths (TL-NL; cyan) were measured and used to calculate the scaling factor to obtain more accurate scaled lesion areas. See text for full details.
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ROI raw measurements:
• Lesion • Integrated density (IDlesion) • Area (Alesion) • Non-lesion • Integrated density (IDnon-lesion) • Area (Anon-lesion) • Background 1 (superior to ONL) • Mean fluorescence • Background 2 (inferior to ONL) • Mean fluorescence
Calculated measurements
• ABF = (Background 1 mean fluorescence + Background 2 mean fluorescence) ÷ 2
• Lesion corrected total fluorescence = IDlesion – (Alesion × ABF)
• Non-lesion corrected total fluorescence = IDnon-lesion – (Anon-lesion × ABF)
• CTF Ratio = lesion CTF ÷ non-lesion CTF
Figure 19. Zpr1 (cone density) quantification strategy. Example image of section stained with Zpr1 antibody (green, labeling cone photoreceptor cells) is shown to demonstrate quantitative approach taken with each section. Lesioned (yellow) and non-lesioned (cyan) areas were drawn to obtain integrated density (ID) and area (A) measurements using Image J software; background measurements were also taken for each image (magenta). Calculations shown for average background fluorescence (ABF), corrected total fluorescence (CTF), and the ratio of CTF measurements in lesioned vs. non-lesioned areas to obtain the proportion of cones regenerated relative to the baseline cone density for each section/image. See text for full details.
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Results
Zebrafish Müller glia upregulate Alcama expression in response to injury
We first wanted to characterize the expression pattern of Alcama before and after injury.
In whole retina tissue, Alcama is expressed at baseline in uninjured retinas and after injury
out to 14 days post lesion (dpl) as we have seen so far with RT-PCR (Fig. 20A). It has been
previously shown that Müller glia specifically express Alcama after injury.123 Upregulation of Alcama in Müller glia in response to injury is likely masked by the baseline expression in other cells such as in the ciliary marginal zone (CMZ) or contaminating RPE in whole retina tissue. We therefore FAC-sorted Müller glia from Tg(apoe:gfp) retinas as described in Chapter 2 and extracted RNA to generate cDNA for RT-PCR. Here, we see that Müller glia do not express Alcama in the uninjured state, but upregulate its expression after injury, as early as 1 dpl and out to 14 dpl (Fig. 20B).
We then sought to demonstrate changes in Alcama expression at the protein level to show Müller glia-specific Alcama expression after injury in a more comprehensive way, as previously published results only showed a few Alcama-positive Müller glia at a single injured time point.123 When trying to replicate these results in our hands, we saw a
striated pattern of Alcama expression spanning the outer nuclear layer, highly
reminiscent of Müller glia in the light-damaged eye at 4 dpl that was absent in the
uninjured tissue (Fig. 20C). This required exhaustive attempts with various antibodies and
staining protocols, however, and we were unable to repeat the result at different time
points or with a Müller glia-specific marker to corroborate with the Müller glia
morphology initially observed with the single label. It is important to note that the
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availability of zebrafish antibodies is quite limited, with three available for Alcama to our
knowledge, all of which we have tried here (Table 4). Two (Zn8, Zn5) come from the same
manufacturer (Zebrafish International Resource Center [ZIRC]) and are thought to actually
be the same antibody. Our previous experience with this manufacturer’s antibodies
suggests different batches of the same antibody can produce very different results, which
we believe may have been a contributing factor to our inability to repeat the published
results with their reported antibody, Zn5, nor Zn8. Our result (Fig. 20C) was achieved with a third antibody from GeneTex. This antibody, however, has only been validated in whole mount embryos and may, in part, explain our limited success with it.
We also wanted to compare Alcama expression in a mammalian system and therefore stained for ALCAM in mouse retinas (Fig. 21). Here, ALCAM expression is restricted to the choroid, inner plexiform layer (IPL), and ganglion cell layer (GCL) both at baseline and at 4 days post injury. It is unknown whether this IPL/GCL staining is from
Müller cell processes/end feet, although literature suggests that it is likely to be from
amacrine and ganglion cell processes.235 Importantly, there is no upregulation of
expression in the outer nuclear layer of mouse retinas that is seen in zebrafish retinas
after injury (Fig. 20). This observation, in combination with the Müller glia-specific
upregulation of Alcama in response to injury, suggests a regenerative role for Alcama in
zebrafish retinas.
Morpholino-mediated knockdown of Alcama
To determine whether Alcama plays a role in regeneration, we first needed to knockdown its expression and see how the process of regeneration changes in response. Because of
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Alcama’s prominent role in various aspects of development, we needed to alter its
expression in the adult fish only and target it to the retina. The tools available to accomplish this in zebrafish, however, are somewhat limited. Genetic editing through
TALENS or, more recently, CRISPR/Cas9 systems, are still relatively new technologies in zebrafish. They are quite time consuming strategies and were out of the scope of this project timeline. There are a few newly-generated Alcama mutant zebrafish lines, but they have only just been published and have yet to be characterized as viable adults.237,238
We therefore used a morpholino-based strategy to knock down Alcama expression as
previously described.236
Morpholinos are synthetic, anti-sense oligonucleotides designed, in our case, to
target and bind Alcama pre-mRNA in a sequence-specific manner at a splice site (Fig. 22A).
This disrupts splicing to introduce a premature stop codon, triggering nonsense-mediated
decay and ultimately leads to degradation of Alcama mRNA. We also utilized a scrambled
control morpholino, not targeting any sequence in the zebrafish genome, for all
knockdown experiments to demonstrate Alcama-specific effects. Both morpholinos are
tagged with lissamine, a charged, fluorescent molecule that allows efficient delivery into
the retina via electroporation and subsequent visualization. By retinal flatmount, we can
see that lissamine signal is well-distributed across the retina; histological sections also
demonstrate that the morpholino is delivered to all retinal layers (Fig. 22B). Taken
together, we can conclude that the lissamine-tagged morpholino is efficiently delivered
to the majority of the adult fish retina after electroporation.
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To confirm that the morpholino was effectively knocking down Alcama expression as expected, we used a FACS-based approach to separate out lissamine-positive and lissamine-negative cells. This was done in order to look specifically at cells that had taken up the morpholino (lissamine-positive) and determine a morpholino-specific effect that could be masked by cells that had not taken up the morpholino when looking at whole retina tissue, especially in cases of less efficient delivery. Furthermore, this would demonstrate whether the morpholino was effectively inhibiting Alcama expression in
Müller glia. For this experiment, retinal cell suspensions were generated from
Tg(apoe:gfp) retinas two days following retinal injury and Alcama morpholino treatment, then subjected to flow cytometric cell sorting as described in Chapter 2.
Following the GFP gating for Müller glia, an additional gating parameter was applied to collect a total of six cell populations based on GFP and lissamine signal.
Subsequent RT-PCR analysis for each of these populations demonstrated that Alcama was not expressed in any of the lissamine-positive cell populations, regardless of whether the lissamine signal was high or low (Fig. 23A). This indicated that the Alcama morpholino was indeed inhibiting Alcama expression not only in Müller glia (GFP-high), but in other cell types as well (GFP-negative, GFP-low). An additional experiment was similarly performed, but with control morpholino treatment. Here, gating was performed based only on lissamine signal, where lissamine-positive and lissamine-negative cell populations were collected. RT-PCR analysis on these populations indicated that the control morpholino was not having an effect on Alcama expression, as it was detectable in both populations
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of cells (Fig. 23B). No changes in Alcama expression with either control or Alcama morpholino treatment were detectable at the whole retina level (Fig. 23C).
Alcama inhibition affects progenitor cell migration during retinal regeneration
In order to test our main hypothesis that Alcama helps to facilitate progenitor cell migration, we used an EdU lineage tracing assay to label and track progenitor cells during regeneration. For this experiment, we began by administering three laser lesions to each fish retina and immediately treated them with either control or anti-Alcama morpholino.
At three days post lesion (dpl), all fish were IP injected with EdU to label progenitor cells when their proliferation is most robust. Eyes were collected at 3, 4, 7, and 14 dpl, sectioned, and stained for EdU (Fig. 24). For both control and anti-Alcama morpholino treatments, the number of EdU-positive cells in each retinal layer were quantified over time in order to determine what effect Alcama inhibition might have on the migration of progenitor cells.
Looking at the proportion of EdU-positive cells in each layer in terms of the total
number of EdU-positive cells in the lesion plotted over time, we can see that most
proliferating cells reside in the inner nuclear layer (INL) at 3 dpl for both morpholino
treatment groups (Fig. 24, Fig. 25). Over time as regeneration is occurring, the progenitors
migrate out of the INL and into the outer nuclear layer (ONL) (Fig. 25). While this trend
remains consistent between control and Alcama groups, there is a distinct separation
between the two. There are statistically significant differences between treatment groups
specifically at 4, 7, and 14 dpl in the INL, and at all time points (3, 4, 7, and 14 dpl) in the
ONL (Fig. 25C), suggesting that Alcama inhibition contributes to a delay in progenitor cell
81 migration. In this way, we also assessed the distribution of EdU-positive cells in the inner plexiform layer (IPL) and nerve fiber/ganglion cell layers (NFL/GCL). Here, the results are less dramatic, and little to no migration into or out of these layers seems to occur after injury (Fig. 25D). Looking more closely, however, there is a statistically significant difference between Alcama and control morpholino treatment groups at 3 dpl in the
NFL/GCL, and at 7 dpl in the IPL. As another way of assessing differences in distribution of progenitors, we looked at the ratio of outer-to-inner nuclear layer EdU-positive cells over time, where a higher ratio indicates more EdU-positive cells in the ONL than in the INL at a particular time point (Fig. 25E). This analysis shows a significantly lower ONL-to-INL ratio at all time points, providing further support for the observation of misguided or delayed migration of progenitor cells with Alcama morpholino treatment compared to control.
We also assessed differences in the total number of EdU-positive cells between treatment groups to determine whether Alcama inhibition had any effect on progenitor cell proliferation (Fig. 26A). The number of EdU-positive cells per section quantified for every lesion was plotted for both treatment groups at each time point. Multiple t-tests show significant differences between the two groups at 3 dpl and 7 dpl, where there are significantly more EdU-positive cells per section in the control morpholino group compared to the Alcama morpholino treatment. This indicates that Alcama inhibition may also be playing a role in the modulation of progenitor cell proliferation in addition to their migration. When looking at the number of EdU-positive cells in the retinal pigment epithelium (RPE) and outer segment zone (OSZ), we did not see any significant differences between the treatment groups (Fig. 26B). This suggests that Alcama inhibition does not
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affect the infiltration of immune cells that are likely the population of cells represented
by the EdU-positive cells in the RPE/OSZ region.
Taken together, these results demonstrate that Alcama contributes to the
migration of progenitor cells during retinal regeneration, as shown by the disruption of
EdU-positive progenitor cell movement with Alcama morpholino treatment compared to
control (Fig. 25). Alcama may also play a role in the initial and/or continued proliferation of progenitor cells, as indicated by differences in total EdU-positive cell counts between
Alcama and control morpholino treatment groups at 3 dpl and 7 dpl (Fig. 26A).
The regenerative process may be inhibited or delayed with Alcama knockdown
We then wanted to determine whether Alcama inhibition had an effect on overall
regeneration, perhaps as a result of the misguided migration and/or proliferation of
progenitor cells. Using confocal scanning laser ophthalmoscopy (SLO) imaging, we can
obtain in vivo images of fish retinas. In this way, we can visualize laser lesions and track
retinal regeneration of the same lesions in the same fish over time. By quantifying the size
of these lesions as regeneration is occurring, we can determine whether there are
changes to overall regeneration when Alcama is inhibited.
For this analysis, retinas were injured with three laser lesions and immediately imaged by SLO for a 0d post-injury time point, then treated with either control or anti-
Alcama morpholino. Retinas were imaged every two days afterward until the terminal time point of 14 dpl (Fig. 27A, Fig. 28). All lesions at every time point from both treatment groups were quantified by tracing the lesion and measuring the area with Image J. After scaling the measurements appropriately (Fig. 18), measurements were plotted in terms
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of percentage change from baseline by dividing all scaled areas by the area measurement
from the same lesion at day 0, utilizing measurements from three individual graders (Fig.
27B). Analyzing lesion area measurements in this way demonstrates a distinct separation between the two treatment groups, with statistically significant differences at days 8, 10, and 12. Though not significant during the earlier time points, lesions in Alcama morpholino-treated retinas are consistently larger than those treated with control morpholino throughout the time-course. The separation between the two groups begins to increase over the next several days, with the largest difference in lesion sizes occurring around 12 dpl (Fig. 27B). By 14 dpl, however, there is no significant difference between
the two treatment groups, although the p value of 0.052243 is very close to the alpha
level of 0.05. This suggests that regeneration in the Alcama morpholino-treated retinas
was stalled or delayed but was able to catch up, to some extent, with the control group’s
rate of regeneration. Additional time points in future experiments may help clarify how
and why this is occurring.
Alcama inhibition decreases cone regeneration
The eyes from the SLO analysis were collected at their terminal time point of 14 dpl and
used for histological analysis as a measure of regeneration on a cellular level (Fig. 27A).
Sections were stained with Zpr1 antibody to label cone photoreceptor cells to determine
what effect Alcama inhibition might be having on regeneration of cones specifically (Fig.
29A). The cone density within and outside the lesioned area was measured and corrected
total fluorescence (CTF) was calculated for each region (Fig. 19). Final results are plotted
as CTF ratios of lesion/non-lesion CTF measurements to take differences between
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baseline cone densities into account (Fig. 29B). Overall, lesions treated with the Alcama
morpholino show a significantly lower CTF ratio, indicating that less cones have
regenerated at 14 dpl compared to those treated with the control morpholino. This
suggests that Alcama inhibition contributes to a reduction in cone regeneration, which
could be the direct or indirect result of disrupted migration, proliferation, or
differentiation of progenitor cells. Future studies will be needed to determine the
mechanisms underlying this result.
Because the 14 dpl results demonstrate no difference in lesion size as measured
by SLO, but there is a significant difference in cone density for the same lesions at the
same time point, we performed a comparative analysis of these results to determine if
there was a relationship between the measurements (Fig. 30). Linear regression indicates
a slight negative correlation between lesion area and cone density, but the results are not
significant, suggesting that these two measurements are indicative of different aspects of
regeneration (Fig. 30B). This is further demonstrated when comparing the overall change
in lesion size at 14 dpl and CTF ratio, which shows virtually no correlation between the
two measurements (Fig. 30C). These discrepancies are likely because the area of the
lesion as measured by SLO is actually reflective of multiple facets of regeneration,
including inflammation, edema, overall cellular organization, autofluorescence, and other
imaging artifacts. This is demonstrated, for example, by the apparent slight increase in
lesion size at 2 dpl compared to baseline (Fig. 27B). The lesion itself is likely not increasing in size, rather inflammation or autofluorescence becomes detectable by SLO at this time and obscures the lesion itself, causing the boundary lines to be drawn further out and the
85 lesion appears to be larger. Conversely, Zpr1 staining represents a more specific measurement (i.e. cone density), as indicated by corrected total fluorescence. These results underscore the importance of taking all analyses into account: while SLO is helpful to look at, particularly for tracking how lesions change over time, it is hard to pinpoint exactly what is changing. On the other hand, immunofluorescence is more limiting experimentally but does show changes to a specific cell type.
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A
B
C Uninjured Injured (4 dpl)
Alcama
(+DAPI) Overlay
Figure 20. Alcama expression is specifically upregulated by Müller glia in response to injury in the zebrafish retina. A) RT-PCR analysis of whole retina tissue, either uninjured (UN) or injured and collected at various days post lesion (dpl). nT, no template negative control. B) RT-PCR analysis of Müller glia isolated via fluorescence activated cell sorting (FACS) from cell suspensions of 6-8 retinas each that were either uninjured (UN) or injured and collected at various days post lesion (dpl). nT, no template negative control. A, B) Representative of three independent experiments. C) Stained sections (10 μm) of retinas that were either uninjured or injured by light lesion and collected 4 days post lesion (dpl). Arrow indicates outer nuclear layer, where Müller glia-like processes are visible when staining for Alcama in the injured retina. Counterstained with DAPI (blue, overlay). 63x magnification; scale bars = 25 μm.
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Uninjured Injured (4 dpl)
ALCAM
PI)
Overlay (+
Figure 21. ALCAM expression in mouse retinas is restricted to the inner plexiform layer, ganglion cell layer, and choroid before and after injury. Stained sections (10 μm) of adult mouse retinas at baseline (uninjured) or 4 days after laser lesion injury (4 dpl). ALCAM expression (red) is observed in the inner plexiform layer, ganglion cell layer, and choroid. This expression pattern does not change with injury at 4 dpl, with no upregulation in the outer nuclear layer that is observed in the fish retina. Counterstained with propidium iodide (PI; blue, uninjured overlay; white, injured overlay). 40x magnification; scale bars = 50 μm.
