Optic Nerve Regeneration: How Will We Get There?

Background: Restoration of vision in patients blinded by advanced optic neuropathies requires technologies that can either 1) salvage damaged and prevent further degeneration of retinal ganglion cells (RGCs), or 2) replace lost RGCs. Evidence acquisition: Review of scientific literature. Results: In this article, we discuss the different barriers to cell-replacement based strategies for optic nerve regeneration and provide an update regarding what progress that has been made to overcome them. We also provide an update on current stem cell-based therapies for optic nerve regeneration. Conclusions: As neuro-regenerative and cell-transplantation based strategies for optic nerve regeneration continue to be refined, researchers and clinicians will need to work together to determine who will be a good candidate for such therapies.

State-of-the-Art Review Section Editors: Fiona Costello, MD, FRCP(C) Sashank Prasad, MD Optic Nerve Regeneration: How Will We Get There? Kimberly K. Gokoffski, MD, PhD, Phillip Lam, BS, Basheer F. Alas, BS, Micalla G. Peng, BS, Heidi R. R. Ansorge Background: Restoration of vision in patients blinded by advanced optic neuropathies requires technologies that can either 1) salvage damaged and prevent further degeneration of retinal ganglion cells (RGCs), or 2) replace lost RGCs. Evidence Acquisition: Review of scientific literature. Results: In this article, we discuss the different barriers to cellreplacement based strategies for optic nerve regeneration and provide an update regarding what progress that has been made to overcome them. We also provide an update on current stem cell-based therapies for optic nerve regeneration. Conclusions: As neuro-regenerative and cell-transplantation based strategies for optic nerve regeneration continue to be refined, researchers and clinicians will need to work together to determine who will be a good candidate for such therapies. Journal of Neuro-Ophthalmology 2020;40:234-242 doi: 10.1097/WNO.0000000000000953 © 2020 by North American Neuro-Ophthalmology Society R estoration of vision in patients blinded by advanced optic neuropathies requires technologies that are neuroprotective approaches (prevent further degeneration of retinal ganglion cells [RGCs]), neuroregenerative approaches (salvage damaged RGCs), or cell replacement approaches (replace lost RGCs). Neuroprotective and neuroregenerative approaches are more naturally suited for chronic, progressive optic neuropathies such as glaucoma, hereditary, toxic, and metabolic optic neuropathies than for acute optic neuropathies such as traumatic, ischemic, demyelinating, and inflammatory conditions. Although there may be a critical window in which patients with acute Department of Ophthalmology, Roski Eye Institute, University of Southern California, Los Angeles, California. K. K. Gokoffski is supported by a KL2 career development award from the SC-CTSI (NCATS UL1TR001855) and an unrestricted grant to USC Roski Eye Institute from Research to Prevent Blindness). M. G. Peng and B. F. Alas were supported by the Keck Summer Research Fellowship. The authors report no conflicts of interest. Address correspondence to Kimberly K. Gokoffski, Department of Ophthalmology, Roski Eye Institute, University of Southern California, 1450 San Pablo Street, 4th Floor, Los Angeles, CA 91208; E-mail: [e-mail] 234 optic neuropathies could benefit from neuroprotective therapies, they are more likely to require cell replacement therapies. There are a number of barriers that must be overcome before cell replacement strategies can be translated to the clinical setting (Fig. 1). First and foremost, transplantation approaches require a source of healthy RGCs to transplant. Second, these cells must be integrated into the native retina, so that they may interface with amacrine and bipolar cells. Integrated cells must then sprout an axon that will grow into the nerve fiber layer of the retina, toward the optic nerve head, and then through the lamina cribrosa into the optic nerve toward various synaptic partners in the diencephalon, without taking detours. Finally, once reaching the optic tectum, the axons of these RGCs must generate new, retinotopic appropriate synaptic connections. In this article, we will discuss these different challenges and provide an update regarding the progress that has been made to overcome them. We will finish with a discussion of current stem cell-based optic nerve regeneration methods that are being evaluated in clinical trials. LIMITED ENDOGENOUS REGENERATION IN THE ADULT MAMMALIAN CENTRAL NERVOUS SYSTEM In mammals, axon regeneration after injury is the rule for the peripheral nervous system (PNS) but the exception when it comes to the central nervous system (CNS). The limited regenerative capacity of the CNS was believed to be mostly cell intrinsic until experiments demonstrated that CNS neurites readily grow into peripheral nerve grafts (1,2). The notion that environmental cues in the CNS limit regeneration, rather that factors solely intrinsic to the neurons, was further supported by experiments demonstrating