Zebrafish may hold the secret to regenerating damaged retinas in humans

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The tiny zebrafish may hold the secret to regenerating damaged retinas in humans, Vanderbilt University researchers reported last week in the journal Cell Reports.

Currently there are few effective treatments for retinal degenerative diseases such as age-related macular degeneration (AMD), the most common cause of blindness in millions of Americans over age 55.

The prognosis is equally dim for another 100,000 Americans with retinitis pigmentosa. Many of them are legally blind by age 40.

Mammals cannot spontaneously regenerate retinal neurons that are lost or damaged by disease.

But in bony fish like the 1-inch-long zebrafish, which is widely used as a “model organism” in biological and medical research studies, retinal damage triggers a spontaneous regenerative response that restores both retinal structure and function.

Since the cells and structure of the retina are highly conserved among vertebrates, from fish to humans, understanding how zebrafish regenerate damaged retinas could lead to new ways to enhance retinal regeneration in people.

One key factor explored by James Patton, Ph.D., and colleagues in the Department of Biological Sciences at Vanderbilt University is a microRNA, miR-216a, which regulates the expression of Dot1l, an enzyme involved in regulating gene expression.

Suppressing miR-216a, they showed, initiates the differentiation and proliferation of Müller glia (MG), the source of regenerated neurons in the zebrafish retina, in part by releasing inhibition of Dot1l.

A next step is to test whether suppressing miR-216a can induce MG reprogramming and proliferation in mammals.

“Müller glia constitute an adult stem cell in the zebrafish retina and our goal is to identify pathways and genes that could be activated to induce similar behavior in the human retina,” said Patton, Stevenson Professor of Biological Sciences and director of the Interdisciplinary Graduate Program at Vanderbilt.


Similar to many organs, the eye is structurally well-conserved between zebrafish and mammals. For example, the eyes of zebrafish have the same gross structure as human and other mammalian eyes, and contain a cornea, lens, vitreous, retina, pigment epithelium, choroid and sclera (Figure ​Figure1A).

Furthermore, the development of the eye during embryogenesis is also conserved, and complimentary work in zebrafish, mice and other species has helped to delineate the key developmental events in eye morphogenesis across vertebrates (Bibliowicz et al., 2011Stenkamp, 2015).

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FIGURE 1
Structure of the zebrafish eye and retina. (A) Anatomy of the zebrafish eye: DIC image of a cryosection from a 6 days post-fertilization (6 dpf) larval eye highlighting the main structures of the vertebrate eye including cornea, lens, retina, and retinal pigment epithelium (RPE). (B) The larval retina is organized into highly structured layers: Overlay of a DIC image and the fluorescent nuclear marker DAPI of a cryosection from a 6 dpf larva showing the different retinal layers, including the outer nuclear layer (ONL) which contains the cell bodies of photoreceptors (rods and cones). Photoreceptors make synapses in the outer plexiform layer (OPL) with bipolar and horizontal cells. The inner nuclear layer (INL) contains the cell bodies of horizontal, bipolar and amacrine cells, while the ganglion cell layer (GCL) contains the cell bodies of retinal ganglion cells (RGC). Bipolar cells provide excitatory synaptic input to RGC in the inner plexiform layer (IPL), while amacrine cells modulate this input both pre- and post-synaptically. (C) The adult retina retains the same layered structure: Overlay of a DIC image and DAPI of a cryosection from an adult zebrafish. (D) The zebrafish retina contains 4 subtypes of cones: DIC image of a cryosection from an adult zebrafish showing the short-single or ultraviolet-wavelength sensitive cones (UV-cones), the long-single or short-wavelength sensitive cones (S-cones), and the double cones which correspond to the middle- and long-wavelength sensitive cones (M- and L-cones). (E) Mosaic arrangement of zebrafish cone photoreceptors: Confocal image of a whole-mounted retina of a double-reporter transgenic lines to identify UV-cones [Tg(sws1:GFP)kj9, magenta] and S-cones [tg(sws2:mCherry)ua3011, blue] overlayed with DAPI (gray), allowing the identification of the nuclei of M- and L-cones between the rows of UV- and S-cones. (F) Diagram of the vertebrate retina and the retinal cells. Inset highlights the synapse between cones and horizontal and bipolar cells, where the cone synaptic terminal contains synaptic vesicles (white) attached to the synaptic ribbon (black). In close apposition to the ribbon, the dendrites on On-bipolar cells (On-BCs) invaginate into the synaptic terminal and are flanked by two horizontal cell (HC) processes. Off-bipolar cells make more basal contacts in close proximity but not apposed to the synaptic ribbon.