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A
B
C
Figure 22. Morpholino delivery and mechanism of action. A) Schematic of morpholino mechanism of action. B) Schematic of morpholino delivery; adapted from Thummel et al., 2011.236 C) Histological section (left) and whole retina flat mount (right) demonstrating delivery of the lissamine-tagged morpholino to all retinal layers and across most of the retina, respectively. OS, outer segments; ONL, outer nuclear layer; INL, inner nuclear layer; GCL, ganglion cell layer; T, temporal; S, superior; N, nasal; I, inferior; ON, optic nerve.
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A
B C
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Figure 23. Validation of morpholino-mediated Alcama knockdown. A) A cell suspension was generated from retinas that were injured, treated with Alcama morpholino, and collected 2 days later and subjected to fluorescence activated cell sorting (FACS) to isolate cell populations based on GFP and lissamine signal: G(-), GFP-negative; GL, GFPlow; GH, GFPhigh; LL, lissamine low; LH, lissamine high. RT-PCR of the indicated populations shows that Alcama is inhibited in all cell populations containing the morpholino (i.e. lissamine positive); lissamine negative cells were unable to be collected due to limitations of the flow cytometry equipment. B) A cell suspension was generated from retinas that were injured, treated with control morpholino, and collected 2 days later and subjected to FACS to isolate cell populations based on lissamine signal (lissamine-negative, L(-); lissamine-positive (+)). RT-PCR of the indicated populations shows Alcama is expressed in both populations and therefore unaffected by the control morpholino. A,B) Positive control (+ Ctrl) is 3 dpl FAC-sorted Müller glia. See Chapter 2 for full details about sample preparation for flow cytometric cell sorting and subsequent gating to first identify individual, live cells before implementing the gating strategies specific to the experiments above. C) Quantitative PCR in whole retina tissue that was untreated (black), treated with control morpholino (Control MO, blue), or treated with Alcama morpholino (Alcama MO, red) demonstrates that differences in Alcama expression at the RNA level are not detectable in whole retina tissue; multiple t-tests with Holm-Sidak correction show no significant differences between any of the treatment groups at any time point.
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A
B
Figure 24. EdU lineage tracing assay outline and representative images. A) Outline of EdU lineage tracing assay. Fish are administered three laser lesions on day 0 and immediately treated with either control or anti-Alcama morpholino (within 30 minutes of injury). All fish then receive IP injections of EdU on day 3. Eyes are collected 4 hours later on day 3, or on days 4, 7, and 14. Retinas are sectioned through the entire thickness of all lesions and every section is stained for EdU. B) Representative images from this assay. Each image shows a single z-stack of a single 10 μm section. An average of 16 sections/images were used to count the EdU-positive cells per lesion quantified.
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B A
C D
E
Figure 25. Movement of EdU-positive cells within retinal lesions over time. A) Key for B-D: INL, inner nuclear layer; ONL, outer nuclear layer; IPL, inner plexiform layer; NFL/GCL, nerve fiber layer and ganglion cell layer. B) Comparison of all 4 retinal layers assessed for this assay, plotted by retinal layer as a proportion of the total EdU-positive cells quantified. For clarity, no significance marks are included, but are instead indicated on the separated graphs in B and C. C) Movement of EdU-positive cells within retinal lesions over time: INL and ONL. D) Movement of EdU-positive cells within retinal lesions over time: IPL and NFL/GCL. E) ONL/INL ratios of EdU-positive cells over time after injury. B-E) n = 14-18 lesions at each time point in both treatment groups, with 6 fish represented in all groups; dpl, days post lesion. Error bars represent standard error of the mean. Adjusted p-values obtained from multiple t-tests comparing the control and Alcama morpholino treatments with Holm-Sidak correction.
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A
B
Figure 26. Total EdU-positive cell counts in lesions for the EdU lineage tracing assay. A) Total EdU-positive cells quantified within all retinal layers, plotted in terms of number per section. B) Total EdU-positive cells quantified in the lesioned area within the outer segment zone (OSZ) and retinal pigment epithelium (RPE), plotted in terms of number per section. A, B) n = 14-18 lesions at each time point in both treatment groups, with 6 fish represented in all groups; dpl, days post lesion. Error bars represent standard error of the mean. Adjusted p-values obtained from multiple t-tests comparing the control and Alcama morpholino treatments with Holm-Sidak correction.
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A
B
Figure 27. SLO imaging analysis shows increased lesion sizes with Alcama morpholino treatment at some time points. A) Experimental outline. B) Composite results incorporating lesion measurements from three blinded individuals: three graders traced lesions independently from SLO images as described, tracked over time and used to calculate the change in lesion area as a percentage of the baseline measurement for the corresponding lesion. This was carried out by each grader for the same 23 lesions in 8 fish (Alcama morpholino-treated) and 33 lesions in 11 fish (control morpholino-treated); thus, each data point in the Alcama group represents n = 69 lesion area measurements, and in the control group n = 99 lesion area measurements. Error bars represent standard error of the mean. Multiple t-tests comparing the control and Alcama morpholino treatments were performed with Holm-Sidak correction.
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Figure 28. Representative images from SLO imaging analysis. SLO images taken according to outline in Figure 27A. Each set of images, control or Alcama morpholino-treated (0d-6d, top; 8d-14d, bottom), were taken from the same fish. All images taken in IR dark field mode.
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A
B
Figure 29. Immunohistochemistry of cone photoreceptors in control and Alcama morpholino-treated retinas at 14 days post lesion. A) Representative images from 10 μm sections through lesions in retinas treated with either control (top) or Alcama (bottom) morpholino (MO), stained for Zpr1 (cones, green) and DAPI (nuclear stain, blue) at 14 days post lesion. Scale bars = 50 μm, 40x magnification. B) Quantification of all Zpr1 staining results. Corrected total fluorescence (CTF) was calculated from the integrated densities, adjusted for background fluorescence, for regions within the lesion and outside the lesion. CTF ratio = lesion CTF/non-lesion CTF. One data point represents the average CTF ratio across all sections spanning a single lesion. n = 14 lesions in 6 fish for control MO treatment; n = 15 lesions for Alcama MO treatment. Unpaired t-test (equal variance, two-tailed) comparing control and Alcama MO treatment groups indicates significantly lower CTF ratios (less dense cone staining in lesions) in the Alcama MO group. Error bars represent standard error of the mean.
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A
B C
Figure 30. Comparison of in vivo imaging and immunohistochemistry as methods for assessing retinal regeneration. A) Representative comparison of in vivo SLO images (top) and terminal 14 dpl (days post lesion) immunohistochemistry from the same eye (bottom). TL, temporal lesion; CL, central lesion; NL, nasal lesion; ON, optic nerve. B) Linear regression of scaled lesion areas measured from in vivo SLO images and cone densities (corrected total fluorescence, CTF measurements) of corresponding lesions at the terminal time point of 14 dpl; au, arbitrary units. C) Linear regression of the total change in lesion area (percentage of baseline measurement at 14 dpl), measured from in vivo SLO images, and lesion/non-lesion CTF ratio of zpr1 staining from the corresponding lesion at 14 dpl. B, C) Blue data points represent control morpholino (MO) treated lesions; red data points Alcama MO treated lesions; n = 11 lesions in 5 fish for both control and Alcama MO treatment groups.
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Discussion
Understanding the mechanisms of retinal regeneration inherent to zebrafish is an
essential component in developing regenerative therapies for human use. Müller glia have regenerative potential in both fish and mammals: while they proliferate and hypertrophy in response to damage in mammals, zebrafish Müller glia comprise a functional stem cell niche that gives rise to new neurons and reconstitutes the retina.36,113,165 In response to injury, Müller glia in zebrafish dedifferentiate and undergo
a single, asymmetric division giving rise to a Müller glia and a progenitor cell, which
proliferates.36,111 These initial parts of the regenerative process have been the most
extensively studied in the field, regarded as the defining aspects between a gliotic scarring
response in mammals but productive regeneration in fish.114,124 The resulting progenitor
cells then migrate and appropriately differentiate to replace the cells lost to injury. The
mechanisms underlying these aspects of retinal regeneration, though, remain largely
unclear, and much of what we know about cellular migration in general has been gleaned
from similar processes in other organisms, tissue types, or developmental contexts. In this
way, we identified Alcama as an adhesion molecule that could be potentially involved in the specific context of zebrafish retinal regeneration.123,223,226,235 Here, we have executed
a functional analysis of Alcama during adult zebrafish retinal regeneration, the first of its
kind to our knowledge. We have shown that when Alcama is inhibited, there is a
significant reduction in the efficiency of progenitor cell migration and overall regenerative
capacity of the zebrafish retina.
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Alcama expression in the zebrafish retina
First we identified that Müller glia specifically upregulate expression of Alcama in response to injury, expanding upon the previous observation of Alcama expression only at a single time point post-injury in the adult zebrafish retina, formerly the only known study involving Alcama in this context.123 We have seen Alcama expression at the RNA level in both whole retina and FAC-sorted Müller glia, but Müller glia only express Alcama after injury, as early as 1 day post lesion (dpl) and out to 14 dpl to some extent (Fig. 20A-
B). We have also seen a Müller glia-like pattern in injured retinal sections stained for
Alcama, with Alcama-positive processes spanning the outer nuclear layer (Fig. 20C), which is not observed in mouse retinas (Fig. 21). Because of difficulty with immunofluorescent techniques and the generally limited availability of antibodies for zebrafish epitopes, we have unfortunately been unable to expand these observations at the protein level beyond a single time point of 4 dpl. Consequently, it would be highly beneficial to create a custom antibody for zebrafish Alcama to be used in future immunohistochemistry, Western blotting, immunoprecipitations, and other antibody-based assays. This could also be applicable to flow cytometric applications by conjugating a fluorophore to a custom
Alcama antibody with commercially available kits. Flow cytometry may be a particularly useful tool for elucidating Alcama’s mechanism of action, as it could be used as a quantitative assessment of Alcama expression at the protein level in response to various experimental conditions.
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Morpholino-mediated knockdown of Alcama
To determine whether Alcama is playing a role in retinal regeneration, particularly in the
migration of progenitor cells, we wanted to identify the functional consequences of
knocking down its expression during regeneration. Since Alcama has been shown to play
important roles during development both in the retina and elsewhere, we needed a way
to inhibit expression in the adult fish only and preferably target it to the retina. The ideal
solution would have been to develop an inducible, Müller glia-specific Alcama knockout
fish, a fairly straightforward task and readily available option in mice. Because this technology is relatively new in zebrafish and not yet fully developed, the timeline and scope of this project did not allow us to pursue the development of such a tool. We therefore chose to utilize a readily available method that has been previously demonstrated to induce gene-specific knockdown targeted to the adult zebrafish retina involving an antisense morpholino targeting Alcama for degradation.236 Intraocular
injection with subsequent electroporation allows delivery specifically to the retina, made
more efficient by the charged lissamine tag conjugated to the morpholino, which also
allows fluorescent visualization to confirm delivery to all retinal layers across most of the
retina (Fig. 22). We acknowledge that morpholino-mediated knockdown is not necessarily
the ideal method to inhibit expression, but it was previously the best strategy available to us. Morpholinos often carry a bad reputation for their potential off-target effects, especially for their use in zebrafish; however, this likely stems from their extensive use in developmental studies involving morpholino treatment of embryos.239 This is far more
likely to result in unintended consequences because of the significant development that
101 has yet to occur under the influence of the morpholino. Conversely, inhibition of a particular gene targeted to a specific tissue in an adult fish poses much less risk for off- target effects.
We have confirmed morpholino-mediated Alcama knockdown with a flow cytometric cell sorting-based strategy to show that Alcama is not expressed specifically in cells that have taken up the morpholino (Fig. 23A). We used a similar strategy to demonstrate that the control morpholino does not impact Alcama expression (Fig. 23B), and therefore proceeded with this strategy for functional assays to understand the role of Alcama in regeneration. To date, however, we have not been able to repeat these results. A previous repetition of the FAC-sort following Alcama morpholino treatment gave conflicting results to the originally successful demonstration (data not shown); qPCR of whole retina tissue showed no significant differences between untreated, control morpholino-treated, and Alcama morpholino-treated retinas (Fig. 23C), and no splicing changes were detectable in the same whole retina samples on agarose gel (data not shown). These failed validations, however, were performed with morpholino treatments executed by a different person from those used in the validation showing Alcama knockdown (Fig. 23A-B). Importantly, the same person executed the morpholino treatments in the successful validation and all functional assays. The inherent problem in validating this morpholino technique is that there is no way straightforward way to definitively show that the morpholino is both getting in and effectively knocking down expression within the same retina. We attempted to circumvent this problem with the flow cytometric cell sorting, but we have had difficulty in confirming this knockdown, and
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plan to repeat the cell sorting experiments. Not only can there be variability between treatments, but we have also seen differences in validation results when the morpholino treatments are performed by different individuals, which is important for experimental considerations.
In addition to repeating the cell sorting morpholino validation, we also plan to utilize other methods to measure Alcama expression after morpholino treatment. For example, the effects of some morpholinos can only be seen at the protein level; while our morpholino is predicted to act on Alcama pre-mRNA to ultimately result in its degradation, it is possible the actual in vivo effect is not reliably detectable by the PCR- based methods we have so far employed. We will therefore use Western blotting to look for differences in whole retina tissue either left untreated or treated with the control or
Alcama morpholino to assess differences in the resulting Alcama protein, both in quantity and any changes to molecular weight. The caveat here is whether the Alcama antibodies we have had limited success with in immunohistochemistry applications will be effective in a Western blotting application.
Another option to explore is an in vitro validation of the morpholino. Here, a
Müller glia cell line could be used to determine whether the morpholino is enacting its effect. It would first be necessary to check that the cells express Alcama at baseline and that the sequence of the morpholino effectively targets the Alcama that is expressed by the cells, as most available cell lines are mammalian. Ideally, the cell culture method described in Appendix 1 could be utilized for this purpose, as it is derived from zebrafish, but confirmation of Alcama expression in vitro would still need to be performed. Likewise,
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any cell line could be transfected with an expression construct containing zebrafish
Alcama, which may even be designed with a tag to more easily detect differences at the
protein level.
Alternatively, we can use an in situ approach such as RNAScope to simultaneously label Alcama RNA and visualize the morpholino itself, either by visualizing the lissamine signal or probing for the morpholino sequence itself. If executed in Tg(apoe:gfp) retinas, we can potentially also see how much of the effect is occurring in Müller glia specifically
(after injury). In terms of visualizing lissamine and/or GFP signal, we would first have to overcome any quenching effects that may occur during fixation or other parts of the hybridization protocol. Overall, the innate variability of the in vivo electroporation of morpholinos technique, both between individuals and between treatments performed in the same hands, highlights the importance of developing a genetic model for a more consistent Alcama knockout to further study its role in retinal regeneration.
The functional role of Alcama in retinal regeneration
Using morpholino-mediated Alcama knockdown, we have shown that the migration of progenitor cells is consequently disrupted. Although the general pattern of movement from the inner to outer nuclear layers is unaltered, the efficiency of the migration is significantly reduced (Fig. 10, Fig. 24, Fig. 25). This indicates that while Alcama may play a
role in this process, there are likely other molecules acting similarly to facilitate the
movement of cells during regeneration, either in a concurrent or compensatory manner.
In addition, we showed that with Alcama inhibition, there is a reduction in the number of
total proliferating cells at 3 dpl (Fig. 26A), suggesting that Alcama-mediated interactions
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also play a role in signaling progenitor cells to proliferate, and therefore less cells
incorporate EdU. Because this discrepancy between Alcama and control morpholino treatments is not observed at 4 dpl, it suggests that the initial proliferation was delayed
but able to catch up in a relatively short period of time. We again see less EdU-positive
cells in the Alcama group relative to control at 7 dpl (Fig. 26A), conveying the possibility
that there are two distinct proliferative phases for progenitors during retinal
regeneration, and Alcama signaling may play a role in both. Similarly, by 14 dpl, the total
EdU-positive cell numbers in the Alcama morpholino-treated group return to levels
comparable to the control group, again suggestive of compensatory mechanisms at work.
Note that the number of EdU-positive cells appears to decrease between days 4 and 7;
this likely represents some cells proliferating at an increased rate to compensate for the
delay seen at 3 dpl, to such an extent that the EdU within their DNA has been diluted too
much to be detectable. A certain population of cells, then, may not have proliferated in
this earlier phase, but began to proliferate in the later phase, therefore spreading EdU-
incorporated DNA to their progeny at detectable levels, resulting in an increased count of
EdU-positive cells between 7dpl and 14 dpl (Fig. 26A).
To determine whether Alcama inhibition plays a role in overall regeneration,
perhaps as a result of the disruption to progenitor cell migration, we used in vivo SLO
imaging and immunohistochemistry analysis on terminal eyes to obtain two different
readouts of regeneration. By tracking lesion sizes in the same fish over time with SLO, we
have shown that there is less of a change from baseline measurements with Alcama
morpholino treatment compared to control (Fig. 27). The measurements of lesion areas
105 from SLO imaging were performed by three different people, due to the relatively subjective nature of determining the boundaries of lesions from SLO images. Differences in coloration between images make it unreliable to utilize more automated features of
Image J or other imaging software to define a lesion. Because the correlations between features identified by SLO and biological interpretation are largely unknown, results should be cautiously interpreted, especially in the context of a morpholino-mediated knockdown, which may itself may be incomplete.