blockage of sciatic nerve regeneration with optic nerve grafts (3). What these landmark studies also suggested is that extrinsic environmental factors can override intrinsic limitations to regeneration in the CNS. These studies fueled investigations to identify growth permissive molecules within the PNS and growth inhibitory molecules within the CNS, with the hope that manipulating their relative levels Gokoffski et al: J Neuro-Ophthalmol 2020; 40: 234-242 Copyright © North American Neuro-Ophthalmology Society. Unauthorized reproduction of this article is prohibited. State-of-the-Art Review FIG. 1. Schematic of proposed steps and obstacles for developing stem cell-based therapies for optic nerve regeneration. Cell replacement-based strategies for optic nerve regeneration need a source of healthy retinal ganglion cells (RGCs) for transplantation. 1) fibroblasts isolated from patients can be used to develop inducible pluripotent stem cells (iPSCs), which could then be 2) transdifferentiated into RGCs through either 3D retinal organoids or 2D planar cultures. 3) RGCs would then be purified and injected into patient eyes. 4) Currently, transplanted RGCs have poor rates of integration into the retina. 5) Of the RGCs that successfully integrated, limited axonogenesis out of the eye into the optic nerve or past glial scar has been observed. 6) Regenerating axons have also been observed to overshoot their synaptic target. Upon reaching the lateral geniculate nucleus (77), new synaptic connections would need to be made. would unlock the regenerative capacity of the CNS. Decades of work have culminated in the identification of intrinsic regulators of neuronal growth (e.g., cAMP, mechanistic target of rapamycin [mTOR]/PTEN, APC-Cdh1, and KLF4) (4,5) and extrinsic factors such as myelin byproducts (e.g., myelinassociated glycoprotein, Nogo, and oligodendrocyte myelin glycoprotein) and glial scar as playing important roles in limiting regeneration in the CNS. For example, myelin byproducts are shed from injured axons and are cleared by Schwann cells and blood-borne macrophages in the PNS (6-10). The absence of Schwann cells in the CNS is believed to account, in part, for the relatively poor axon regeneration seen in the CNS. CRITICAL BARRIERS TO REGENERATION AND CURRENT CELL TRANSPLANTATION- BASED APPROACHES Source of Retinal Ganglion Cells The first step in developing cell replacement-based strategies for optic nerve regeneration requires a reliable, highvolume source of healthy RGCs. Given the scarcity and ethical dilemmas associated with using cadaveric retinas and human embryonic stem cells (hESCs), the scientific community was challenged with developing an alternative source. In 2006, Yamanaka followed by Thompson reported a remarkable breakthrough technology whereby adult fibroblasts could be reprogrammed into pluripotent stem cells using 4 transcription factors (Oct3/4, Sox2, Gokoffski et al: J Neuro-Ophthalmol 2020; 40: 234-242 c-Myc, and Klf4) (11,12). This landmark development, termed inducible pluripotent stem cells (iPSCs), has turned optic nerve regeneration from science fiction to possibility. These transdifferentiated cells exhibit similar characteristics to hESCs including the ability to propagate indefinitely and the ability to differentiate into many different cell types including RGCs. Although the eye is an immuneprivileged organ, the fact that these cells would have the same HLA make-up as their hosts confer another advantage over hESC-derived RGCs; iPSC-derived cells are less likely to be destroyed by the immune system. Currently, there are 2 different methods for generating RGCs from stem cells (hESCs and iPSCs): 1) organoid differentiation and 2) planar differentiation. Significant work is being performed to understand which of these methods is the most robust for producing RGCs that resemble the native RGC. To date, most of the published protocols to generate RGCs from stem cells have been developed with hESCs, although significant effort is being made to translate these technologies to iPSCs as well. Organoid-Derived Retinal Ganglion Cells Organoids are self-organizing, 3-dimensional miniature organs developed in vitro from pluripotent stem cells. These miniorgans can be used to study disease pathology and developmental biology, and for screening drugs. In 2015, Kuwahara et al (13) devised a protocol to develop 3D retinal organoids from hESCs. Using a cocktail of growth factors including bone morphogenic protein 4 (BMP4) in 235 Copyright © North American Neuro-Ophthalmology Society. Unauthorized reproduction of this article is prohibited. State-of-the-Art Review combination with a nonadherent tissue culture surface, hESCs can be coaxed into differentiating into self-organizing retinal cups. Remarkably, these organoids have been shown to recapitulate many aspects of retinal development in vitro, expressing the same sequence of transcription factors that characterize endogenous retinal development (14,15) and self-organizing into layers similar to the native retina (Fig. 2) (16,17). Mature RGCs are readily seen in culture starting day 20 of differentiation