Within the eye, the retina is of particular interest because it is the site of sensory detection and damage to the retina results in vision loss.

The vertebrate retina is a highly structured neuronal tissue that lines the back of the eye. It is responsible for both the detection and processing of visual information, before it is relayed to higher-order visual centers.

To achieve this, the retina is equipped with a variety of neurons that are arranged into three nuclear layers and project into two synaptic layers (Figures 1B,C).

Within these layers, retinal neurons assemble into multiple, distinct circuits that encode different aspects of the visual information (Gollisch and Meister, 2010).

The encoding of visual information starts when light is detected by the rod and cone photoreceptors.

The highly sensitive rods are mainly used during dim-lighting conditions, while the more adaptable but less sensitive cones function from dawn until dusk.

The retina of the nocturnal rodents commonly used in research like mice and rats is rod dominated (97% rods and 3% cones) (Carter-Dawson Louvenia and Lavail Matthew, 1979), as is the peripheral human retina. In contrast, zebrafish have a cone-dominated retina (∼40% rods and ∼60% cones) (Fadool, 2003), similar to the central human retina, which provides high-acuity vision, and is essential for most day-to-day visual tasks.

Therefore, the zebrafish retina is uniquely positioned to understand the molecular mechanisms relevant to development and regeneration of the photoreceptors that are most relevant for human vision.

The zebrafish retina contains four different cone photoreceptor subtypes (UV-, S-, M-, and L-cones).

Each subtype is defined by specific opsin expression that confers a particular wavelength-sensitivity, and morphology (short or long, and single or paired with another cone type) (Figures 1D,E). UV-cones express sws1, an opsin with peak sensitivity (λmax) in the ultraviolet range (λmax = 354 nm), and are short-single cones morphologically. S-cones express sws2, with peak sensitivity at short wavelengths (λmax = 416 nm), and are long-single cones morphologically. M-cones express opsins of the Rh2 class, which have undergone tandem quadruplications (Rh2-1 to Rh2-4), with peak sensitivities at mid wavelengths (λmax = 467 nm, 476 nm, 488 nm, and 505 nm respectively). L-cones express one of two tandemly duplicated opsin genes from the lws class, with peak sensitivities at longer wavelengths (λmax = 548 nm and 558 nm). M- and L-cones are morphologically arranged as a double cone, where the L-cone is the long (or principal) member of the pair and the M-cone is the short (or accessory) member (Raymond et al., 1996Vihtelic et al., 1999Chinen et al., 2003).

In both rod and cone photoreceptors, light triggers the activation of opsins followed by the rest of the phototransduction cascade.

In zebrafish and in mammals, this cascade ultimately leads to changes in photoreceptor membrane potential, and to modulation of neurotransmitter release in the synaptic terminal.

Visual information is then directly transmitted from photoreceptors to several subtypes of horizontal (inhibitory interneurons that locally modulate photoreceptor synaptic output) and bipolar cells (glutamatergic neurons that transmit light signals into the next processing layer). Information is further processed in the next synaptic layer, where bipolar cells (BC) provide excitation to ganglion cells (glutamatergic spiking neurons), while amacrine cells (local interneurons) provide modulation pre- and/or post-synaptically.

The axons of the retinal ganglion cells (or RGCs) form the optic nerve, and relay the pre-processed visual information to central targets in the brain. Additionally, the retina contains microglia (resident immune cells located primarily in the synaptic layers) and two types of true glial cells: Müller cells (a type of radial glia) and astrocytes (associated with axons of RGCs) (Figure ​Figure1F).

The photoreceptors also closely associate with the retinal pigment epithelium (RPE), which provides structural, trophic and metabolic support and is directly involved in the recycling of opsins.