Using the terminal 14 dpl eyes from the SLO analysis, we also stained retinal sections for Zpr1 to get a less comprehensive but more precise measurement of regeneration in terms of cone cell density (Fig. 29). These results indicated that Alcama inhibition resulted in a decreased density of regenerated cones compared to control.
Because this data was only obtained for a single, terminal time point, we cannot be sure of when this reduced regeneration of cones begins to occur, or how long it persists.
Furthermore, we saw through comparative analyses of SLO and Zpr1 immunofluorescence data that there is little to no correlation between the two (Fig. 30).
This underscores the importance of considering both sets of data as separate pieces of information that indicate different aspects of the regenerative process, but are not directly comparable.
In the future, we plan to perform experiments using cone-labeled fish,
Tg(TαC:gfp), in order to combine the benefits of in vivo imaging with the precision of immunohistochemistry. For example, imaging the cone-labeled fish with SLO over time, after injury and Alcama or control morpholino treatment, would give a snapshot of the
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density of cone cells in the same lesions over time. SLO images of lesioned Tg(TαC:gfp)
fish are much easier to interpret, as the fluorescence given off is specifically from GFP
signal when the appropriate filters are applied (Fig. 38, Appendix 2). These cone-labeled
fish are advantageous not only for providing a more distinct, objective picture of defining lesion boundaries to measure changing area over time, but also offer the opportunity to measure the density of cones regenerating after laser ablation in vivo over time. In the most ideal experiment, this SLO imaging analysis would be performed in fish generated from crosses of Tg(TαC:gfp) and one of the Alcama mutant fish mentioned below to give the clearest readout of how regeneration is changing in response to Alcama dysfunction.
To corroborate the morpholino results, we intend to repeat these studies with one or more genetic models of Alcama knockdown. Very recently, there have been a few
Alcama mutant fish lines made publicly known: alcamasa34926, containing a splice site point
mutation; and alcamasa12510, containing a nonsense mutation. These mutant embryos are now publicly available via the Zebrafish International Resource Center (ZIRC), but it is unknown whether they are viable.237 Similarly, Alcama mutant bns201, containing an 8
bp deletion in exon 10, and bns244, a homozygous mutant lacking the Alcama promoter,
have been reported in a recent publication.238 Because these fish were generated for
developmental studies in embryos, it is not known whether they are viable as adult fish.
Notably, the bns201 and bns244 mutants displayed a compensatory upregulation of
Alcama’s paralogue, Alcamb, but it is unknown whether this would occur in the context of adult fish retinal regeneration, or if it contributes to the regenerative process in a similar way, if at all. We are in the process of obtaining some of these Alcama mutants to
107 determine the viability of heterozygous and homozygous mutations. Ideally, both are viable and can be used for the EdU lineage tracing, SLO, and Zpr1 staining assays to compare with the results from the morpholino-mediated knockdown. In all of these experiments, we would expect more dramatic differences between wild-type and the genetic mutants, especially the homozygous, compared to those between the control and
Alcama morpholino treatments since the latter is not a complete knockdown, nor is it known how long it lasts. An even more informative model would be a Müller glia-specific inducible knockout of Alcama to determine whether it is the Alcama expressed by Müller glia after injury specifically that plays a role in progenitor cell migration and overall regeneration. Such a model is not available to our knowledge and would require us to generate and characterize our own line, a multi-year project in and of itself that may be pursued concurrently with the repetition of the functional assays with the available genetic models (if viable). If the available mutants are not viable, this indicates the necessity of Alcama function during development and an inducible model, whether
Müller glia-specific or globally, may be the only option for an in vivo, genetic comparison to the morpholino results.
Overall, our results thus far indicate that Alcama is playing a role in zebrafish retinal regeneration, at least in part by facilitating the migration of progenitor cells.
Repeating the experiments described herein with a genetic model of Alcama inhibition can confirm these results, likely with more robust outcomes. Future mechanistic studies will help to further elucidate how Alcama is functioning within specific cells or pathways
(Chapter 4).
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CHAPTER 4 CONCLUSIONS AND FUTURE DIRECTIONS
Conclusions
Humans and other mammals can experience vision loss from a multitude of diseases with highly varied pathophysiologies; the lost or damaged cells in all cases cannot be replaced.
By understanding how retinas in animals such as zebrafish inherently regenerate, we may someday be able to translate that knowledge into therapies for human use. In this work, we used zebrafish as a model organism to develop a method for isolating Müller glia and to demonstrate the functional importance of Alcama, both of which contribute to an improved understanding of the regenerative process.
First, we developed a protocol for the preparation of a single cell suspension from zebrafish retinal tissue for subsequent flow cytometric cell sorting of Müller glia (Chapter
2). To our knowledge, there were no previously published studies which described the
dissection, dissociation, and sorting process in sufficient detail to execute the isolation of
zebrafish Müller glia in a reliable and repeatable manner. We reported the key steps of a
clean cell suspension preparation and demonstrated that Müller glia can be collected in
the GFP-high population of cells from Tg(apoe:gfp) fish retinas. The ability to effectively
isolate Müller glia creates the opportunity for downstream applications that can offer
novel insights into the cells that lie at the heart of the regenerative process in zebrafish.
We used this technique, for example, to demonstrate that Müller glia upregulate
expression of Alcama, which was not otherwise detectable at the whole retina level
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(Chapter 3). In addition, the preparation of a cell suspension can not only be applied to
other types of flow cytometry, but can also be used for developing primary cell culture
systems, detailed in Appendix 1. Overall, we have developed an effective method for
isolating zebrafish Müller glia, which serves as an important tool for future use in the field
of retina regeneration research.
We then explored the functional role of the adhesion molecule Alcama in zebrafish retinal regeneration, with the hypothesis that it facilitates the migration of progenitor cells and is therefore important for overall regeneration (Chapter 3). After
demonstrating that Alcama expression was upregulated specifically by Müller glia in
response to injury, we inhibited Alcama expression with morpholino delivery into the
adult fish retina for subsequent functional assays. Alcama inhibition was shown to delay,
but not prevent, the migration of progenitor cells during regeneration using an EdU
lineage tracing assay. In addition, SLO imaging and immunohistochemistry showed an
inhibitory effect on overall regeneration with Alcama morpholino treatments. While we
acknowledge the limitations and caveats of morpholino-mediated knockdown, this work
provides a solid foundation on which to base future experiments that may elucidate yet
unknown or understood mechanisms contributing to the process of retinal regeneration
in both zebrafish and mammals. By showing that Alcama inhibition leads to both a delay
in progenitor cell migration and overall regeneration, we demonstrated the importance
of appropriate progenitor cell movement in subsequent cellular regeneration. From a
therapeutic standpoint, it will be necessary to gain a more comprehensive understanding
of how retinal stem cells are guided to where they are needed and effectively integrate
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into existing retinal circuitry. Because Alcama inhibition was shown to delay or impede
progenitor cell migration and overall regeneration but not prevent it entirely, many other
molecules likely contribute to the regenerative process by also facilitating progenitor cell
movement. These may include other adhesion molecules and extracellular matrix
components. This work serves as a starting point for exploring not only the mechanisms of Alcama signaling, but also other pathways and molecules that function in similar or complementary ways.
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Future directions
The process of retinal regeneration is extremely complex, and much work remains to be done in elucidating how it occurs in animals with innate regenerative capacity, as well as in translational and clinical applications for human therapy. Here, we highlight potential
future studies both within and beyond the context of our work exploring the roles of
Alcama, extracellular matrix, and microglia in the process of retinal regeneration.
Proposed mechanistic studies involving Alcama
As our preliminary characterization has suggested that Alcama plays a role in zebrafish
retinal regeneration, particularly in the migration of progenitors, we next need to
elucidate the specific mechanisms underlying this function. Some of the first steps will
include large-scale, open-ended experiments to identify potential processes or pathways
as a starting point for more targeted mechanistic studies, including RNAsequencing and
immunoprecitation/mass spectrometry. We would likely perform RNA sequencing on
Müller glia isolated from injured retinas treated with control or Alcama morpholino,
compared to untreated, injured retinas. Alternatively, this could be performed in injured
retinas from wild-type or Alcama mutant fish. Bioinformatic analysis would then point to
specific regenerative pathways or processes that are altered with Alcama inhibition.
Immunprecipiation may need to be performed from whole retina tissue as opposed to
FAC-sorted cells due to the practical consideration of the amount of starting material required. Subsequent mass spectrometry could then identify putative binding partners of
Alcama, providing additional information on downstream signaling pathways these
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interactions may stimulate. It would also be useful to perform this on mouse retinas to
determine whether there are any differences in Alcama binding partners between fish
and mammals. These studies could conceivably be performed in vitro on Müller glia
cultured from both fish and mouse retinas, which could be a more practical, alternative
strategy.
While we search for extracellular interaction partners and intracellular pathways
involved with Alcama signaling, literature may offer some insights into a specific,
hypothesis-driven pathway to explore. Wnt signaling offers a path forward, which has
been implicated in extracellular signaling, regeneration, and stem cell proliferation.240–243
For example, Müller glia and other cells may respond to injury by releasing signals, such
as Wnt ligands, that induce expression of various molecules involved in regeneration,
including Alcama. The resulting Müller glia-derived progenitors may themselves express
Alcama, or a potential interaction partner such as E-cadherin, with an extracellular
domain that could conceivably interact with Alcama. Co-localization between ALCAM and
E-cadherin has been observed, but direct interactions have not been shown.226,228 The
cytosolic domain of E-cadherin interacts with β-catenin; upon stimulation with Wnt ligands via Frizzled receptors, β-catenin translocates to the nucleus to induce transcription factors that initiate processes including cell adhesion and proliferation.241,244
Following injury in chick retinas, nuclear accumulations of β-catenin have been found in
progenitor cells, peaking at 3 days post injury, when the proliferation of progenitors in
zebrafish is also the most pronounced.132,243 Although Wnt signaling promotes progenitor cell proliferation, it inhibits differentiation,147,241 and a timing component must be
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considered in future experiments to determine how and why these signals stop.
Furthermore, E-cadherin has been shown to facilitate the movements of cell clusters
through Rac1 signaling and cytoskeletal rearragnement.217 Similarly, Rac1 is required for
Müller glia proliferation after injury in rats.245 In a mammalian retina, this Rac1-driven
Müller glia proliferation may be inappropriately robust (i.e. gliosis), but in a fish retina,
this proliferation may be limited to progenitor cells that must proliferate to reconstitute
the damaged retina. Homophilic Alcama interactions are also known to induce
cytoskeletal changes that mediate cell-cell and cell-ECM contacts.221,225 Such cytoskeletal
changes may not only facilitate interactions between Müller glia and progenitor cells, but
also initiate changes within the Müller glia themselves. These changes include interkinetic
nuclear migration or modulation of cell polarity, which also have important implications
in regeneration and are facilitated by Rho/Rock signaling downstream of Wnt11.202,246
Incidentally, zebrafish Müller glia upregulate expression of Wnt11 and Rho after injury.247
Thus, a proposed mechanism of action may be that the Alcama expressed by
Müller glia in response to injury interacts with E-cadherin and/or Alcama expressed by
progenitor cells to simultaneously mediate progenitor cell proliferation and migration
(Fig. 31). Continued stimulation between Müller glia and progenitor cells may also induce
positive feedback loops, resulting in release of additional Wnt ligands or other growth
factors. This model can serve as a guide for our immediate future directions to test the hypothesis that Alcama expressed by Müller glia interacts with E-cadherin expressed by
Müller glia-derived progenitor cells to initiate or continue intracellular Wnt signaling. A
similar mechanism may be at play with other cadherins, such as N-cadherin, previously
114 shown to play a role in neurogenic cluster formation in the zebrafish retina.123 Some parts of this mechanism may be clarified or confirmed by the aforementioned RNA sequencing and immunoprecipitation experiments, in addition to identifying other hypotheses to explore. E-cadherin (or N-cadherin) would be an expected binding partner to support our proposed mechanism, identified by an anti-Alcama immunoprecipitation. Furthermore,
RNA sequencing can reveal downstream signaling targets of Alcama and related regenerative pathways for identifying and testing novel hypotheses. Notably, a recent study has reported differences between Müller glia from zebrafish, chick, and mouse retinas using single cell RNA sequencing.247 This study corroborates our finding that
Alcama expression is upregulated in zebrafish Müller glia, but not in mouse Müller glia, in response to outer nuclear layer injury. In addition, zebrafish Müller glia exhibit increased expression of Dkk’s, a readout for Wnt signaling.247 Importantly, though, this study was performed in Müller glia, but we will need to specifically identify Müller glia-derived progenitor cells to fully test our mechanistic hypothesis.
In vitro assays will also serve as valuable tools for conducting preliminary mechanistic and functional studies. Continued development of our cell culture method
(Appendix 1) will be necessary to be able to study fish cells in culture. In the meantime, we can use CRISPR/Cas9 technology in immortalized human Müller cell lines to knockout or overexpress ALCAM. (Note that ‘Alcama’ refers to the zebrafish protein while ‘ALCAM’ refers to the human/mammalian homolog). In these systems, injury can be simulated by mechanical disturbances or with addition of factors such as IL-6 or HB-EGF that have been shown to be necessary and sufficient for Müller glia activation.136,142,149 This will allow us
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to compare effects of ALCAM expression at various levels with subsequent mechanistic
analyses through Western blotting or qPCR of molecules involved in the pathways hypothesized to be involved with ALCAM signaling. In the example of our proposed mechanism (Fig. 31), if ALCAM is downregulated, we would expect to see downregulation of Dkk’s as a readout for Wnt signaling, or a reduction in β-catenin accumulation in progenitors (if present in culture). Scratch assays would be particularly useful to elucidate effects of ALCAM expression on cell migration in mammalian Müller cells, with the scratch serving as a pseudo-injury. Here, we might expect that without ALCAM, cells would migrate to fill the scratch more slowly than control cells and more quickly when ALCAM is overexpressed. Ultimately, we will need to compare the function of ALCAM/Alcama in both fish and mammalian systems to fully understand how this adhesion molecule is functioning during regeneration. In vitro, this may occur with primary cell cultures of mammalian and zebrafish Müller glia. This will likely not be a problem for mammalian cells as there are readily available cell lines from both humans and mice. While we have made some progress in developing a primary culture system of zebrafish retinal cells, and it may be a useful system as-is for some applications (Appendix 1), it is unknown whether a pure culture of zebrafish Müller glia is possible at present.
In the most straightforward interpretation of these future in vitro results, we might expect that ALCAM must be overexpressed to have an effect, if any, on downstream signaling or cell migration, to enact the effects that occur inherently in zebrafish Müller cells or cells that express zebrafish Alcama. Similarly, ALCAM knockdown would not differ greatly from wild-type ALCAM-expressing cells, which may not induce detectable
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signaling or express ALCAM at all, as we have not seen ALCAM expression in mouse Müller
glia with or without injury (Fig. 21). These predictions are based on the assumption that
there is something unique about Alcama expression in zebrafish specifically, whether it is
Alcama itself, the proteins with which it interacts, and/or the downstream signaling
pathways it may activate. On the other hand, it is very possible that none of these result
can be easily interpreted within the context of regeneration. We can simulate damage in
vitro to some extent as mentioned previously, but we cannot be sure whether this
appropriately recapitulates the necessary circumstances of injury in vivo. For example,
immortalized Müller glia cell lines do not produce progenitor cells since the Müller glia
themselves must continually proliferate to maintain the cell lines. This is comparable in
some ways to what occurs in mammals in vivo, but is far from the reactive gliosis that
leads to retinal scar formation. Conversely, after zebrafish retinal injury, an essential
component of the regenerative process is generation of progenitor cells through robust
self-renewal, which may be an important part in Alcama signaling within and between
Müller glia and the progenitor cell population. The in vitro results, however, can point to
Alcama-dependent mechanisms that can be applied to in vivo experiments. For example, we may induce expression of ALCAM in a mouse or rat model after laser ablation injury to determine whether ALCAM itself could reduce the scarring response and/or induce a shift toward a regenerative response. Another strategy might be to induce endogenous expression of ALCAM with injection of retinal stem cells to help them move within the retina.
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It is important to note that Alcama is probably not the ultimate solution to retinal regeneration; however, these experiments serve as a valuable starting point to identify particular adhesion molecules that play a role in the process. We are beginning to improve our understanding of the role of migration in regeneration for future experiments and applications for therapeutic options, such as creating scaffolds for implanted stem cells.