and continue to develop through day 34. Healthy organoids can be kept alive in culture for many months, although RGCs start to degenerate after 3 months in culture (16). RGCs can be readily purified from organoids. Photoreceptor-driven action potentials have been recorded from iPSC-derived RGCs, albeit sluggish compared with native RGCs (18). This suggests that iPSC-derived retinal organoids develop functional, albeit rudimentary, retinal circuitry. As these technologies continue to be refined, iPSC organoids may serve as a reliable source from which large volumes of immune compatible, healthy RGCs can be purified and transplanted into patients. Planar-Derived Retinal Ganglion Cells An alternative to organoids, RGCs, can be transdifferentiated from stem cells using traditional two-dimensional cell-culture techniques. Reh devised a protocol to induce hESCs into a retinal lineage, giving rise to the major retinal cell types in vitro (19). Mature RGCs can be identified as soon as 15 days in culture and can be reliably purified after 30 days in culture (20). hESC-derived RGCs were shown to have electrophysiological properties similar to mature, native RGCs (20). In addition, whole-cell voltage-clamp recordings demonstrate that hESC-derived RGCs are glutamate responsive. Although the hESC-derived RGCs display morphologic characteristics of native RGCs, these 2D cultures do not produce other retinal cell types and do not develop the retinal circuitry seen in organoid-based cultures. A major limitation to using autologous iPSCs as a source of RGCs is that any underlying genetic predispositions to neurodegeneration would still be present in the cells used to generate new RGCs. For example, Ohlemacher extracted skin fibroblasts from a patient containing a mutation in the optineuron (OPTN) gene, which has been implicated in normal tension glaucoma (21). He reprogrammed these cells into iPSCs and then differentiated them into RGCs. These iPSC-derived RGCs exhibited increased apoptosis relative to control RGCs but could be rescued by the addition of neuroprotective factors. Although this established an in vitro model of glaucoma, these patients are unlikely to be able to serve as autologous donors. Instead, iPSC donor banks would need to be established. Although production of RGCs from hESCs is discussed here in general terms, there are over 30 different subtypes of RGCs (22). These RGC subtypes include ON-OFF directionally selective ganglion cells (ONOFF DSGCs), ON directionally sensitive ganglion cells (ON DSGCs), intrinsically photosensitive retinal ganglion cells, and alpha-RGCs (aRGCs) among others. Studies have shown that not all types of RGCs are uniformly lost in different forms of optic neuropathies. For instance, 14 days after optic nerve crush, Duan demonstrated 80% of aRGCs survived (23). Daniel et al (24) found a subtype of aRGCs (transient OFF aRGCs) to be the most vulnerable to optic nerve crush injury compared with 3 other RGC subtypes. Similar results were demonstrated in a mouse model of glaucoma where intraocular pressure was transiently elevated (25,26). Interestingly, different subtypes of RGCs are not produced at equal rates in hESC cultures. Among hESC-derived RGCs, ON-OFF and ON DSGCs consist of about 17% and 30% of the total RGC population, respectively, whereas aRGCs were found in about 21% of the population (27). A continued understanding of which subtypes of RGCs are damaged in different types of optic neuropathies may allow for the development of more targeted therapies. Integration of Transplanted Retinal Ganglion Cells Great strides have been made to develop technologies that allow for large volume production of iPSC-derived RGCs FIG. 2. Retinal organoids derived from human embryonic stem cell line. A, C. Self-organizing retinal organoids derived from the H9 human embryonic stem cell line, day 25 of differentiation. Cell-line courtesy of Dr. Donald Zack (Johns Hopkins University), pictures courtesy of Dr. Narine Harutyunyan (Children's Hospital Los Angeles Stem Cell Core). B. Red fluorescent cells represent mature retinal ganglion cells (RGCs) expressing tdTomato under the control of the Brn3B promoter (a class-4 POU domain transcription factor expressed in mature RGCs). Scale bar = 100 mm. 236 Gokoffski et al: J Neuro-Ophthalmol 2020; 40: 234-242 Copyright © North American Neuro-Ophthalmology Society. Unauthorized reproduction of this article is prohibited. State-of-the-Art Review for cell transplantation. Unfortunately, simply injecting RGCs into the vitreous does not lead to meaningful levels of RGC integration into the ganglion cell layer (GCL) of the retina. Goldberg reported between 1% and 7% of RGCs integrated into the GCL after intravitreal injection with 40- 60,000 RGCs (28). Interestingly, improved integration rates were noted when smaller amounts of RGCs were injected into the eye. When Goldberg cotransplanted RGCs with iPSCs into the vitreous, they saw retinal engraftment rates on the order of 20% (29). RGCs derived from healthier cultures or younger retinas were also associated with higher integration rates (28,30). There are a number of