Therapies for RD

In humans, photoreceptor loss in RD is permanent (regardless of their diverse causes and speed of progression) and therefore remain largely untreatable and lead to progressive loss of vision and ultimately blindness. In early stages of RD in humans, current treatments include neuroprotective agents (Trifunovic et al., 2012) and antibody therapies in cases where the underlying mutation is well characterized (Lazic and Gabric, 2007). Unfortunately, these treatments only slow down the progression of disease and have variable outcomes (Pardue and Allen, 2018). In addition to these treatments, gene therapy has been used to improve vision in patients with LCA caused by mutations in RPE65, an RPE-specific protein involved in the recycling of retinoids (Cideciyan, 2010Jacobson et al., 2012), but the improvement may not be long-lasting (Jacobson et al., 2015).

In late stages of RD in humans, when there is widespread photoreceptor loss, two distinct approaches for treatment exist.

The first seeks to bypass the need for photoreceptors. This can be achieved by either making the surviving retinal bipolar or ganglion cells photosensitive using optogenetics (Busskamp et al., 2010Yue et al., 2016) or synthetic photo-switchable compounds (Polosukhina et al., 2012Tochitsky et al., 2017). Additionally, retinal prostheses capable of stimulating RGCs directly have been developed, attempting to encode visual information directly into these output neurons (da Cruz et al., 2016Lewis et al., 2016).

Of note, the use of retinal prostheses for blindness was approved by the European Union in 2011, and by the FDA in 2013. Use of these prostheses has led to some successful reacquisition of very basic visual functions but only for limited periods of time (Mills et al., 2017).

The second and more promising approaches aims to replace the lost photoreceptors by transplantation or by stimulating regeneration. These later approaches have received special attention because they have the capability of renewing the native function of the retina, and could provide a real cure for RD. Due to this potential for complete functional recovery, and because of recent and important developments in the field, transplantation and regeneration will be a focus of this review.

Barriers in Photoreceptor Transplantation as a Therapy for RD

Just a decade ago, the prospect of producing photoreceptors from stem cells seemed like an overly daunting task (Adler, 2008).

Nevertheless, in the last few years, several laboratories have successfully developed protocols to produce eyecup-like structures from induced-pluripotent stem cells (iPSCs) in the span of weeks. Some of these eyecups are able to acquire a layered structure reminiscent of the retina, with photoreceptor-like cells that contain outer segments, express phototransduction proteins (Wahlin et al., 2017), and have some capacity for light responsiveness (Zhong et al., 2014), and vesicular release (Wahlin et al., 2017).

The successful development of these eyecups opens the possibility of harvesting cells from an individual to generate iPSCs, and re-differentiate them into photoreceptors that could be then transplanted back into patients with RD. Based on these prospects, recent work in the retinal field has focused on using mice to explore photoreceptor transplantation as a therapy for RD.

Initial transplantation studies in mice attempted to introduce rod photoreceptors, with the best rates of integration (never surpassing a few percent) achieved by transplanting immature rod precursors (MacLaren et al., 2016).

Follow-up studies presented equally promising examples of integration, and in some cases demonstrated functional recovery of vision (Santos-Ferreira et al., 2015Smiley et al., 2016).

However, it has recently been discovered that many of these results are due to the exchange of cytoplasmic material (including RNA and/or proteins) between donor cells and the host retina, and not integration of transplanted photoreceptors (Pearson et al., 2016Santos-Ferreira et al., 2016Singh et al., 2016Ortin-Martinez et al., 2017). In light of this recent discovery, it will be important to carefully interpret how functional recovery of vision occurred after transplantation/cytoplasmic exchange in degenerated or degenerating retinas (Homma et al., 2013Singh et al., 2013Santos-Ferreira et al., 2015).

Even if cytoplasmic exchange results in recovery photoreceptor function, it occurs at very low rates (a few percent of host positive cells, for transplantations of tens to hundreds of thousands of donor cells). Such low rates casts doubt on the prospect of leveraging this process as a viable therapeutic strategy, especially in advanced cases of degeneration.