Genetic models of Alcama mutations or knockdowns in zebrafish will likely result in more robust changes compared to morpholino treatment, which is not a complete knockdown across the entire retina. This is an important consideration when analyzing regeneration within individual lesions that may or may not have been completely targeted by the morpholino. Another option for future experiments would be to develop double mutants or inducible knockouts of Alcama in combination with other adhesion molecules that may play roles in retinal regeneration. These could be identified through the aforementioned
RNA sequencing or immunoprecipitation experiments, which would point to molecules that interact directly or indirectly with Alcama, and knocking them down with Alcama would likely intensify the effect of an Alcama knockdown alone on regeneration.
Additional research may identify molecules involved in progenitor cell migration unrelated to Alcama, which will offer equally important avenues to explore in the further elucidation of progenitor cell migration mechanisms in zebrafish retinal regeneration.
Doing so will subsequently lead to comparative studies in mammalian systems that will ultimately inform future therapeutic strategies for proper guidance and integration of retinal stem cells in humans.
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The extracellular matrix and retinal regeneration
As discussed previously, there is an intimate association between adhesion molecules and
the extracellular matrix (ECM), particularly where cell migration is involved. There is a
similarly close association between inflammation and the extracellular matrix, which must respond appropriately to signals of damage just as the cells do in order to facilitate productive repair mechanisms.248 In addition to exploring what these interactions may be
with Alcama specifically, the overall involvement of the extracellular matrix in retinal
regeneration represents an important and largely understudied aspect of the process,
parallel to the scarce attention cell migration has received in the field. The retinal ECM is
a diverse and dynamic composition of various proteoglycans, laminins, collagens,
glycoproteins, and other molecules that differ depending on the developmental stage and
retinal location.178–180,249
Specific components of the ECM contribute to scar formation and may be
responsible for preventing regeneration in mammals, at least in part by preventing
cellular and axonal movement to the necessary locations.203,206 For example, tenascin-C
has been shown to support the de-differentiation of Müller glia but inhibit retinal
stem/progenitor cell proliferation during development in embryonic mouse retinas.178,250
Interestingly, this may have been due, in part, to interactions with the Wnt signaling pathway, contributing to the differentiation of the stem/progenitor cell population.250
Tenascin-C has also been shown to contribute to stem cell niche formation and influence
cell adhesion during development.250–252 Its potential interactions with Alcama
throughout the process of zebrafish retinal regeneration may shed light on both adhesion
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molecules and the ECM for their importance in creating an environment conducive to
productive regeneration or stem cell therapies in a mammalian retina. Tenascin-C
knockdown, for example, results in an upregulation of a particular chondroitin sulfate
proteoglycan (CSPG) that is inhibitory to pathfinding of growing neurites’ axonal
projections.178,250,253 Furthermore, CSPGs are major components of glial scars in the
mammalian CNS in response to damage, protecting the retina from further damage at the
cost of inhibiting regeneration.254 Conversely, ECM components such as laminins and enzymes including matrix metalloproteinase 9 (MMP-9) promote a pro-growth
environment conducive to regeneration.178,248,255 Overall, it will be critical to fully
understand the roles of specific extracellular matrix components in relation to their
associated adhesion molecules throughout the processes of retinal regeneration in fish
and scarring in mammals. The appropriate constituents must then be appropriately
manipulated away from scarring tendencies and toward productive regeneration and
implemented accordingly in future regenerative therapies such as ECM hydrogels and
stem cell transplantation.178,256
A preliminary in vitro experiment may involve a scratch assay with Müller cell cultures, similar to what was described in the preceding section, in order to determine how interactions between Alcama and the ECM may contribute to cell migration. Cells
would be compared expressing or not expressing Alcama while plated on different
combinations of substrates including laminin, collagens, tenascins, and proteoglycans.
After scratching, cells would be monitored for their ability to fill in the gap and migrate
toward one another. There would be several caveats in the interpretation of these results.
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First and foremost, these experiments would be initially executed in mammalian Müller
cell cultures (e.g. MIO-M1, a spontaneously immortalized human Müller cell line). This means that the cells may behave differently than they would in vivo, and different still from zebrafish Müller cells. Furthermore, the in vitro cultures will likely not produce progenitor cells that would normally be migrating along the Müller glia, whereas the migration of Müller glia themselves would be analyzed in the culture system.
Nevertheless, in vitro assays will serve as a valuable starting point for determining particular ECM components that may be contributing to the movement of retinal cells.
These findings could then be translated to other models such as zebrafish retinal cell cultures, retinal explants, or in vivo mammalian systems.
The role of microglia in retinal regeneration
Although beyond the scope of this work, an important avenue to explore in future projects involves the role of inflammation, microglia, and other immune cells in retinal regeneration, which may include functions of Alcama in slightly different contexts than previously discussed. We have seen that microglia infiltrate the injury sight within a day post lesion, and begin to retreat as regeneration is underway (Fig. 32). While these immune cells may be cleaning up debris from the damage, this is likely not their sole function. There is significant evidence to suggest that microglia play an important part in the regenerative response, both independently and through interactions with Müller glia.25,31,257,258 Indeed, microglia ablation leads to reduced formation of Müller glia- derived progenitor cells and a decreased rate of photoreceptor regeneration.259,260 These
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responses may be mediated by a release of pro-inflammatory cytokines including IL-6
family members, known to induce Müller glia activation.149,261
In regards to a potential connection between Alcama: the name itself – activated leukocyte cell adhesion molecule – points to its potential connection to immune cell function. It was first identified in activated leukocytes and is required for T-cell activation and proliferation through interaction with CD6 at dendritic cell/T-cell contact zones.226,262,263 It would therefore be interesting to ask whether a similar interaction
might be occurring in the retina. Perhaps the retinal or choroidal vasculature expresses
Alcama and is involved in the recruitment and migration of immune cells into the retina,
previously shown to promote trafficking of lymphocytes and monocytes across the blood brain barrier into the central nervous system.264 Alternatively, Alcama could be expressed by resident microglia either in response to injury, helping them migrate to the damaged areas, or constitutively expressed, aiding the movement of other cells by upregulating expression of Alcama interaction partners in response to injury.
The large scale experiments mentioned previously (e.g. RNA-seq,
immunoprecipitation) may again help to shed light on these hypotheses, as well as
develop others related to the role of the immune system in zebrafish retinal regeneration.
More targeted experiments would likely rely heavily on flow cytometry analysis, a
valuable and well-established tool for immune cell characterization, in which case a
reliable Alcama antibody would become essential. For example, initial experiments may
consist of running flow panels for retinal cells before and after injury at various time
points, including markers for immune cells and Alcama to gain a preliminary
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understanding of the immune cell profile in relation to Alcama expression over the course
of the regenerative process. These experiments could be carried out with cells from
Tg(mpeg1:gfp), a fish line in our care whose microglia express GFP. This also brings up the
potential utility of generating a transgenic fish expressing a fluorescent marker under control of the Alcama promoter. While such a fish provides slightly less flexibility than an antibody in labeling experiments such as flow cytometry and immunohistochemistry and would be more time consuming to generate initially, it may ultimately be the most versatile and reliable option for detecting Alcama expression consistently in a variety of applications. In this way, we can run the described flow panels to identify which cell types express Alcama, overlaid onto the immune cell profile in a regenerating zebrafish retina.
Once a clear picture in zebrafish begins to take form, we can also run similar flow cytometry panels on mouse retinas to identify differences in immune cell responses, further contributing to the understanding of how the balance may be tipped from a gliotic, scarring response in mammalian retinas toward a productive, regenerative response inherent to zebrafish retinas.
Concluding remarks
Overall, this work has contributed to the field of retinal regeneration in several ways. First,
we have developed a reliable, reproducible method for isolating Müller glia from
zebrafish retinas for subsequent analysis of a single cell type with the inherent capacity
for regeneration. This serves as a valuable research tool for use in a variety of applications
such as flow cytometry analysis, RNA sequencing, or cell culture that can improve our
understanding of the regenerative process. Secondly, we have shown for the first time
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that the adhesion molecule, Alcama, plays a role in zebrafish retinal regeneration. Alcama is upregulated by Müller glia in response to injury and, when inhibited, leads to reduced efficiency of progenitor cell migration and decreased capacity for overall regeneration.
This work serves as a foundation on which to base future studies confirming the role of
Alcama in retinal regeneration with a genetic zebrafish model. Ultimately, we hope to convey the importance of understanding the cells and mechanisms underlying all parts of the regenerative process, including Müller glia and progenitor cell migration, which will be essential for developing therapies to effectively restore lost vision.
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Figure 31. Proposed model for Alcama-mediated mechanism of action. Müller glia upregulate Alcama expression in response to injury stimuli, primarily on the Müller cell processes that span the inner nuclear layer (INL) to outer nuclear layer (ONL) when the injury occurs to the ONL. We hypothesize that Müller glia- expressed Alcama interacts with E-cadherin and/or Alcama expressed on the surface of progenitor cells, with the Müller glia acting as a scaffold to direct their migration from INL to ONL. Inset: intracellular signaling in the progenitor cells (purple membrane) and Müller glia (green membrane) in response to Alcama/E-cadherin and Alcama/Alcama interactions. In progenitor cells, β-catenin interacts with the cytoplasmic tail of E-cadherin; in response to Wnt pathway activation (via Frizzled receptors), β-catenin translocates and accumulates in the nucleus to enact cellular changes including proliferation and migration. Homophilic Alcama interactions stimulate cytoskeletal changes via interactions with the cytoplasmic tail of Alcama and Rho/Rac signaling, both in the progenitor cells and Müller glia. E-cadherin-mediated stimulation of Müller glia-expressed Alcama may also activate downstream pathways that results in a positive feedback loop of signaling molecule release, including Wnt ligands. Created with BioRender.com.
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Baseline 1 dpl 3 dpl 7 dpl
Figure 32. Microglia infiltrate the injury site after laser ablation of photoreceptors. All zebrafish retina sections (10 μm) are stained for microglia (green, 4c4 antibody) and counterstained with propidium iodide. Fish received OCT-guided laser photocoagulation and were sacrificed at the indicated days post lesion (dpl). Microglia are present, though difficult to distinguish, at baseline (uninjured) because of their resting state morphology. 40x magnification; scale bar = 100 μm.
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APPENDIX 1
Primary culture of zebrafish retinal cells
Using the dissociation procedure detailed in Chapter 2, we began developing a primary cell culture method for zebrafish retinal cell suspensions. The original goal was to obtain pure cultures of Müller glia; while this has not yet been achieved, we are able to culture whole retina cell suspensions from zebrafish for several weeks. This primary cell culture system can serve as a valuable in vitro tool for studying the regenerative potential of zebrafish retinas. Although additional work needs to be done to fully characterize the system before constituting a comprehensive project or paper, its current status is reported herein.
Cell culture methods
Cell suspensions were obtained from Tg(apoe:gfp) retinas using the dissociation protocol described previously (Materials and methods, Chapter 2). The final suspension was made in room temperature complete cell culture media rather than sorting buffer: 10% FBS, 1%
GlutaMax, and 1% penicillin/streptomycin in DMEM, 0.22 μm-filtered. Cell suspensions were then plated onto chamber slides coated with 10 μg/cm2 laminin at 250,000 cells/cm2 and incubated in a humidified environment at 28 °C, 5% CO2. The incubation temperature was chosen to match the temperature at which our fish are raised and maintained. Cells are left undisturbed for at least 7 days before changing the media. We have been able to maintain these cultures for 3-4 weeks. We have not yet been able to optimize the media
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change schedule: these cells seem to be particularly sensitive to the mechanical
disturbance that comes with media changes, but if there is too much time between
feedings, the cells become unhealthy and begin to die off.
The culture substrate was chosen after comparing the morphologies of the cells,
particularly Müller glia, when cultured on three different materials: collagen IV, laminin, and fibronectin (Fig. 33). The GFP-positive Müller glia grown on fibronectin are quite flat and globular, and they do not resemble the expected morphology exhibited in vivo
(Chapter 2, Fig. 7). Furthermore, there are significant amounts of RPE cells also stuck to
the plate; while this may be interesting for cultures of organoids or similar in vitro
recapitulation of the full retinal environment, for our purposes, this makes visualization
difficult and does not promote native Müller glia characteristics. The collagen IV substrate
yields similar results, although some elongated, adherent GFP+ Müller glia are visible
here. Nearly all Müller cells grown on laminin exhibit their typical morphology: elongated
cell bodies with extensive processes, and there are minimal adherent RPE. Laminin was
therefore chosen as the growth substrate for all subsequent experiments.
Immunocytochemistry methods
For standard immunocytochemistry staining, we have primarily utilized the proceeding
protocol, which produced positive signal for most of the antigens we have probed for.
Depending on the antigen/antibodies of interest in future, however, this may need to be adjusted or further optimized. To fix the cells, media was removed and 4% paraformaldehyde (PFA) in 1x PBS was immediately and incubated at room temperature.
Cells should not be rinsed with PBS before fixing, as they will very easily detach. Fixative
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was removed and cells washed three times, 10 minutes each in 1x PBST (0.1% Tween-20
in 1x PBS). Cells were then incubated in 0.1% Triton X-100 for 10 minutes before washing
in 1x PBST three times, five minutes each. To block non-specific binding, cells were
incubated for 1-2 hours at room temperature in blocking solution: 5% goat serum, 1%
DMSO, 1% Tween-20, and 2% BSA in 1x PBS. Cells were incubated overnight at 4 °C in
primary antibody (Table 4) diluted in blocking solution, or in blocking solution for
secondary-only controls. Primary antibodies were removed and washed three times for
10 minutes each in 1x PBST before incubating in secondary antibody (Table 4) diluted in
blocking solution for one hour at room temperature in the dark. Cells were then washed
three times, 10 minutes each in 1x PBST before incubating in DAPI solution (Thermo
Scientific, #62248, 1 mg/mL) diluted 1:1000 in 1x PBS for 10 minutes at room temperature in the dark. The chamber slide walls were removed, and the slide was rinsed briefly in 1x
PBST before mounting with Fluoromount-G mounting media: this must be applied in
excess before covering with glass coverslip to avoid bubbles around the hydrophobic
barriers of the chamber slide and allowed to dry at least 1-2 days before imaging.
To label the cultures with BrdU, we used a BrdU Labeling and Detection Kit (Roche,
#11444611001). BrdU Labeling Reagent was added to cell cultures without changing the
media on days 1, 2, and 3 in culture by diluting in cell culture medium and adding to wells
so that the final concentration in culture was 10 μM, as recommended by the
manufacturer. For detection, media was removed and ice cold fixative immediately
added: 50 mM glycine in 70% ethanol, pH = 2.0. Cells were incubated at -20 °C for 20
minutes to fix, then washed three times, five minutes each in 1x PBS at room
129 temperature. Cells were incubated for 30 minutes at 37 °C in anti-BrdU antibody diluted
1:10 in Incubation Buffer, both of which are provided in the kit. Cells were washed three times, five minutes each in 1x PBS before incubating for 30 minutes at 37 °C in secondary antibody. We used goat anti-mouse IgG, AlexaFluor-633 diluted 1:500 in 1x PBS instead of the fluorescein provided in the kit since we would be co-labeling for GFP-positive
Müller glia (Table 4). The cells were washed twice in PBS for five minutes each and twice in PBST for ten minutes each. Cells were then stained for GFP following the standard immunocytochemistry protocol described above, beginning with the blocking step.
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A
Fibronectin Collagen IV Laminin
B
Figure 33. Comparison of zebrafish whole retina primary cultures grown on various substrates. A) Top: overlay of transmitted light with GFP fluorescence. Bottom: GFP fluorescence (ApoE:GFP, Müller glia). Fibronectin (left) gives flattened, pancake-like Müller glia morphology with excessive RPE adherence. Collagen IV (middle) promotes round, flat cells and some with elongated Müller glia-like morphology, as well as some RPE growth. Laminin (right) allows almost exclusive growth of GFP+ cells with characteristic Müller glia morphology and minimal RPE adherence. Images taken at 3-4 days in culture; 10x magnification; scale bars = 200 μm. B) Whole retina primary cultures grown on laminin substrate days demonstrating large clusters of small, round cells growing around the GFP-positive Müller glia. Left: overlay of transmitted light with GFP fluorescence. Right: GFP fluorescence (ApoE:GFP Müller glia). Images taken at 6 days in culture; 63x magnification; scale bars = 25 μm.
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Preliminary characterization of primary zebrafish retinal cell cultures
We first wanted to determine whether the GFP-positive Müller glia maintained their
identity in culture. Cells were fixed after six days in culture and stained for GFP, GFAP, and
DAPI (Fig. 34). There is near complete overlap of the GFP and GFAP signals, keeping in
mind that GFAP is an intermediate filament protein and appears filamentous, while GFP
is cytoplasmic with some perinuclear localization. In addition, clear differences can be seen in the nuclei of Müller glia and the other GFP-negative cells: Müller glia nuclei are
diffusely staining with distinct nucleoli, and they are surrounded by these GFP-negative
cells with small, round nuclei. These cultures were reminiscent of neurogenic clusters of
progenitors surrounding Müller glia observed in vivo after injury. We hypothesized that
the act of taking the cells out of the body and putting them into an in vitro environment could be simulating injury.