factors that may account for the poor rates of RGC integration: 1) structural obstacles to the GCL, such as the retinal inner limiting membrane (ILM), 2) postinjection inflammatory response, 3) extracellular matrix (ECM), and 4) immune-mediated clearance of the transplanted cells. The ILM, which serves as a vitreoretinal barrier, is formed by astrocytes and end feet of Müller glial cells that are covered in a thick basement membrane comprising ECM proteins. The finding that greater than 99% of RGCs injected into the eye lay at the vitreous ILM interface implicated the ILM as a possible barrier to RGC integration (31). This notion is supported by ex vivo cultures, which showed that the amount of mesenchymal stem cell (MSC) retinal integration correlated with the amount of ILM peeled. Interestingly, ILM peeling was also correlated with reduction in glial reactivity, which represents another barrier to cell integration, as discussed below (32). By contrast, digestion of the ILM with collagenase and plasmin did not demonstrate a significant effect on MSC proliferation, survival, or migration into the retina (32). In addition to the ILM, intravitreal injections have been found to elicit a gliotic reaction in the retina that may inhibit RGC integration. Astrocytes and Müller glia have been shown to upregulate glial fibrillary acidic protein and vimentin after intravitreal injections, creating an inhospitable environment for RGC integration (33). When the glial cell reaction was suppressed by aminoadipic acid treatment, there was a 4-fold increase in MSC migration in vivo (34). This treatment, however, may not be feasible for translation to human application because aminoadipic acid treatment impairs normal physiological function of Müller cells. On a similar vein, Goldberg showed that RGC integration rates in ex vivo retinal cultures whose optic nerves were previously axotomized were superior in the 28-day group than earlier groups (30). They proposed that more effective RGC integration can be achieved in an RGC-depleted retina after the inflammatory response has cleared. This is supported by experiments by Singhal who successfully used antiinflammatory and ECM-degrading matrix agents to increase integration (35). A less understood obstacle to RGC integration is the inhospitable ECM environment that develops in the retina as Gokoffski et al: J Neuro-Ophthalmol 2020; 40: 234-242 it matures. ECM molecules including CD44 and neurocan, a chondroitin sulfate proteoglycan, have been shown to increase in concentration as the retina matures (36,37). Reducing levels of retinal CD44 and neurocan in MRL/ MpJ mice was associated with enhanced integration of photoreceptors into the mature retina and increased regeneration (38). Singhal also saw significantly improved integration of Müller stem cells into all retinal layers when chondroitinase ABC (and concomitant immunosuppression) was used to degrade chondroitin sulfate proteoglycans (39). Thus, it may be worthwhile to consider ECM degradation treatment as an adjuvant to future RGC transplantation approaches. Although the eye is an immune-privileged organ, it is possible that immune-mediated clearance of injected RGCs contributes to poor integration rates. Bull found immunosuppression to improve rates of transplanted cell survival (40). Upon successful migration into the GCL, another challenge to RGC integration involves the formation of appropriate synaptic connections with retinal amacrine and bipolar interneurons. Remarkably, Goldberg noted that RGCs that migrated into the GCL displayed normal RGClike arboritec structures (28). Furthermore, using synaptic marker staining and light stimulation, they demonstrated the formation of novel and functional synapses between donor RGCs and host retina. These connections, however, were found to be weaker than native RGCs because increased stimulation was needed to elicit an action potential in transplanted RGCs. Given experiences with photoreceptor transplantation, one concern regarding experiments reporting RGC integration is whether true RGC integration even occurs. Transplantation studies with GFP-labeled photoreceptors have found that rather than integrating, transplanted photoreceptors were transferring cytoplasmic material (i.e., green fluorescent tracing material) to endogenous photoreceptors, giving the false appearance of cellular integration. Although the mechanism is unknown, it is believed that photoreceptors transfer cytoplasmic material through exosome vesicular transport (41,42). This phenomenon of cytoplasmic transfer, however, is believed to be unique to photoreceptors who routinely shed their outer segments and does not to apply to RGCs. This argument is bolstered by reports from Goldberg who observed 1) morphological differences between the axon terminals of GFP-labeled donor cells and mature host RGC termini and 2) only noted single nuclei in transplanted RGCs (28). Future studies involving interspecies transplantation (i.e., human iPSC-derived RGCs into rat retina) will be able to use markers of donor genetic material to demonstrate true integration. Axon Growth As detailed