Even with viable evidence for successful photoreceptor transplantation (Waldron et al., 2018), there are additional concerns for this type of therapy. For example, subretinal injection of a mass of cells, the most common transplantation method, causes inflammation and scarring, and inhibits the migration of transplanted cells (Barber et al., 2013).

Additionally, it is also unclear if transplanted photoreceptors are capable of rewiring properly into the host retina.

This problem is further compounded by our incomplete understanding of how photoreceptors normally wire during development. To date, only a handful of genes are known to be involved in synapse formation between photoreceptors and their postsynaptic targets (Simmons et al., 2017Zhang et al., 2017Miller et al., 2018Sarria et al., 2018Ueno et al., 2018).

Despite this work, we still have little insight on the molecules that drive the initial recognition between these cells, or on the processes that promote, inhibit or refine synapse formation. Unveiling genes involved in photoreceptor synapse formation during normal development in zebrafish could provide direct therapeutic targets to promote rewiring of transplanted photoreceptors.

Using Zebrafish to Explore Retinal Regeneration as a Therapy for RD

Cumulatively, work on transplantation therapies has highlighted that alternative therapies, such as photoreceptor regeneration, could be a promising alternative. Unfortunately, in mammals there is no regeneration in the retina after damage or RD. In contrast, the zebrafish retina has the innate capacity for regeneration. This capacity may be due to the continued growth of the zebrafish retina into adulthood, as well as the ability of the zebrafish to maintain a population of multipotent stem cells within the retina.

Larval zebrafish form a functional visual system by 4 days post fertilization (4 dpf), and are able to perform complex visual guided behaviors (like prey capture, see below) by 5 dpf (Patterson et al., 2013). This rapid onset of sensory function is critical to survival of the animal. As larvae progress into adulthood, zebrafish continue to grow in size. This growth requires organs like the eye and retina to grow as well. In the retina this growth occurs in the ciliary marginal zone (CMZ).

The CMZ maintains a niche of pluripotent cells at the edge of the retina that continually adds neurons in peripheral concentric rings (Centanin et al., 2011). In addition to this continual growth, zebrafish can also regenerate their retinas after injury. In fact, robust retinal regeneration and rewiring have been demonstrated in genetic models of RD and in other models that incur retinal injury.

Overall, given that zebrafish is a genetically tractable model with active retinal regeneration, it is poised to uncover the molecular processes that control retinal regeneration and rewiring.

In teleost fish, retinal regeneration after injury has a rich history. It was first reported in goldfish (Lombardo, 1968) and later in cichlids (Johns and Fernald, 1981) and trout (Julian et al., 1998). In zebrafish retinal regeneration is robust after resection (Cameron, 2000), mechanical damage (Fausett and Goldman, 2006), light damage (Bernardos et al., 2007Thomas et al., 2012), thermal damage (Raymond et al., 2006), pharmacological damage (Fimbel et al., 2007Sherpa et al., 2008Nagashima et al., 2013Tappeiner et al., 2013Sherpa et al., 2014) and selective ablation of particular cell-types (Montgomery et al., 2010D’Orazi et al., 2016Hagerman et al., 2016Yoshimatsu et al., 2016White et al., 2017). In teleosts, regeneration can occur from cells generated in the CMZ (Raymond et al., 2006), a dedicated population of progenitors that are committed to a rod fate (Bernardos et al., 2007Morris et al., 2008), and the Müller glia (Fausett and Goldman, 2006Bernardos et al., 2007Fimbel et al., 2007).

Due to its location at the edge of the retina, the CMZ is only involved in regeneration if the injury involves the peripheral retina. During regeneration, the CMZ is capable of giving rise to all retinal neurons except rod photoreceptors (Stenkamp et al., 2001Raymond et al., 2006). Instead, rods originate from rod-specific progenitors.

These progenitors were first identified in goldfish and cichlids (Johns and Fernald, 1981) and were later found in other teleost fish including trout and zebrafish (Julian et al., 1998). Rod-specific progenitors are thought to be important for maintaining the density of rods as the eye grows, and lineage tracing revealed that these rod progenitors derive from Müller cells that slowly and continuously divide in the normal retina (Otteson et al., 2001Raymond et al., 2006Bernardos et al., 2007Nelson et al., 2008).