Assessing the proliferative capacity of primary zebrafish retinal cell cultures
To address this hypothesis, we then stained for Nestin, a neural stem cell marker, and
PCNA, a marker of proliferation to determine if these cells were actively proliferating progenitors (Fig. 35). After fixing and staining the cells after 4 days in culture, we observed that nearly every cell appeared to be Nestin-positive (Fig. 35A), which can be interpreted in multiple ways. It is possible the antibody is too non-specific and merely sticks to almost every part of the cell or to the GFP antibody. The staining is not an exact match to the GFP signal, however, indicating that it could be true labeling. The brightest signal, for example, appears to be coming from the small, round clusters of cells, as well as a few Müller glia themselves. This could mean that these cells are indeed exhibiting stem cell
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characteristics. They could also be stuck between Müller glia and stem cell identity, but
unable to commit either way because of the in vitro environment. When staining for
PCNA, however, we observed the opposite of the Nestin staining results, and no cells
appeared to be PCNA-positive (Fig. 35B). This could simply mean the antibody itself was not effective, or the staining conditions and protocol were not optimal for that antibody.
The cells could also be truly negative for PCNA when stained here at 4 days (Fig. 35B), nor at days 1, 2, or 3 in culture (data not shown). In turn, that can either mean the cells are not proliferating, or they are being captured in a phase of the cell cycle in which PCNA expression is not easily detectable, as PCNA is most highly expressed in S-phase.265
We wanted to further characterize the proliferative capacity of these cells and
determine whether they can be stimulated to proliferate. Because IL-6 has previously
been shown to be necessary and sufficient to induce Müller glia proliferation in an
uninjured zebrafish retina,149 we chose to incubate cultures in media containing rhIL-6
(10, 100, or 1000 ng/mL) for 4 days. BrdU was spiked into the cultures without changing
the media on days 1, 2, and 3. This was to maximize the chances of seeing BrdU-positive
cells, rather than to pinpoint their proliferative timeline. While BrdU-positive Müller glia
were evident in these cultures at all concentrations of IL-6, all of which were above the
expected saturation point, the same was true for cultures incubated with the vehicle
control of PBS/BSA instead of IL-6 (Fig. 36). It is evident that the cells incorporating BrdU
are Müller glia rather than the surrounding clusters of small, round cells. Because BrdU-
positive Müller glia are observed even in the control conditions, they might be
proliferating at baseline without stimulation. Serum starvation may be necessary before
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adding a stimulant such as IL-6 in order to see a difference. While cells are positive for
BrdU, it indicates that the cells are entering the cell cycle by going through S-phase,
synthesizing new DNA, and therefore incorporating BrdU into their nuclei; however, these
cells may be stuck in G2 phase and not continuing to actively divide. This hypothesis is
supported by the lack of PCNA staining (Fig. 35B), which is highly expressed in S-phase,
but downregulated in G2/M phase.265
Cell type-specific markers
Finally, we sought to determine what the identity of the non-Müller cells in culture might
be. We therefore fixed and stained cells after about five days in culture for various cell type-specific markers (Fig. 37). We found that the cell clusters consist of multiple cell types, the most prominent of which is a HuC/HuD-positive population, labeling amacrine
and ganglion cells (Fig. 37A). There was also an abundance of bipolar cells, positive for
PKC (protein kinase C) (Fig. 37B). To a lesser extent, some cone cells were also detected
by the zpr1 antibody (Fig. 37C). Fish are a cone-dominant species, and because very few
cones and virtually no rods were detected (zpr3 antibody, data not shown), these culture
conditions are likely not supportive for photoreceptors.
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GFP DAPI
GFAP Merge
Figure 34. GFAP co-labels with ApoE:GFP expression to confirm Müller glia identity in culture. GFP antibody labels GFP-expressing cells (green) from Tg(apoe:gfp) fish retinas, labeling Müller glia. GFAP antibody (Zrf1) labels glial fibrillary acidic protein (red), which almost entirely overlaps GFP signal, confirming the GFP-positive cells to be Müller glia in culture. DAPI nuclear stain (blue) labels the large, flat nuclei of Müller glia with distinct nucleoli and surrounding clusters of small, round cell nuclei. Cells fixed after 6 days in culture; 40x magnification; scale bars = 50 μm.
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A B
GFP GFP
Nestin PCNA
DAPI DAPI
Merge Merge
Figure 35. Zebrafish whole retina primary cultures are Nestin-positive but PCNA-negative. Green: GFP antibody labels GFP-expressing cells from Tg(apoe:gfp) fish retinas, labeling Müller glia. Blue: DAPI nuclear stain. Red: Nestin (A) labels neural stem cells; PCNA (B) labels proliferating cells. Most cells, both Müller glia and others, appear Nestin-positive (A). No cells appear to be PCNA-positive (B). See text for details. Cells fixed after 4 days in culture; 40x magnification; scale bars = 50 μm.
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DAPI GFP BrdU Merge +IL-6
BSA/PBS
Figure 36. Primary retinal cultures incorporate BrdU when stimulated with IL-6 and at baseline. Green: GFP antibody labels GFP-expressing cells from Tg(apoe:gfp) fish retinas, labeling Müller glia. Blue: DAPI nuclear stain. Red: BrdU incorporated by cells during S-phase. BrdU was spiked into the culture media (10 μM) on days 1, 2, and 3. Recombinant human IL-6 (rhIL-6) was present in the media for duration of culture at 10, 100, or 1000 ng/mL. No significant differences were observed for these concentrations; representative images are therefore shown here from 10 ng/mL IL-6 treatment. Vehicle control treatment (BSA/PBS solution) also showed BrdU incorporation to a similar extent. Cells fixed after 4 days in culture; 40x magnification; scale bars = 50 μm.
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DAPI GFP CTSM Merge A
HuC/HuD
B
PKC
C
Zpr1
Figure 37. Cell clusters around Müller glia primarily consist of amacrine/ganglion cells, bipolar cells, and cones. Green: GFP antibody labels GFP-expressing cells from Tg(apoe:gfp) fish retinas, labeling Müller glia. Blue: DAPI nuclear stain. Red: cell type-specific marker (CTSM); A) HuC/HuC, amacrine/ganglion cells; B) PKC (protein kinase C), bipolar cells; C) Zpr1, cone photoreceptors. Cells fixed after 5 days in culture; 40x magnification; scale bars = 50 μm.
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Applications and future directions
While these cultures need to be studied further before being fully utilized as a research
tool, we have learned a lot from these initial results. For the whole retina cell culture
system described herein, we have seen clusters of cells surrounding Müller glia. Given the
staining results, these clusters are comprised of a mixture of amacrine/ganglion cells,
bipolar cells, and cones (Fig. 37). The staining of all these markers, however, does not account for all of the cells in these clusters. This observation, in conjunction with the
Nestin, PCNA, and BrdU results suggests that there may also be some stem-like cells in these clusters, including the Müller glia themselves. Most of the cells appeared to be
Nestin-positive; no cells appeared to be PCNA-positive, but even without stimulation, incorporated BrdU (Fig. 35, Fig. 36). This suggests that the act of taking cells out of the body and into an in vitro environment is enough to activate the cells to enter a stem cell- like state. This may be true for both the Müller glia progenitor-like cells potentially derived from them.
The Müller glia may have entered the cell cycle enough to undergo S-phase and incorporate BrdU, and even give rise to a progenitor-like cell. The environment of the cell culture dish is not sufficient, however, to maintain their stemness to an extent that allows their continued proliferation and may require further stimulation to proliferate and/or exit the cell cycle. Those cells that do express markers of cones, bipolar, amacrine, or ganglion cells are therefore not likely to be differentiated from Müller glia-derived progenitor cells, but are instead mature cells that have been dissociated and plated. While
the cells were plated as single cells initially and these large clusters are not observed for
139 several days, it is possible that they converge on Müller glia over time in culture for metabolic or signaling support.
This could be tested in the future with some co-labeling experiments. For example, the cell type-specific marker antibodies could all be used in conjunction and labeled with the same secondary antibody, while Nestin and other stem cell markers are labeled with another color. If none of the cell type-specific labels overlay with the stem cell-like labels, it would support the hypothesis that these clusters surrounding the Müller glia are comprised of a mixture of mature cells from the dissociation and stem-like cells potentially derived from the Müller glia, stimulated to re-enter the cell cycle through the act of dissociation and plating. To test the hypothesis that the Müller glia might be initially dividing enough to incorporate BrdU, potentially giving rise to some stem cell-like cells that have arrested in their cell cycle, a lineage tracing assay involving BrdU and EdU may be useful. Since PCNA staining was not observed even at 1 day in culture, the proliferation is likely occurring in the first 24 hours of culture and then arresting. A spike-in of BrdU at
8 hours followed by a spike-in of EdU at 16 hours, for example, might offer more insight into when and how the Müller glia are dividing and/or giving rise to progenitor cells. Full media changes would be ideal rather than spike-ins, which may make interpretation difficult though still informative. So far, however, we have not been able to change the media without disturbing the cultures, especially at this early time point.
Dissociation and primary culture of mouse Müller glia
We have further expanded this cell culture application to murine cultures of Müller glia.
Unlike the zebrafish cultures, we were able to obtain up to 90% pure Müller glia after
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several days in culture with routine media changes. Primary cultures of murine Müller glia
were developed in collaboration with another group, recently published as part of a
comprehensive project on Müller glia metabolism, not discussed in depth here.266 While
the same general dissociation and culture procedure described previously was used,
some key adjustments were made. Dissociated cells were obtained from p11-p12 mouse
retinas, as opposed to adult fish retinas. Incubation in the papain/DNase solution was
longer since the size difference between the animals’ eyes meant there was substantially
more tissue to dissociate. Tissue was still incubated at 37 °C with 160 rpm agitation, but
for a total of about 35 minutes, broken up into 15 minute, 15 minute, and 5 minute
incubations, triturating gently after each as described previously. The centrifugation speed was also reduced to 500-550 x g, since mouse retinal cells do not seem to be as buoyant as those from zebrafish. This also meant the resuspension could be less traumatic, increasing the ability of Müller glia to adhere to the culture substrate without shearing their processes.
Cells were cultured in complete medium: DMEM-high glucose, without L-glut, with
NaPy; 1% GlutaMax; 1% penicillin/streptomycin; 10% FBS. Dissociated cells were 83%
viable, on average, at the time of plating and were plated at a density of 1.1 million
2 cells/cm on laminin-coated plates, incubated at 37 °C and 5% CO2. The first media change
occurred after four days in culture, followed by media changes every two days. After 8-
12 days in culture following this media change schedule, cultures primarily consisted of
Müller glia that had expanded to near confluence. Cells were harvested for experiments before passaging was required, which may have caused transformation of the cells that
141 was not desired for the purposes of this study. Cell identity was confirmed with Western blot and used for metabolic assays described by Singh, et al., 2020.266
A few strategies for optimizing the cell culture method of primary zebrafish Müller glia can be learned from how we were able to grow mouse Müller cells in vitro. For example, the media used for the mouse cell cultures is almost identical to the media used in zebrafish cultures except that it is high glucose. This may have contributed to the successful growth of Müller glia specifically in the mouse cultures while pure cultures of
Müller glia were never obtained from fish because Müller cells are so metabolically active.2,39 Perhaps the biggest reason the Müller glia grew and expanded was because of the age of the animals from which they were obtained. Müller glia from young mice likely retain some of their proliferative capacity from the still developing retina. While we may be able to more easily culture Müller glia from very young fish eyes, it may be difficult to obtain sufficient numbers from such small amounts of tissue. Furthermore, we wanted to develop an in vitro tool to study the regenerative capacity of adult zebrafish Müller glia, which may be concealed or distorted if using cultures obtained from young fish. An alternative strategy going forward may therefore be to develop an injury method for primary retinal cell cultures.
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APPENDIX 2
Table 3. Primer sequences for RT-PCR experiments.
Product Gene Forward Primer (5’ – 3’) Reverse Primer (5’ – 3’) Length (bp) alcama AGGCACAGAAAGATGATCCG ACATTCCCCCAACTGCGTC 257 gfap GGATGAGATCCAGATGCTGAAGG CAGATCCTTCCTCTCCGTAGTGG 298 rpl13a TCTGGAGGACTGTAAGAGGTATGC AGACGCACAATCTTGAGAGCAG 150
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Table 4. Antibodies used in immunohistochemistry or immunocytochemistry applications.
Antibody Species Dilution Manufacturer (Catalogue #) Zpr1 Mouse 1:100 ZIRC (zpr1) Alcama Rabbit 1:200 GeneTex (GTX128399) Zn5 Mouse 1:4 – 1:500 ZIRC (zn-5) Zn8 Mouse 1:4 – 1:500 ZIRC (zn-8) 4c4 Mouse 1:200 Raymond Lab (Univ. Michigan) Zrf1 Mouse 1:100 ZIRC (zrf1) GFP Rabbit 1:500 Torrey Pines (TP401) GFP Chicken 1:100 Abcam (ab13970) PCNA Rabbit 1:50 SantaCruz (sc-7907) Nestin Rabbit 1:50 AnaSpec (AS-55818) PKC Mouse 1:50 Santa Cruz (sc-80) HuC/HuD Mouse 1:100 Invitrogen (A21271) BrdU Mouse 1:50 Santa Cruz (sc-32323) ALCAM/CD166 Rabbit 1:500 SinoBiological (50005-RP02) Anti-mouseIgG-AF488 Goat 1:200 (IHC) 1:500 (ICC) Invitrogen (A11029) Anti-rabbitIgG-AF488 Goat 1:500 Invitrogen (A11008) Anti-mouseIgG-AF633 Goat 1:500 Abcam (ab150119) Anti-mouseIgG-AF568 Goat 1:500 Invitrogen (A11004) Anti-rabbitIgG-AF568 Goat 1:500 Invitrogen (A11011) Anti-chickenIgY-AF488 Goat 1:200 Invitrogen (A11039) Anti-rabbitIgG-AF555 Goat 1:500 Invitrogen (A21429)
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Figure 38. SLO images of cone-labeled fish after laser injury. Tg(TαC:gfp) zebrafish retinas with three laser lesions superior to the optic nerve at 1, 7, and 14 days post lesion (dpl). SLO images taken in autofluorescence mode, showing the GFP-positive cone photoreceptors, which are visible in their characteristic cone mosaic pattern (below a shadow of the retinal vasculature). These images demonstrate the utility of the Tg(TαC:gfp) fish, with not only increased clarity of defining distinctive lesion area boundaries, but also the for simultaneous measurement of cone density within the lesioned area, offering an improved readout for cellular regeneration as it occurs over time.
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LITERATURE CITED
1. London, A., Benhar, I. & Schwartz, M. The retina as a window to the brain—from eye
research to CNS disorders. Nature Reviews Neurology 9, 44–53 (2013).
2. Vecino, E., Rodriguez, F. D., Ruzafa, N., Pereiro, X. & Sharma, S. C. Glia–neuron interactions in
the mammalian retina. Progress in Retinal and Eye Research 51, 1–40 (2016).
3. Doly, M. Transduction of the light message: from the photon to the optic nerve. Fundamental
& Clinical Pharmacology 8, 147–154 (1994).
4. Pfrieger, F. W. & Barres, B. A. New views on synapse—glia interactions. Current Opinion in
Neurobiology 6, 615–621 (1996).
5. Purves, D. et al. The Retina. Neuroscience. 2nd edition (2001).
6. Hoon, M., Okawa, H., Santina, L. D. & Wong, R. O. L. Functional Architecture of the Retina:
Development and Disease. Prog Retin Eye Res 42, 44–84 (2014).
7. Masri, R. A., Percival, K. A., Koizumi, A., Martin, P. R. & Grünert, U. Survey of retinal ganglion
cell morphology in marmoset. Journal of Comparative Neurology 0, (2016).
8. Seung, H. S. & Sümbül, U. Neuronal Cell Types and Connectivity: Lessons from the Retina.
Neuron 83, 1262–1272 (2014).
9. Lukowski, S. W. et al. A single-cell transcriptome atlas of the adult human retina. EMBO J. 38,
e100811 (2019).
10. Molday, R. S. & Moritz, O. L. Photoreceptors at a glance. J Cell Sci 128, 4039–4045
(2015).
11. Purves, D. et al. Phototransduction. Neuroscience. 2nd edition (2001).
12. Lamb, T. D. & Pugh, E. N. Phototransduction, Dark Adaptation, and Rhodopsin
Regeneration The Proctor Lecture. Invest. Ophthalmol. Vis. Sci. 47, 5138–5152 (2006).
146
13. Pugh, E. N. & Lamb, T. D. Amplification and kinetics of the activation steps in
phototransduction. Biochimica et Biophysica Acta (BBA) - Bioenergetics 1141, 111–149
(1993).
14. Kolb, H., Nelson, R., Ahnelt, P. & Cuenca, N. Chapter 1 Cellular organization of the
vertebrate retina. in Progress in Brain Research vol. 131 3–26 (Elsevier, 2001).
15. Boycott, B. B. & Hopkins, J. M. Microglia in the retina of monkey and other mammals; Its
distinction from other types of glia and horizontal cells. Neuroscience 6, 679–688 (1981).