above, when RGCs were injected into the vitreous, few integrated into the retina. Of the RGCs that successfully integrated into the GCL, many sprouted axons 237 Copyright © North American Neuro-Ophthalmology Society. Unauthorized reproduction of this article is prohibited. State-of-the-Art Review that grew toward the optic nerve head but few grew past the lamina cribrosa (w10%) (28). Whether poor axonogenesis is a result of absence of external guidance cues or presence of inhibitory molecules is under active investigation. During development, surface proteins including ephrin-A and their respective ligands, EphA, are expressed in gradients within the superior colliculus and RGC population, respectively (43). These molecules have been implicated in directing axonogenesis during optic nerve development, helping establish the topographic map between the retina and superior colliculus. Experiments aimed at exploiting these pathways to recapitulate development and thereby direct optic nerve regeneration, however, have been met with limited success; for example, ephrin-A2 expression in the superior colliculus was found to have an inhibitory effect on RGC axon regeneration (44). A similar effect was reported with semaphorin-3A (45). Failure to direct optic nerve regeneration may stem from the fact that these pathways work in not only spatial but also temporal gradients, which are difficult to recapitulate in vivo. Other approaches to promote axonogenesis include providing RGCs with neurotrophic support such as with nerve growth factor and brain-derived neurotrophic factor (46). The rationale for these approaches stem from the finding that neurons depend on retrograde transport of endosomes containing neurotrophic factors for survival (47). By providing neurotrophic support, RGCs can be kept alive longer, increasing the opportunity for axon regeneration. The problem, however, is that increased neuronal survival did not equate to new axon formation (48,49). More targeted approaches aimed at resetting cell intrinsic programs to proaxonogenic states have been met with greater success. Activation of growth-promoting molecules such as mTOR and Janus kinase/signal transducers and activators of transcription (JAK/STAT) has been associated with long-distance RGC axon regeneration after optic nerve crush injury (50-52). There are a number of caveats, however, associated with these approaches. First, these therapies seem to benefit only a subset of RGCs: activation of mTOR or overexpression of oncomodulin and insulin-like growth factor 1 selectively promoted regeneration of alpha-RGCs, which comprise only 6% of the total RGC population (23). Second, the efficacy of mTOR and JAK/STAT in directing optic nerve regeneration have only been demonstrated when their levels were upregulated before or concurrently with axon injury (50,53), which may not be clinically practical. Moreover, given that RGC axon degeneration after crush injury occurs over the course of weeks (54), it is difficult to determine whether these experiments promoted de novo axon growth or instead prevented axon degeneration. Third, axons have been observed to grow toward aberrant targets (e.g., into the contralateral optic nerve or back on itself) (50,55,56), implying a need for not just cues that promote but also ones that direct growth. 238 The approach that our group is taking to overcome the problem of limited axonogenesis is to use electric fields (EFs) to direct RGC axon regeneration. Often overlooked, electrical gradients coexist with chemoattractants and repellants and have been found to play an important role in directing tissue patterning during development (57) and wound healing (58). Recently, we demonstrated that RGC axons exhibit cathode-directed growth in vitro (59). In vivo, Borgens demonstrated that transected spinal cords of sea lampreys and guinea pigs who underwent continuous EF stimulation had more action potentials and more axons traversing across the transection site, respectively, than controls (60,61). These approaches lost momentum, in large part, from an inability to translate well into large animal studies. Borgens applied a direct current, which is well known to be toxic above even minute levels, and others tried alternating current, which was found to be ineffectual (62). The latter finding was not surprising, given our work showed RGC axons making U-turns in response to 180° changes in electric field polarity, a condition that mimics alternating current (59). Since these studies, advances in electrical engineering have fueled the development of hybrid currents such as asymmetric charge-balanced waveforms that allow for both safe and effective activation neurons in the CNS (63). The efficacy of such currents on directing RGC axon regeneration in vivo is currently under active investigation in our laboratory, and preliminary experiments show promising results. A major advantage that may be gained from taking an electrical over molecular approach is that EFs can override endogenous molecular directional cues: in vivo EF application redirected neural stem cells moving within the rostral migratory