During regeneration, there is an expansion in the number of photoreceptor progenitors, but these mainly derive from actively dividing Müller glia. In fact, in zebrafish the Müller glia are the primary source of regenerated neurons after injury. During regeneration they can act as multipotent stem cells, dividing and differentiating into any retinal cell type (Ramachandran et al., 2010b).

In contrast to zebrafish, in humans and other mammals Müller glia do not remain multipotent and therefore cannot readily replace lost neurons in the retina. Because Müller glia are the primary source of regenerated retinal neurons and can regenerate all retinal neurons, considerable work has been dedicated to understanding the differences between the Müller glia of zebrafish and mammals.

A series of studies that investigated the response of zebrafish Müller glia to retinal injury, have unveiled the key transcription factors in a gene regulatory network that controls retinal repair. Shortly after injury, cytokines and growth factors activate the beta-catenin and stat3 pathways (Kassen et al., 2007Wan et al., 2014).

These pathways upregulate the expression of ascl1 (Fausett et al., 2008), a key transcription factor that (through lin-28) leads to the suppression of let7 microRNA (Ramachandran et al., 2010a).

In the uninjured retina, let7 normally represses the expression of many regeneration-induced genes (including ascl1 and lin-28), closing the loop of a system poised to control Müller glia response to injury (Wan and Goldman, 2016) (Figure ​Figure2A2A).

In contrast, mammalian Müller glia does not readily divide (Wan et al., 2008) and responds to retinal injury with an inflammatory response known as reactive gliosis, characterized by an increase in size and overproduction of intermediate filaments, and leading to distortion of the architecture of the retina without repair (Dyer and Cepko, 2000Bringmann et al., 2009Thomas et al., 2016) (Figure ​Figure2C2C). Significant efforts have been devoted to understanding the differences between these species, in the hope of stimulating regeneration in mammals.

This work has shown that ascl1 is not upregulated in mice after retinal injury (Karl et al., 2008), but ascl1 overexpression in mammalian Müller cells in vitro is sufficient to induce production of neurons (Pollak et al., 2013). Moreover, induction of expression of ascl1 in Müller cells in vivo, followed by retinal injury, induces division and production of all classes of retinal neurons, but only in young mice (Ueki et al., 2015).

Recently, a successful report of regeneration in adult mice has shown that new bipolar- and amacrine-like cells derived from Müller glia can rewire into the retina. For this work, in addition to overexpression of ascl1, inhibition of histone deacetylation was also required (Jorstad et al., 2017). Unfortunately, no other retinal cell types are produced with this protocol (Figure ​Figure2D2D).

Further insight into retinal regeneration has been gained from studying a related telost, medaka, which shows a restricted capacity for regeneration. In medaka fish, after injury, Müller glia do not readily proliferate, and new retinal progenitors commit almost exclusively to a photoreceptor fate.

Comparisons in the Müller glia response to retinal injury in both medaka fish and zebrafish concluded that sustained expression of the transcription factor sox2 in adult Müller cells is key for maintaining multipotency (Lust and Wittbrodt, 2018) (Figure ​Figure2B2B). While it is clear that important strides have been taken to attempt retinal regeneration in mammals, before regeneration can be used as a viable therapy, we need a deeper understanding on the mechanisms that maintain cells with a regenerative potential in zebrafish throughout adulthood.

Insights from Zebrafish on Rewiring After Regeneration

Even after successful transplantation or regeneration of photoreceptors, the biggest hurdle in these RD therapies is ensuring that the new photoreceptors rewire into the appropriate retinal circuits so that they are able to restore normal visual function.

Once again, zebrafish has offered a unique opportunity to study rewiring after injury and regeneration, in both larvae and adults. Cumulatively, this work has demonstrated that the extent and time course of regeneration and rewiring is determined by lesion-specific differences, in particular the extent of injury and the number of cells that need to be replaced.

Adult teleosts are able to regenerate their retinas even after extensive retinal damage. Early seminal studies in adult goldfish, and later studies in adult zebrafish, showed robust retinal regeneration and rewiring after surgical retinal extirpation (Hitchcock et al., 1992Cameron, 2000) or pharmacologically induced death of all retinal neurons (Raymond et al., 1988Sherpa et al., 2008).