16. Ling, E. A. & Leblond, C. P. Investigation of glial cells in semithin sections. II. Variation
with age in the numbers of the various glial cell types in rat cortex and corpus callosum. J.
Comp. Neurol. 149, 73–81 (1973).
17. Rathnasamy, G., Foulds, W. S., Ling, E.-A. & Kaur, C. Retinal microglia - A key player in
healthy and diseased retina. Prog. Neurobiol. 173, 18–40 (2019).
18. Hume, D. A., Perry, V. H. & Gordon, S. Immunohistochemical localization of a
macrophage-specific antigen in developing mouse retina: phagocytosis of dying neurons and
differentiation of microglial cells to form a regular array in the plexiform layers. J. Cell Biol.
97, 253–257 (1983).
19. Lee, J. E., Liang, K. J., Fariss, R. N. & Wong, W. T. Ex vivo Dynamic Imaging of Retinal
Microglia using Time-lapse Confocal Microscopy. Invest Ophthalmol Vis Sci 49, 4169–4176
(2008).
20. Katz, L. C. & Shatz, C. J. Synaptic activity and the construction of cortical circuits. Science
274, 1133–1138 (1996).
21. Li, L., Eter, N. & Heiduschka, P. The microglia in healthy and diseased retina. Exp. Eye
Res. 136, 116–130 (2015).
147
22. Schafer, D. P. et al. Microglia Sculpt Postnatal Neural Circuits in an Activity and
Complement-Dependent Manner. Neuron 74, 691–705 (2012).
23. Saijo, K. & Glass, C. K. Microglial cell origin and phenotypes in health and disease. Nature
Reviews Immunology 11, 775–787 (2011).
24. O’Koren, E. G., Mathew, R. & Saban, D. R. Fate mapping reveals that microglia and
recruited monocyte-derived macrophages are definitively distinguishable by phenotype in
the retina. Scientific Reports 6, 1–12 (2016).
25. Mitchell, D. M., Lovel, A. G. & Stenkamp, D. L. Dynamic changes in microglial and
macrophage characteristics during degeneration and regeneration of the zebrafish retina. J
Neuroinflammation 15, 163 (2018).
26. Combadière, C. et al. CX3CR1-dependent subretinal microglia cell accumulation is
associated with cardinal features of age-related macular degeneration. J. Clin. Invest. 117,
2920–2928 (2007).
27. O’Koren, E. G. et al. Microglial Function Is Distinct in Different Anatomical Locations
during Retinal Homeostasis and Degeneration. Immunity 50, 723-737.e7 (2019).
28. Geissmann, F., Jung, S. & Littman, D. R. Blood monocytes consist of two principal subsets
with distinct migratory properties. Immunity 19, 71–82 (2003).
29. Feng, L. et al. Microglial proliferation and monocyte infiltration contribute to
microgliosis following status epilepticus. Glia 67, 1434–1448 (2019).
30. Käufer, C. et al. Chemokine receptors CCR2 and CX3CR1 regulate viral encephalitis-
induced hippocampal damage but not seizures. PNAS 115, E8929–E8938 (2018).
31. London, A. et al. Neuroprotection and progenitor cell renewal in the injured adult
murine retina requires healing monocyte-derived macrophages. Journal of Experimental
Medicine 208, 23–39 (2011).
148
32. Faustino, J. et al. CX3CR1-CCR2-dependent monocyte-microglial signaling modulates
neurovascular leakage and acute injury in a mouse model of childhood stroke. J. Cereb. Blood
Flow Metab. 39, 1919–1935 (2019).
33. O’Sullivan, M. L. et al. Astrocytes Follow Ganglion Cell Axons to Establish an Angiogenic
Template During Retinal Development. Glia 65, 1697–1716 (2017).
34. Koke, J. R., Mosier, A. L. & García, D. M. Intermediate filaments of zebrafish retinal and
optic nerve astrocytes and Müller glia: differential distribution of cytokeratin and GFAP. BMC
Res Notes 3, 50 (2010).
35. Lyons, D. A. & Talbot, W. S. Glial Cell Development and Function in Zebrafish. Cold
Spring Harb Perspect Biol 7, (2015).
36. Raymond, P. A., Barthel, L. K., Bernardos, R. L. & Perkowski, J. J. Molecular
characterization of retinal stem cells and their niches in adult zebrafish. BMC Dev Biol 6, 36
(2006).
37. Conrad, M., Lemb, K., Schubert, T. & Markl, J. Biochemical identification and tissue-
specific expression patterns of keratins in the zebrafish Danio rerio. Cell Tissue Res. 293, 195–
205 (1998).
38. Bringmann, A. et al. Müller cells in the healthy and diseased retina. Prog Retin Eye Res
25, 397–424 (2006).
39. Sorrentino, F. S., Allkabes, M., Salsini, G., Bonifazzi, C. & Perri, P. The importance of glial
cells in the homeostasis of the retinal microenvironment and their pivotal role in the course
of diabetic retinopathy. Life Sciences 162, 54–59 (2016).
40. Garlipp, M. A. & Gonzalez-Fernandez, F. Cone outer segment and Müller microvilli
pericellular matrices provide binding domains for interphotoreceptor retinoid-binding
protein (IRBP). Experimental Eye Research 113, 192–202 (2013).
149
41. Höfer, D. & Drenckhahn, D. Molecular heterogeneity of the actin filament cytoskeleton
associated with microvilli of photoreceptors, Müller’s glial cells and pigment epithelial cells of
the retina. Histochemistry 99, 29–35 (1993).
42. Toft-Kehler, A. K., Skytt, D. M. & Kolko, M. A Perspective on the Müller Cell-Neuron
Metabolic Partnership in the Inner Retina. Mol. Neurobiol. 55, 5353–5361 (2018).
43. Newman, E. & Reichenbach, A. The Müller cell: a functional element of the retina.
Trends in Neurosciences 19, 307–312 (1996).
44. Sarthy, V. & Ripps, H. The Retinal Müller Cell: Structure and Function. (Springer Science
& Business Media, 2001).
45. Tout, S., Chan-Ling, T., Holländer, H. & Stone, J. The role of Müller cells in the formation
of the blood-retinal barrier. Neuroscience 55, 291–301 (1993).
46. Reichenbach, A. & Robinson, S. R. Phylogenetic constraints on retinal organisation and
development. Progress in Retinal and Eye Research 15, 139–171 (1995).
47. Bringmann, A. & Wiedemann, P. Müller Glial Cells in Retinal Disease. OPH 227, 1–19
(2012).
48. Scott, A. W., Bressler, N. M., Ffolkes, S., Wittenborn, J. S. & Jorkasky, J. Public Attitudes
About Eye and Vision Health. JAMA Ophthalmol 134, 1111–1118 (2016).
49. Landau, K. & Kurz-Levin, M. Retinal disorders. Handb Clin Neurol 102, 97–116 (2011).
50. Retinal Degenerative Diseases | Nano Retina. http://www.nano-retina.com/retinal-
degenerative-diseases/.
51. Low Vision Resources Center — Numbers of People with Macular Degeneration and
Other Retinal Diseases. http://lowvision.preventblindness.org/eye-conditions/numbers-of-
people-with-macular-degeneration-and-other-retinal-diseases/.
150
52. Veleri, S. et al. Biology and therapy of inherited retinal degenerative disease: insights
from mouse models. Dis Model Mech 8, 109–129 (2015).
53. Zarbin, M. Cell-Based Therapy for Retinal Disease: The New Frontier. Methods Mol. Biol.
1834, 367–381 (2019).
54. Age-Related Macular Degeneration: Facts & Figures. BrightFocus Foundation
https://www.brightfocus.org/macular/article/age-related-macular-facts-figures (2015).
55. Fritsche, L. G. et al. Age-related macular degeneration: genetics and biology coming
together. Annu Rev Genomics Hum Genet 15, 151–171 (2014).
56. Wong, W. L. et al. Global prevalence of age-related macular degeneration and disease
burden projection for 2020 and 2040: a systematic review and meta-analysis. The Lancet
Global Health 2, e106–e116 (2014).
57. Swaroop, A., Chew, E. Y., Rickman, C. B. & Abecasis, G. R. Unraveling a multifactorial
late-onset disease: from genetic susceptibility to disease mechanisms for age-related macular
degeneration. Annu Rev Genomics Hum Genet 10, 19–43 (2009).
58. Age-Related Eye Disease Study Research Group. Risk factors associated with age-related
macular degeneration. A case-control study in the age-related eye disease study: Age-Related
Eye Disease Study Report Number 3. Ophthalmology 107, 2224–2232 (2000).
59. Vingerling, J. R., Hofman, A., Grobbee, D. E. & de Jong, P. T. Age-related macular
degeneration and smoking. The Rotterdam Study. Arch. Ophthalmol. 114, 1193–1196 (1996).
60. Age-Related Eye Disease Study Research Group. A randomized, placebo-controlled,
clinical trial of high-dose supplementation with vitamins C and E, beta carotene, and zinc for
age-related macular degeneration and vision loss: AREDS report no. 8. Arch. Ophthalmol.
119, 1417–1436 (2001).
151
61. Haines, J. L. et al. Complement factor H variant increases the risk of age-related macular
degeneration. Science 308, 419–421 (2005).
62. Chen, W. et al. Genetic variants near TIMP3 and high-density lipoprotein–associated loci
influence susceptibility to age-related macular degeneration. Proc Natl Acad Sci U S A 107,
7401–7406 (2010).
63. Fritsche, L. G. et al. Seven new loci associated with age-related macular degeneration.
Nature Genetics 45, 433–439 (2013).
64. Zając-Pytrus, H. M., Pilecka, A., Turno-Kręcicka, A., Adamiec-Mroczek, J. & Misiuk-Hojło,
M. The Dry Form of Age-Related Macular Degeneration (AMD): The Current Concepts of
Pathogenesis and Prospects for Treatment. Adv Clin Exp Med 24, 1099–1104 (2015).
65. Pennington, K. L. & DeAngelis, M. M. Epidemiology of age-related macular degeneration
(AMD): associations with cardiovascular disease phenotypes and lipid factors. Eye Vis (Lond)
3, (2016).
66. Heesterbeek, T. J., Lorés-Motta, L., Hoyng, C. B., Lechanteur, Y. T. E. & den Hollander, A.
I. Risk factors for progression of age-related macular degeneration. Ophthalmic and
Physiological Optics n/a, (2020).
67. Campochiaro, P. A., Soloway, P., Ryan, S. J. & Miller, J. W. The pathogenesis of choroidal
neovascularization in patients with age-related macular degeneration. Mol. Vis. 5, 34 (1999).
68. Kovach, J. L., Schwartz, S. G., Flynn, H. W. & Scott, I. U. Anti-VEGF Treatment Strategies
for Wet AMD. J Ophthalmol 2012, (2012).
69. Wolff, B. et al. Ten-year outcomes of anti-vascular endothelial growth factor treatment
for neovascular age-related macular disease: a single center French study. Clinical &
Experimental Ophthalmology n/a,.
152
70. Ludwig, P. E., Freeman, S. C. & Janot, A. C. Novel stem cell and gene therapy in diabetic
retinopathy, age related macular degeneration, and retinitis pigmentosa. Int J Retina Vitreous
5, 7 (2019).
71. Hartong, D. T., Berson, E. L. & Dryja, T. P. Retinitis pigmentosa. The Lancet 368, 1795–
1809 (2006).
72. Bravo-Gil, N. et al. Unravelling the genetic basis of simplex Retinitis Pigmentosa cases.
Scientific Reports 7, 1–10 (2017).
73. Prokofyeva, E., Troeger, E., Wilke, R. & Zrenner, E. Age of Visual Symptoms Onset in
Different Types of Inherited Retinal Degenerations. Invest. Ophthalmol. Vis. Sci. 51, 3548–
3548 (2010).
74. Tsujikawa, M. et al. Age at onset curves of retinitis pigmentosa. Arch. Ophthalmol. 126,
337–340 (2008).
75. RetNet: Summaries. https://sph.uth.edu/retnet/sum-dis.htm#B-diseases.
76. Sergouniotis, P. I. Inherited Retinal Disorders: Using Evidence as a Driver for
Implementation. OPH 242, 187–194 (2019).
77. Diakatou, M., Manes, G., Bocquet, B., Meunier, I. & Kalatzis, V. Genome Editing as a
Treatment for the Most Prevalent Causative Genes of Autosomal Dominant Retinitis
Pigmentosa. Int J Mol Sci 20, (2019).
78. Tanna, P., Strauss, R. W., Fujinami, K. & Michaelides, M. Stargardt disease: clinical
features, molecular genetics, animal models and therapeutic options. British Journal of
Ophthalmology 101, 25–30 (2017).
79. Burke, T. R. & Tsang, S. H. Allelic and Phenotypic Heterogeneity in ABCA4 mutations.
Ophthalmic Genet 32, 165–174 (2011).
153
80. Takkar, B., Bansal, P. & Venkatesh, P. Leber’s Congenital Amaurosis and Gene Therapy.
Indian J Pediatr 85, 237–242 (2018).
81. Miraldi Utz, V., Coussa, R. G., Antaki, F. & Traboulsi, E. I. Gene therapy for RPE65-related
retinal disease. Ophthalmic Genet. 39, 671–677 (2018).
82. Le Meur, G. et al. Safety and Long-Term Efficacy of AAV4 Gene Therapy in Patients with
RPE65 Leber Congenital Amaurosis. Mol Ther 26, 256–268 (2018).
83. Lee, R., Wong, T. Y. & Sabanayagam, C. Epidemiology of diabetic retinopathy, diabetic
macular edema and related vision loss. Eye Vis (Lond) 2, (2015).
84. Diabetes. https://www.who.int/news-room/fact-sheets/detail/diabetes.
85. Cheung, N., Mitchell, P. & Wong, T. Y. Diabetic retinopathy. The Lancet 376, 124–136
(2010).
86. Wang, W. & Lo, A. C. Y. Diabetic Retinopathy: Pathophysiology and Treatments. Int J Mol
Sci 19, (2018).
87. Roy, S., Amin, S. & Roy, S. Retinal Fibrosis in Diabetic Retinopathy. Exp Eye Res 142, 71–
75 (2016).
88. Bek, T. Diameter Changes of Retinal Vessels in Diabetic Retinopathy. Curr. Diab. Rep. 17,
82 (2017).
89. Frydkjaer-Olsen, U. et al. Correlation between Retinal Vessel Calibre and
Neurodegeneration in Patients with Type 2 Diabetes Mellitus in the European Consortium for
the Early Treatment of Diabetic Retinopathy (EUROCONDOR). ORE 56, 10–16 (2016).
90. Simó, R., Hernández, C. & Retinopathy (EUROCONDOR)*, on behalf of the E. C. for the E.
T. of D. Neurodegeneration is an early event in diabetic retinopathy: therapeutic implications.
British Journal of Ophthalmology 96, 1285–1290 (2012).
154
91. Graca, A. B., Hippert, C. & Pearson, R. A. Müller Glia Reactivity and Development of
Gliosis in Response to Pathological Conditions. Adv. Exp. Med. Biol. 1074, 303–308 (2018).
92. Belecky-Adams, T. L., Chernoff, E. C., Wilson, J. M. & Dharmarajan, S. Reactive Muller
Glia as Potential Retinal Progenitors. Neural Stem Cells - New Perspectives (2013)
doi:10.5772/55150.
93. Singh, M. S. et al. Retinal stem cell transplantation: Balancing safety and potential.
Progress in Retinal and Eye Research 75, 100779 (2020).
94. Jin, Z.-B. et al. Stemming retinal regeneration with pluripotent stem cells. Prog Retin Eye
Res 69, 38–56 (2019).
95. Hirami, Y. et al. Generation of retinal cells from mouse and human induced pluripotent
stem cells. Neurosci. Lett. 458, 126–131 (2009).
96. Osakada, F., Ikeda, H., Sasai, Y. & Takahashi, M. Stepwise differentiation of pluripotent
stem cells into retinal cells. Nat. Protocols 4, 811–824 (2009).
97. Jin, Z.-B. & Takahashi, M. Generation of retinal cells from pluripotent stem cells. Prog.
Brain Res. 201, 171–181 (2012).
98. Zhong, X. et al. Generation of three-dimensional retinal tissue with functional
photoreceptors from human iPSCs. Nat Commun 5, 4047 (2014).
99. Capowski, E. E. et al. Reproducibility and staging of 3D human retinal organoids across
multiple pluripotent stem cell lines. Development 146, (2019).
100. Hallam, D. et al. Human-Induced Pluripotent Stem Cells Generate Light Responsive
Retinal Organoids with Variable and Nutrient-Dependent Efficiency. Stem Cells 36, 1535–
1551 (2018).
101. Mandai, M. et al. iPSC-Derived Retina Transplants Improve Vision in rd1 End-Stage
Retinal-Degeneration Mice. Stem Cell Reports 8, 69–83 (2017).