Under these lesion paradigms, all retinal cell types were regenerated. Importantly, with regards to rewiring, the retinal lamination was reestablished (Raymond et al., 1988Sherpa et al., 2008), and synaptic connections were reformed between retinal neurons (Hitchcock and Cirenza, 1994) and between RGCs and the brain (Stuermer et al., 1985).

Parallel work demonstrated that after extensive retinal damage in adult goldfish and zebrafish, visual function is also recovered (Mensinger and Powers, 19992007Lindsey and Powers, 2007Sherpa et al., 2008). Nevertheless, because the Müller glia are the main source of regenerated retinal neurons, complete ocular excision prevents regeneration (Mensinger and Powers, 2007).

In adult teleosts, although there is robust regeneration after extensive retinal injury, the time required for regeneration and functional recovery depends on the extent of injury. For example, the differences in regeneration and rewiring were examined after surgical extirpation of ∼25, 50, and 75% of the adult retina.

This work found that after surgical extirpation of 25% of the retina, 17 weeks (120 days) are required for regeneration and reestablishment of lamination and 25 weeks (180 days) for functional recovery. Additional extirpation lengthened the time and extent of recovery for both lamination and functional recovery (Mensinger and Powers, 2007).

Similar results were observed in adults after pharmacological damage and death of all retinal neurons (induced by high doses of intraocular ouabain). In this study functional recovery started 5 weeks after the injury with further improvement by weeks 7 – 10, albeit with decreased sensitivity (Mensinger and Powers, 1999Lindsey and Powers, 2007).

In adult zebrafish, the extent and time course of retinal regeneration and rewiring is also dependent on the injury.

Extensive pharmacological damage of all retinal layers with ouabain leads to regeneration, and the newly formed cells are capable to reorganize into the three distinct nuclear layers by week 3 after injury.

After 14 weeks, the regenerated retinas are well laminated (clear nuclear and plexiform layers), the optic nerve has regrown, and there is functional recovery (Sherpa et al., 2008).

Interestingly, with pharmacological damage (using lower doses of ouabain) that spares photoreceptors and Müller glia but still induces a loss of cells in the INL and GCL, regeneration is faster, with significant recovery of function after only 8 weeks (Sherpa et al., 2014McGinn et al., 2018). In this lesion paradigm, rewiring of a specific (but heterogeneous) subset of regenerated BCs was closely examined.

The regenerated BCs had largely normal morphology, and, as a population, were able to reproduce the diversity of connectivity patterns observed in the surviving photoreceptors, with only a few errors in lamination or abnormal dendritic or axonal arborization, again emphasizing the robust regeneration of zebrafish (McGinn et al., 2018).

Nevertheless, in the context of extensive damage, regeneration in adult teleosts is far from perfect.

Several structural defects are common including areas with defects in the formation or absence of plexiform layers, disorganization of nuclear layers, presence of cells in the incorrect layer (e.g., RGCs in INL), failure to reestablish the photoreceptor mosaic, formation of photoreceptor rosettes, overproduction of neurons, and the generation of cell types that were not initially damaged (Raymond et al., 1988Hitchcock et al., 1992Cameron, 2000Vihtelic Thomas and Hyde David, 2000Stenkamp et al., 2001Stenkamp and Cameron, 2002Sherpa et al., 2008Powell et al., 2016).

More recently, the genetic tractability of zebrafish has enabled researchers to damage specific retinal cell types and study their rewiring after regeneration.

This work was accomplished by using the recently developed nitroreductase–metronidazole (NTR–MTZ) system. For this method, transgenic zebrafish are created expressing the NTR gene (nfsb) under the control of a cell-specific promoter.

When these transgenic zebrafish are treated with the compound MTZ, the NTR converts MTZ into a cytotoxic compound. Because this compound does not diffuse to neighboring cells, the resulting ablation is restricted to the NTR-expressing cells.

Importantly, this process is reversible, and removal of MTZ solution makes it possible to examine regeneration and rewiring. To date, the NTR-MTZ system has been used to selectively ablate rods, and specific subtypes of cones, bipolar cells and glial cells (Zhao et al., 2009Ariga et al., 2010Montgomery et al., 2010Fraser et al., 2013D’Orazi et al., 2016Hagerman et al., 2016White et al., 2017). Importantly, several of these studies demonstrated that other cells in the retina that did not express NTR were not ablated after MTZ application. In addition, these treatments did not affect the surrounding retinal architecture (Zhao et al., 2009). This highlights the specificity and power of the NTR-MTZ system.

After genetic ablation and removal of MTZ, in each instance the targeted cells regenerated after several days, although the exact time-course varied depending on the ablated cell type and the age of zebrafish treated.

For example, after using the NTR–MTZ to completely ablate rods in adult zebrafish, newly generated rods were identified within a week after removal of MTZ, and repopulation of rods attained pre-injury levels within 4 weeks (Montgomery et al., 2010) – a very similar time course required for the regeneration of cones in adults (Raymond et al., 2006Bernardos et al., 2007).

In larvae, the regeneration of cells occurs at a faster timescale. For example, after ablation of rods using the NTR-MTZ system in 5 dpf larvae, newly formed rods attained control levels in just 6 days (White et al., 2017). Similarly, cones ablated between 4 and 6 dpf regenerate in 7 – 10 days (Fraser et al., 2013Yoshimatsu et al., 2016) and BCs ablated at 7 dpf regenerate in 13 days (D’Orazi et al., 2016).

In the majority of these studies, regeneration was confirmed morphologically. In a subset of studies, after regeneration, the analysis was extended to include behavior or rewiring. For example, in one study, either the UV- or S-cones were ablated (in 7 dpf larvae) and the optomotor response (OMR) (see below) was assayed after ablation and following UV- or S-cone regeneration respectively (Hagerman et al., 2016).

The OMR was reduced immediately after ablation of either UV- or S-cones. Surprisingly, while the OMR recovery took 4 days for the UV-cone ablation, the OMR recovered in just 1 day following S-cone ablation, before new S-cones were produced.

These differences in behavioral recovery suggest that there may be a capacity for plasticity amongst the remaining cells, used to compensate for the ablated cells during the recovery phase. It is possible that this short-term plasticity relies on activity from other cone subtypes and/or on synaptic remodeling.

Evidence for such synaptic remodeling has been reported in a parallel study (Yoshimatsu et al., 2016). In this study, a subtype of horizontal cell (H3) that normally connects preferentially to UV- and S-cones, was able to reconnect to UV-cones after UV-cone specific ablation and regeneration.

Nevertheless, if UV-cone regeneration was delayed, the H3 made additional contacts with S-cones and even M- and L-cones, suggesting functional compensation at the level of rewiring. Another recent study examined rewiring after selective loss of a subpopulation of BC using the NTR-MTZ system in 5 dpf larvae (D’Orazi et al., 2016).

Thirteen days after the ablation of these BC, the majority of regenerated BC were morphologically normal but the rewiring did not fully recapitulate development, with a relative loss of selectivity for specific cone subtypes. Additionally, BC axons contained significantly more synapses.

As a whole, work in this field proves that retinal regeneration in zebrafish is a robust process, but also suggests that some of the developmental cues required to refine synapse number or proportion of photoreceptor subtypes innervated may not be present during regeneration.

In the future, it will be important to further understand what cues are present during development that enable photoreceptors to wire into different retinal circuits. It will be especially important to understand how specific photoreceptor subtypes recognize the different bipolar- and horizontal-cell subtypes, and the factors required for the formation of these synapses.

It will also be important to recognize and examine the developmental and environmental differences between larval and adult zebrafish retinas. This knowledge will provide a comprehensive understanding of the differences between development and regeneration, between wiring, rewiring and remodeling, and likely uncover manipulations that could be used to modify or refine rewiring in the context of treatments for RD.


More information: Nergis Kara et al. The miR-216a-Dot1l Regulatory Axis Is Necessary and Sufficient for Müller Glia Reprogramming during Retina Regeneration, Cell Reports (2019). DOI: 10.1016/j.celrep.2019.07.061

Journal information: Cell Reports
Provided by Vanderbilt University

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