155
102. Gagliardi, G., Ben M’Barek, K. & Goureau, O. Photoreceptor cell replacement in macular
degeneration and retinitis pigmentosa: A pluripotent stem cell-based approach. Prog Retin
Eye Res 71, 1–25 (2019).
103. Garita-Hernandez, M. et al. AAV-Mediated Gene Delivery to 3D Retinal Organoids
Derived from Human Induced Pluripotent Stem Cells. Int J Mol Sci 21, (2020).
104. Yoshida, T. et al. The use of induced pluripotent stem cells to reveal pathogenic gene
mutations and explore treatments for retinitis pigmentosa. Mol Brain 7, 45 (2014).
105. Cuevas, E., Parmar, P. & Sowden, J. C. Restoring Vision Using Stem Cells and
Transplantation. Adv. Exp. Med. Biol. 1185, 563–567 (2019).
106. Ortin-Martinez, A. et al. A Reinterpretation of Cell Transplantation: GFP Transfer From
Donor to Host Photoreceptors. Stem Cells 35, 932–939 (2017).
107. Mead, B. et al. Stem cell treatment of degenerative eye disease. Stem Cell Res 14, 243–
257 (2015).
108. Jorstad, N. L. et al. Stimulation of functional neuronal regeneration from Müller glia in
adult mice. Nature 548, 103–107 (2017).
109. Jorstad, N. L. et al. STAT Signaling Modifies Ascl1 Chromatin Binding and Limits Neural
Regeneration from Muller Glia in Adult Mouse Retina. Cell Rep 30, 2195-2208.e5 (2020).
110. Beach, K. M., Wang, J. & Otteson, D. C. Regulation of Stem Cell Properties of Müller Glia
by JAK/STAT and MAPK Signaling in the Mammalian Retina. Stem Cells International vol. 2017
e1610691 https://www.hindawi.com/journals/sci/2017/1610691/ (2017).
111. Lenkowski, J. R. & Raymond, P. A. Müller glia: Stem cells for generation and
regeneration of retinal neurons in teleost fish. Prog Retin Eye Res 40, 94–123 (2014).
112. Goldman, D. Müller glia cell reprogramming and retina regeneration. Nat Rev Neurosci
15, 431–442 (2014).
156
113. Thummel, R. et al. Characterization of Müller glia and neuronal progenitors during adult
zebrafish retinal regeneration. Exp Eye Res 87, 433–444 (2008).
114. Gallina, D., Todd, L. & Fischer, A. J. A comparative analysis of Müller glia-mediated
regeneration in the vertebrate retina. Experimental Eye Research 123, 121–130 (2014).
115. Chhetri, J., Jacobson, G. & Gueven, N. Zebrafish—on the move towards
ophthalmological research. Eye (Lond) 28, 367–380 (2014).
116. Bibliowicz, J., Tittle, R. K. & Gross, J. M. Towards a better understanding of human eye
disease: insights from the zebrafish, Danio rerio. Prog Mol Biol Transl Sci 100, 287–330
(2011).
117. Richardson, R., Tracey-White, D., Webster, A. & Moosajee, M. The zebrafish eye—a
paradigm for investigating human ocular genetics. Eye 31, 68–86 (2017).
118. Howe, K. et al. The zebrafish reference genome sequence and its relationship to the
human genome. Nature 496, 498–503 (2013).
119. Wang, H. et al. Portrait of glial scar in neurological diseases. Int J Immunopathol
Pharmacol 31, (2018).
120. Lundkvist, A. et al. Under stress, the absence of intermediate filaments from Müller cells
in the retina has structural and functional consequences. J. Cell. Sci. 117, 3481–3488 (2004).
121. Lu, Y.-B. et al. Biomechanical properties of retinal glial cells: comparative and
developmental data. Exp. Eye Res. 113, 60–65 (2013).
122. Lewis, G. P. & Fisher, S. K. Up-regulation of glial fibrillary acidic protein in response to
retinal injury: its potential role in glial remodeling and a comparison to vimentin expression.
Int. Rev. Cytol. 230, 263–290 (2003).
157
123. Nagashima, M., Barthel, L. K. & Raymond, P. A. A self-renewing division of zebrafish
Müller glial cells generates neuronal progenitors that require N-cadherin to regenerate
retinal neurons. Development 140, 4510–4521 (2013).
124. Bernardos, R. L., Barthel, L. K., Meyers, J. R. & Raymond, P. A. Late-Stage Neuronal
Progenitors in the Retina Are Radial Müller Glia That Function as Retinal Stem Cells. J.
Neurosci. 27, 7028–7040 (2007).
125. Powell, C., Cornblath, E., Elsaeidi, F., Wan, J. & Goldman, D. Zebrafish Müller glia-derived
progenitors are multipotent, exhibit proliferative biases and regenerate excess neurons.
Scientific Reports 6, 24851 (2016).
126. Fausett, B. V. & Goldman, D. A role for alpha1 tubulin-expressing Müller glia in
regeneration of the injured zebrafish retina. J. Neurosci. 26, 6303–6313 (2006).
127. Fimbel, S. M., Montgomery, J. E., Burket, C. T. & Hyde, D. R. Regeneration of Inner
Retinal Neurons after Intravitreal Injection of Ouabain in Zebrafish. J. Neurosci. 27, 1712–
1724 (2007).
128. Maier, W. & Wolburg, H. Regeneration of the goldfish retina after exposure to different
doses of ouabain. Cell Tissue Res. 202, 99–118 (1979).
129. McGinn, T. E. et al. Rewiring the Regenerated Zebrafish Retina: Reemergence of Bipolar
Neurons and Cone-Bipolar Circuitry Following an Inner Retinal Lesion. Front. Cell Dev. Biol. 7,
(2019).
130. Wu, D. M. et al. Cones regenerate from retinal stem cells sequestered in the inner
nuclear layer of adult goldfish retina. Invest. Ophthalmol. Vis. Sci. 42, 2115–2124 (2001).
131. DiCicco, R. M. et al. Retinal regeneration following OCT-guided laser injury in zebrafish.
Invest. Ophthalmol. Vis. Sci. 55, 6281–6288 (2014).
158
132. Vihtelic, T. S. & Hyde, D. R. Light-induced rod and cone cell death and regeneration in
the adult albino zebrafish (Danio rerio) retina. Journal of Neurobiology 44, 289–307 (2000).
133. Otteson, D. C., D’Costa, A. R. & Hitchcock, P. F. Putative stem cells and the lineage of rod
photoreceptors in the mature retina of the goldfish. Dev. Biol. 232, 62–76 (2001).
134. Raymond, P. A. & Rivlin, P. K. Germinal cells in the goldfish retina that produce rod
photoreceptors. Dev. Biol. 122, 120–138 (1987).
135. Cameron, D. A. Cellular proliferation and neurogenesis in the injured retina of adult
zebrafish. Vis. Neurosci. 17, 789–797 (2000).
136. Wan, J. & Goldman, D. Retina regeneration in zebrafish. Curr Opin Genet Dev 40, 41–47
(2016).
137. Rao, M. B., Didiano, D. & Patton, J. G. Neurotransmitter-Regulated Regeneration in the
Zebrafish Retina. Stem Cell Reports 8, 831–842 (2017).
138. Wang, M., Ma, W., Zhao, L., Fariss, R. N. & Wong, W. T. Adaptive Müller cell responses
to microglial activation mediate neuroprotection and coordinate inflammation in the retina.
Journal of Neuroinflammation 8, 173 (2011).
139. Conedera, F. M., Pousa, A. M. Q., Mercader, N., Tschopp, M. & Enzmann, V. Retinal
microglia signaling affects Müller cell behavior in the zebrafish following laser injury
induction. Glia 67, 1150–1166 (2019).
140. Sifuentes, C. J., Kim, J.-W., Swaroop, A. & Raymond, P. A. Rapid, Dynamic Activation of
Müller Glial Stem Cell Responses in Zebrafish. Invest Ophthalmol Vis Sci 57, 5148–5160
(2016).
141. Wan, J. & Goldman, D. Opposing actions of Fgf8a on Notch signaling distinguish two
Muller glial cell populations that contribute to retina growth and regeneration. Cell Rep 19,
849–862 (2017).
159
142. Wan, J., Ramachandran, R. & Goldman, D. HB-EGF is necessary and sufficient for Müller
glia dedifferentiation and retina regeneration. Dev Cell 22, 334–347 (2012).
143. Todd, L., Volkov, L. I., Zelinka, C., Squires, N. & Fischer, A. J. Heparin-binding EGF-like
growth factor (HB-EGF) stimulates the proliferation of Müller glia-derived progenitor cells in
avian and murine retinas. Mol. Cell. Neurosci. 69, 54–64 (2015).
144. Wan, J., Zhao, X.-F., Vojtek, A. & Goldman, D. Retinal injury, growth factors, and
cytokines converge on β-catenin and pStat3 signaling to stimulate retina regeneration. Cell
Rep 9, 285–297 (2014).
145. Kassen, S. C. et al. CNTF induces photoreceptor neuroprotection and Müller glial cell
proliferation through two different signaling pathways in the adult zebrafish retina. Exp. Eye
Res. 88, 1051–1064 (2009).
146. Wang, Y., Smith, S. B., Ogilvie, J. M., McCool, D. J. & Sarthy, V. Ciliary neurotrophic factor
induces glial fibrillary acidic protein in retinal Müller cells through the JAK/STAT signal
transduction pathway. Curr. Eye Res. 24, 305–312 (2002).
147. Meyers, J. R. et al. β-catenin/Wnt signaling controls progenitor fate in the developing
and regenerating zebrafish retina. Neural Dev 7, 30 (2012).
148. Ramachandran, R., Zhao, X.-F. & Goldman, D. Ascl1a/Dkk/beta-catenin signaling
pathway is necessary and glycogen synthase kinase-3beta inhibition is sufficient for zebrafish
retina regeneration. Proc. Natl. Acad. Sci. U.S.A. 108, 15858–15863 (2011).
149. Zhao, X.-F. et al. Leptin and IL-6 family cytokines synergize to stimulate Müller glia
reprogramming and retina regeneration. Cell Rep 9, 272–284 (2014).
150. Coorey, N. J., Shen, W., Zhu, L. & Gillies, M. C. Differential Expression of IL-6/gp130
Cytokines, Jak-STAT Signaling and Neuroprotection After Müller Cell Ablation in a Transgenic
Mouse Model. Invest. Ophthalmol. Vis. Sci. 56, 2151–2161 (2015).
160
151. Xue, W. et al. Ciliary Neurotrophic Factor Induces Genes Associated with Inflammation
and Gliosis in the Retina: A Gene Profiling Study of Flow-Sorted, Müller Cells. PLoS One 6,
(2011).
152. Elsaeidi, F. et al. Notch Suppression Collaborates with Ascl1 and Lin28 to Unleash a
Regenerative Response in Fish Retina, But Not in Mice. J. Neurosci. 38, 2246–2261 (2018).
153. Fausett, B. V., Gumerson, J. D. & Goldman, D. The proneural bHLH gene ascl1a is
required for retina regeneration. J Neurosci 28, 1109–1117 (2008).
154. Allan, K., DiCicco, R., Ramos, M., Asosingh, K. & Yuan, A. Preparing a Single Cell
Suspension from Zebrafish Retinal Tissue for Flow Cytometric Cell Sorting of Müller Glia.
Cytometry Part A n/a,.
155. McKinnon, K. M. Flow Cytometry: An Overview. Curr Protoc Immunol 120, 5.1.1-5.1.11
(2018).
156. Davies, D. Cell Sorting by Flow Cytometry. in Flow Cytometry: Principles and Applications
(ed. Macey, M. G.) 257–276 (Humana Press, 2007). doi:10.1007/978-1-59745-451-3_11.
157. Sun, C., Mitchell, D. M. & Stenkamp, D. L. Isolation of photoreceptors from mature,
developing, and regenerated zebrafish retinas, and of microglia/macrophages from
regenerating zebrafish retinas. Experimental Eye Research 177, 130–144 (2018).
158. Sun, C., Galicia, C. & Stenkamp, D. L. Transcripts within rod photoreceptors of the
Zebrafish retina. BMC Genomics 19, 127 (2018).
159. Qin, Z., Barthel, L. K. & Raymond, P. A. Genetic evidence for shared mechanisms of
epimorphic regeneration in zebrafish. Proc. Natl. Acad. Sci. U.S.A. 106, 9310–9315 (2009).
160. Glaviano, A. et al. A method for isolation of cone photoreceptors from adult zebrafish
retinae. BMC Neurosci 17, (2016).
161
161. Hoang, T. et al. Cross-species transcriptomic and epigenomic analysis reveals key
regulators of injury response and neuronal regeneration in vertebrate retinas. bioRxiv 717876
(2019) doi:10.1101/717876.
162. Ramachandran, R., Fausett, B. V. & Goldman, D. Ascl1a regulates Muller glia
dedifferentiation and retinal regeneration through a Lin-28-dependent, let-7 microRNA
signalling pathway. Nat Cell Biol 12, 1101–1107 (2010).
163. Stenkamp, D. L. Neurogenesis in the Fish Retina. Int Rev Cytol 259, 173–224 (2007).
164. Thummel, R. et al. Characterization of Müller glia and neuronal progenitors during adult
zebrafish retinal regeneration. Exp Eye Res 87, 433–444 (2008).
165. Goldman, D. Muller glial cell reprogramming and retina regeneration. Nat Rev Neurosci
15, 431–442 (2014).
166. Reichard, A. & Asosingh, K. Best Practices for Preparing a Single Cell Suspension from
Solid Tissues for Flow Cytometry. Cytometry Part A 95, 219–226 (2019).
167. Hussain, R. Z. et al. Defining standard enzymatic dissociation methods for individual
brains and spinal cords in EAE. Neurol Neuroimmunol Neuroinflamm 5, (2018).
168. Huettner, J. E. & Baughman, R. W. Primary culture of identified neurons from the visual
cortex of postnatal rats. J. Neurosci. 6, 3044–3060 (1986).
169. Bader, C. R., MacLeish, P. R. & Schwartz, E. A. Responses to light of solitary rod
photoreceptors isolated from tiger salamander retina. Proc. Natl. Acad. Sci. U.S.A. 75, 3507–
3511 (1978).
170. Townes-Anderson, E., MacLeish, P. R. & Raviola, E. Rod cells dissociated from mature
salamander retina: ultrastructure and uptake of horseradish peroxidase. J. Cell Biol. 100, 175–
188 (1985).
162
171. Lam, D. M. K. Biosynthesis of Acetylcholine in Turtle Photoreceptors. Proc Natl Acad Sci
U S A 69, 1987–1991 (1972).
172. Papain Dissociation System - Worthington Enzyme Manual. http://www.worthington-
biochem.com/PDS/.
173. Wu, S. M. Synaptic Organization of the Vertebrate Retina: General Principles and
Species-Specific Variations: The Friedenwald Lecture. Invest. Ophthalmol. Vis. Sci. 51, 1264–
1274 (2010).
174. Hippert, C. et al. Müller Glia Activation in Response to Inherited Retinal Degeneration Is
Highly Varied and Disease-Specific. PLOS ONE 10, e0120415 (2015).
175. Pasha, S. P. B. S. et al. Retinal cell death dependent reactive proliferative gliosis in the
mouse retina. Scientific Reports 7, 9517 (2017).
176. Gestri, G., Link, B. A. & Neuhauss, S. C. The Visual System of Zebrafish and its Use to
Model Human Ocular Diseases. Dev Neurobiol 72, 302–327 (2012).
177. Avanesov, A. & Malicki, J. Analysis of the Retina in the Zebrafish Model. Methods Cell
Biol 100, 153–204 (2010).
178. Reinhard, J., Joachim, S. C. & Faissner, A. Extracellular matrix remodeling during retinal
development. Experimental Eye Research 133, 132–140 (2015).
179. Al-Ubaidi, M. R., Naash, M. I. & Conley, S. M. A Perspective on the Role of the
Extracellular Matrix in Progressive Retinal Degenerative Disorders. Invest Ophthalmol Vis Sci
54, 8119–8124 (2013).
180. Taylor, L., Arnér, K., Engelsberg, K. & Ghosh, F. Scaffolding the retina: the interstitial
extracellular matrix during rat retinal development. Int. J. Dev. Neurosci. 42, 46–58 (2015).
181. Hunter, D. D. & Brunken, W. J. Beta 2 laminins modulate neuronal phenotype in the rat
retina. Mol. Cell. Neurosci. 10, 7–15 (1997).
163
182. Perez, R. G. & Halfter, W. Tenascin in the Developing Chick Visual System: Distribution
and Potential Role as a Modulator of Retinal Axon Growth. Developmental Biology 156, 278–
292 (1993).
183. McAdams, B. D. & McLoon, S. C. Expression of chondroitin sulfate and keratan sulfate
proteoglycans in the path of growing retinal axons in the developing chick. Journal of
Comparative Neurology 352, 594–606 (1995).
184. Hodel, C., Neuhauss, S. C. F. & Biehlmaier, O. Time course and development of light
adaptation processes in the outer zebrafish retina. The Anatomical Record Part A: Discoveries
in Molecular, Cellular, and Evolutionary Biology 288A, 653–662 (2006).
185. Bishop, P. N. The role of extracellular matrix in retinal vascular development and
preretinal neovascularization. Exp. Eye Res. 133, 30–36 (2015).
186. Vecino, E., Heller, J. P., Veiga-Crespo, P., Martin, K. R. & Fawcett, J. W. Influence of
extracellular matrix components on the expression of integrins and regeneration of adult
retinal ganglion cells. PLoS ONE 10, e0125250 (2015).
187. Bryan, C. D., Chien, C.-B. & Kwan, K. M. Loss of laminin alpha 1 results in multiple
structural defects and divergent effects on adhesion during vertebrate optic cup
morphogenesis. Dev Biol 416, 324–337 (2016).
188. Ali, M. A. [Retinomotor response: characteristics and mechanisms]. Vision Res. 11,
1225–1288 (1971).
189. Tennant, J. R. EVALUATION OF THE TRYPAN BLUE TECHNIQUE FOR DETERMINATION OF
CELL VIABILITY. Transplantation 2, 685–694 (1964).
190. Martinez-De Luna, R. I. et al. Müller glia reactivity follows retinal injury despite the
absence of the glial fibrillary acidic protein gene in Xenopus. Developmental Biology 426,
219–235 (2017).
164
191. Raviola, E. & Gilula, N. B. Gap Junctions between Photoreceptor Cells in the Vertebrate
Retina. PNAS 70, 1677–1681 (1973).
192. Goodenough, D. A. & Paul, D. L. Gap Junctions. Cold Spring Harb Perspect Biol 1, (2009).
193. Roy, K., Kumar, S. & Bloomfield, S. A. Gap junctional coupling between retinal amacrine
and ganglion cells underlies coherent activity integral to global object perception. Proc Natl
Acad Sci U S A 114, E10484–E10493 (2017).
194. Klaassen, L. J., de Graaff, W., van Asselt, J. B., Klooster, J. & Kamermans, M. Specific
connectivity between photoreceptors and horizontal cells in the zebrafish retina. J.
Neurophysiol. 116, 2799–2814 (2016).
195. Kántor, O. et al. Bipolar cell gap junctions serve major signaling pathways in the human
retina. Brain Struct Funct 222, 2603–2624 (2017).
196. Kong, Y., Naggert, J. K. & Nishina, P. M. The Impact of Adherens and Tight Junctions on
Physiological Function and Pathological Changes in the Retina. in Retinal Degenerative
Diseases (eds. Ash, J. D. et al.) 545–551 (Springer International Publishing, 2018).
197. Alves, C. H., Pellissier, L. P. & Wijnholds, J. The CRB1 and adherens junction complex
proteins in retinal development and maintenance. Progress in Retinal and Eye Research 40,
35–52 (2014).
198. Tolun, G. et al. Paired octamer rings of retinoschisin suggest a junctional model for cell-
cell adhesion in the retina. Proc. Natl. Acad. Sci. U.S.A. 113, 5287–5292 (2016).
199. Reichenbach, A. et al. The structure of rabbit retinal Müller (glial) cells is adapted to the
surrounding retinal layers. Anatomy and Embryology 180, 71–79 (1989).
200. Skytt, D. M. et al. Glia-Neuron Interactions in the Retina Can Be Studied in Cocultures of
Müller Cells and Retinal Ganglion Cells. Biomed Res Int 2016, (2016).
165
201. Kassen, S. C. et al. Time course analysis of gene expression during light-induced
photoreceptor cell death and regeneration in albino zebrafish. Developmental Neurobiology
67, 1009–1031 (2007).
202. Lahne, M. & Hyde, D. R. Interkinetic Nuclear Migration in the Regenerating Retina. Adv.
Exp. Med. Biol. 854, 587–593 (2016).
203. Barros, C. S., Franco, S. J. & Müller, U. Extracellular Matrix: Functions in the Nervous
System. Cold Spring Harb Perspect Biol 3, (2011).
204. Muncie, J. M. & Weaver, V. M. Chapter One - The Physical and Biochemical Properties of
the Extracellular Matrix Regulate Cell Fate. in Current Topics in Developmental Biology (eds.
Litscher, E. S. & Wassarman, P. M.) vol. 130 1–37 (Academic Press, 2018).
205. Gattazzo, F., Urciuolo, A. & Bonaldo, P. Extracellular matrix: A dynamic
microenvironment for stem cell niche. Biochim Biophys Acta 1840, 2506–2519 (2014).
206. Rolls, A., Shechter, R. & Schwartz, M. The bright side of the glial scar in CNS repair. Nat.
Rev. Neurosci. 10, 235–241 (2009).
207. Walters, N. J. & Gentleman, E. Evolving insights in cell-matrix interactions: elucidating
how non-soluble properties of the extracellular niche direct stem cell fate. Acta Biomater 11,
3–16 (2015).
208. Herder, C. et al. ArhGEF18 regulates RhoA-Rock2 signaling to maintain neuro-epithelial
apico-basal polarity and proliferation. Development 140, 2787–2797 (2013).
209. McBeath, R., Pirone, D. M., Nelson, C. M., Bhadriraju, K. & Chen, C. S. Cell Shape,
Cytoskeletal Tension, and RhoA Regulate Stem Cell Lineage Commitment. Developmental Cell
6, 483–495 (2004).
210. Amano, M. et al. Phosphorylation and Activation of Myosin by Rho-associated Kinase
(Rho-kinase). J. Biol. Chem. 271, 20246–20249 (1996).
166
211. Pietri, T. et al. Conditional beta1-integrin gene deletion in neural crest cells causes
severe developmental alterations of the peripheral nervous system. Development 131, 3871–
3883 (2004).
212. Jacques, T. S. et al. Neural precursor cell chain migration and division are regulated
through different beta1 integrins. Development 125, 3167–3177 (1998).
213. de Lucas, B., Pérez, L. M. & Gálvez, B. G. Importance and regulation of adult stem cell
migration. J Cell Mol Med 22, 746–754 (2018).
214. Bian, S. Cell Adhesion Molecules in Neural Stem Cell and Stem Cell- Based Therapy for
Neural Disorders. Neural Stem Cells - New Perspectives (2013) doi:10.5772/55136.
215. Zhang, Y., Sivasankar, S., Nelson, W. J. & Chu, S. Resolving cadherin interactions and
binding cooperativity at the single-molecule level. Proc. Natl. Acad. Sci. U.S.A. 106, 109–114
(2009).
216. Erez, N., Bershadsky, A. & Geiger, B. Signaling from adherens-type junctions. Eur. J. Cell
Biol. 84, 235–244 (2005).
217. Cai, D. et al. Mechanical feedback through E-cadherin promotes direction sensing during
collective cell migration. Cell 157, 1146–1159 (2014).
218. Xu, J.-C. et al. Transplanted L1 expressing radial glia and astrocytes enhance recovery
after spinal cord injury. J. Neurotrauma 28, 1921–1937 (2011).
219. Wang, Y. et al. The promotion of neural progenitor cells proliferation by aligned and
randomly oriented collagen nanofibers through β1 integrin/MAPK signaling pathway.
Biomaterials 32, 6737–6744 (2011).
220. Stepniak, E., Radice, G. L. & Vasioukhin, V. Adhesive and Signaling Functions of
Cadherins and Catenins in Vertebrate Development. Cold Spring Harb Perspect Biol 1,
a002949 (2009).
167
221. von Lersner, A., Droesen, L. & Zijlstra, A. Modulation of cell adhesion and migration
through regulation of the immunoglobulin superfamily member ALCAM/CD166. Clin. Exp.
Metastasis 36, 87–95 (2019).
222. Degen, W. G. et al. MEMD, a new cell adhesion molecule in metastasizing human
melanoma cell lines, is identical to ALCAM (activated leukocyte cell adhesion molecule). Am J
Pathol 152, 805–813 (1998).
223. Diekmann, H. & Stuermer, C. A. O. Zebrafish neurolin-a and -b, orthologs of ALCAM, are
involved in retinal ganglion cell differentiation and retinal axon pathfinding. J. Comp. Neurol.
513, 38–50 (2009).
224. Choudhry, P., Joshi, D., Funke, B. & Trede, N. Alcama mediates Edn1 signaling during
zebrafish cartilage morphogenesis. Dev Biol 349, 483–493 (2011).
225. van Kempen, L. C. et al. Molecular basis for the homophilic activated leukocyte cell
adhesion molecule (ALCAM)-ALCAM interaction. J. Biol. Chem. 276, 25783–25790 (2001).
226. Hansen, A. G., Swart, G. W. & Zijlstra, A. ALCAM. AFCS Nat Mol Pages 2011, (2011).
227. Heterophilic interactions of DM-GRASP: GRASP-NgCAM interactions involved in neurite
extension. J Cell Biol 133, 657–666 (1996).
228. Tomita, K., van Bokhoven, A., Jansen, C. F., Bussemakers, M. J. & Schalken, J. A.
Coordinate recruitment of E-cadherin and ALCAM to cell-cell contacts by alpha-catenin.
Biochem. Biophys. Res. Commun. 267, 870–874 (2000).
229. Heffron, D. S. & Golden, J. A. DM-GRASP Is Necessary for Nonradial Cell Migration during
Chick Diencephalic Development. J. Neurosci. 20, 2287–2294 (2000).
230. Tudor, C. et al. Syntenin-1 and ezrin proteins link activated leukocyte cell adhesion
molecule to the actin cytoskeleton. J. Biol. Chem. 289, 13445–13460 (2014).
168
231. Gilsanz, A. et al. ALCAM/CD166 adhesive function is regulated by the tetraspanin CD9.
Cell. Mol. Life Sci. 70, 475–493 (2013).
232. Thelen, K., Georg, T., Bertuch, S., Zelina, P. & Pollerberg, G. E. Ubiquitination and
endocytosis of cell adhesion molecule DM-GRASP regulate its cell surface presence and affect
its role for axon navigation. J. Biol. Chem. 283, 32792–32801 (2008).
233. Laessing, U. & Stuermer, C. A. Spatiotemporal pattern of retinal ganglion cell
differentiation revealed by the expression of neurolin in embryonic zebrafish. J. Neurobiol.
29, 65–74 (1996).
234. Ott, H., Diekmann, H., Stuermer, C. A. O. & Bastmeyer, M. Function of Neurolin (DM-
GRASP/SC-1) in Guidance of Motor Axons during Zebrafish Development. Developmental
Biology 235, 86–97 (2001).
235. Weiner, J. A. et al. Axon fasciculation defects and retinal dysplasias in mice lacking the
immunoglobulin superfamily adhesion molecule BEN/ALCAM/SC1. Mol. Cell. Neurosci. 27,
59–69 (2004).
236. Thummel, R., Bailey, T. J. & Hyde, D. R. In vivo Electroporation of Morpholinos into the
Adult Zebrafish Retina. JoVE (Journal of Visualized Experiments) e3603–e3603 (2011)
doi:10.3791/3603.
237. ZFIN.
https://zfin.org/search?q=&fq=category%3A%22Mutation+%2F+Tg%22&fq=xref%3A%22ZDB-
GENE-990415-30%22.
238. El-Brolosy, M. A. et al. Genetic compensation triggered by mutant mRNA degradation.
Nature 568, 193–197 (2019).
239. Bedell, V. M., Westcot, S. E. & Ekker, S. C. Lessons from morpholino-based screening in
zebrafish. Brief Funct Genomics 10, 181–188 (2011).
169
240. Zhu, L. et al. Characterization of canonical Wnt signalling changes after induced
disruption of Müller cell in murine retina. Exp. Eye Res. 175, 173–180 (2018).
241. Angbohang, A. et al. Downregulation of the Canonical WNT Signaling Pathway by TGFβ1
Inhibits Photoreceptor Differentiation of Adult Human Müller Glia with Stem Cell
Characteristics. Stem Cells Dev 25, 1–12 (2016).
242. Sugimura, R. et al. Noncanonical Wnt Signaling Maintains Hematopoietic Stem Cells in
the Niche. Cell 150, 351–365 (2012).
243. Gallina, D., Palazzo, I., Steffenson, L., Todd, L. & Fischer, A. J. Wnt/β-catenin-signaling
and the formation of Müller glia-derived progenitors in the chick retina. Dev Neurobiol 76,
983–1002 (2016).
244. Tian, X. et al. E-Cadherin/β-Catenin Complex and the Epithelial Barrier. J Biomed
Biotechnol 2011, (2011).
245. Nomura-Komoike, K., Saitoh, F. & Fujieda, H. Phosphatidylserine recognition and Rac1
activation are required for Müller glia proliferation, gliosis and phagocytosis after retinal
injury. Scientific Reports 10, 1–11 (2020).
246. Marlow, F., Topczewski, J., Sepich, D. & Solnica-Krezel, L. Zebrafish Rho kinase 2 acts
downstream of Wnt11 to mediate cell polarity and effective convergence and extension
movements. Curr. Biol. 12, 876–884 (2002).
247. Hoang, T. et al. Gene regulatory networks controlling vertebrate retinal regeneration.
Science (2020) doi:10.1126/science.abb8598.
248. Silva, N. J. et al. Inflammation and matrix metalloproteinase 9 (Mmp-9) regulate
photoreceptor regeneration in adult zebrafish. Glia n/a,.
249. Reinhard, J. et al. Ischemic injury leads to extracellular matrix alterations in retina and
optic nerve. Sci Rep 7, 43470 (2017).
170
250. Besser, M., Jagatheaswaran, M., Reinhard, J., Schaffelke, P. & Faissner, A. Tenascin C
regulates proliferation and differentiation processes during embryonic retinogenesis and
modulates the de-differentiation capacity of Müller glia by influencing growth factor
responsiveness and the extracellular matrix compartment. Dev. Biol. 369, 163–176 (2012).
251. Garcion, E., Halilagic, A., Faissner, A. & ffrench-Constant, C. Generation of an
environmental niche for neural stem cell development by the extracellular matrix molecule
tenascin C. Development 131, 3423–3432 (2004).
252. Husmann, K., Carbonetto, S. & Schachner, M. Distinct sites on tenascin-C mediate
repellent or adhesive interactions with different neuronal cell types. Cell Adhes. Commun. 3,
293–310 (1995).
253. Ichijo, H. Proteoglycans as cues for axonal guidance in formation of retinotectal or
retinocollicular projections. Mol. Neurobiol. 30, 23–33 (2004).
254. Busch, S. A. & Silver, J. The role of extracellular matrix in CNS regeneration. Current
Opinion in Neurobiology 17, 120–127 (2007).
255. Tucker, B., Klassen, H., Yang, L., Chen, D. F. & Young, M. J. Elevated MMP Expression in
the MRL Mouse Retina Creates a Permissive Environment for Retinal Regeneration. Invest.
Ophthalmol. Vis. Sci. 49, 1686–1695 (2008).
256. Ren, T., van der Merwe, Y. & Steketee, M. B. Developing Extracellular Matrix Technology
to Treat Retinal or Optic Nerve Injury(1,2,3). eNeuro 2, (2015).
257. Wang, M. & Wong, W. T. Microglia-Müller Cell Interactions in the Retina. Adv Exp Med
Biol 801, 333–338 (2014).
258. Conedera, F. M., Pousa, A. M. Q., Mercader, N., Tschopp, M. & Enzmann, V. Retinal
microglia signaling affects Müller cell behavior in the zebrafish following laser injury
induction. Glia 67, 1150–1166 (2019).
171
259. White, D. T. et al. Immunomodulation-accelerated neuronal regeneration following
selective rod photoreceptor cell ablation in the zebrafish retina. Proc. Natl. Acad. Sci. U.S.A.
114, E3719–E3728 (2017).
260. Fischer, A. J., Zelinka, C., Gallina, D., Scott, M. A. & Todd, L. Reactive microglia and
macrophage facilitate the formation of Müller glia-derived retinal progenitors. Glia 62, 1608–
1628 (2014).
261. Smith, J. A., Das, A., Ray, S. K. & Banik, N. L. Role of pro-inflammatory cytokines released
from microglia in neurodegenerative diseases. Brain Res. Bull. 87, 10–20 (2012).
262. Zimmerman, A. W. et al. Long-term engagement of CD6 and ALCAM is essential for T-
cell proliferation induced by dendritic cells. Blood 107, 3212–3220 (2006).
263. Cloning, mapping, and characterization of activated leukocyte-cell adhesion molecule
(ALCAM), a CD6 ligand. J Exp Med 181, 2213–2220 (1995).
264. Cayrol, R. et al. Activated leukocyte cell adhesion molecule promotes leukocyte
trafficking into the central nervous system. Nat. Immunol. 9, 137–145 (2008).
265. Kurki, P., Vanderlaan, M., Dolbeare, F., Gray, J. & Tan, E. M. Expression of proliferating
cell nuclear antigen (PCNA)/cyclin during the cell cycle. Exp Cell Res 166, 209–219 (1986).
266. Singh, C. et al. Hyperoxia induces glutamine-fuelled anaplerosis in retinal Müller cells.
Nature Communications 11, 1–11 (2020).
172