stream to novel targets (64). This suggests that in vivo application of EFs may be able to override endogenous inhibitors of axon regeneration (e.g., myelin byproducts and glial scar) (46). Alternative strategies for directing optic nerve regeneration include fabricated scaffolds and autologous grafts. Recently, 3D printers have been used to print microgrooved and nanogrooved scaffolds that can be used to direct the orientation of axonal growth (20,65,66). Other groups hope to exploit the regenerative capacity of the PNS by using sciatic nerve grafts to facilitate optic nerve regeneration (67). Ultimately, optic nerve regeneration is going to require a combinatorial approach with signals that provide not only the "drive" to grow but also directional cues that can "steer" axons toward proper targets. New Synapse Formation The question of whether regenerated axons will establish new, functional synapses with the diencephalon remains a significant question in the field of optic nerve regeneration. This task is further complicated by the necessity of maintaining the retinotopic map. Progress in this field has been hampered by limited long-distance axonogenesis Gokoffski et al: J Neuro-Ophthalmol 2020; 40: 234-242 Copyright © North American Neuro-Ophthalmology Society. Unauthorized reproduction of this article is prohibited. State-of-the-Art Review (see previous section). To circumvent this, Bei lesioned the optic tract just proximal to the superior colliculus (51). They found that upregulation of mTOR and JAK/STAT or co-overexpression of osteopontin (OPN)/insulin-like growth factor (IGF1)/ciliary neurotrophic factor (CNTF) was associated with increased formation of functional synapses with the superior colliculus compared with controls. These mice, however, did not show improvement on visual aptitude testing. They found newly formed axons to be unmyelinated, but that partial restoration of visual function could be achieved with administration of 4-aminopyridine (4-AP), a voltage-gated potassium channel blocker. Other approaches to promote synaptogenesis exploit processes that directed synapse formation during development. In utero, before direct stimulation by light, spontaneous neural activity termed "retinal waves" prime the developing visual system for eventual light stimulation (68). This organizational process is believed to be driven by voltage-dependent calcium influx that mediates transcriptional changes that then drive new synapse formation. After birth, visual stimulation refines established visual neural networks, which dictate retinotopy and ocular dominance. Retinal wave disruption, either pharmacologically or genetically, interferes with retinotopic mapping and eye-specific segregation in the lateral geniculate nucleus and superior colliculus in mice (69). Building on this concept, recent data show that high-contrast visual stimulation in conjunction with mTOR activation enhanced RGC regeneration after crush injury and lead to partial recovery of visual function including the optokinetic reflex, the visual cliff test, and the looming avoidance response (53). In these experiments, Lim crushed one optic nerve and sutured the nonlesioned eye shut, forcing all light input to the visual cortex to come from the lesioned optic nerve. These mice demonstrated increased RGC regeneration, electrical activ- ity, and synaptogenesis compared with lesioned mice in which both eyelids were left open. CURRENT CLINICAL TRIALS To date, few active stem cell-based clinical trials for optic nerve regeneration are listed on clinicaltrials.gov (Table 1). As emphasized above, this is because the field is still in early stages of development. This is well exemplified by the recent disastrous outcomes of patients with age-related macular degeneration who lost significant vision after intravitreal injection with adipose-derived stem cells (70). These trials were able to circumvent FDA regulation on the technicality that they were transplanting autologous cells. Rather than cell replacement, current stem cell-based therapies likely work by providing neurotrophic support to host tissue. Although researchers and clinicians alike want the best for their patients who are debilitated by their vision loss, the community must proceed with caution when endorsing these trials to prevent public loss of faith in the potential of stem cell-based therapies. CONCLUSIONS In parallel to the progress being made to develop neuroregenerative and cell transplantation-based strategies for optic nerve regeneration, researchers and clinicians together will need to determine who will be a good candidate for such therapies. Trans-synaptic retrograde and anterograde degeneration have been well described in the visual pathway. For example, patients with advanced glaucoma have documented degeneration in the lateral geniculate nucleus (71). Patients with posterior visual pathway lesions have been shown to have correlated degeneration of their retinal nerve fiber layer, suggesting anterograde degeneration (72,73). Given this, the TABLE 1. Stem cell-based clinical trials for optic nerve regeneration Clinicaltrials.gov ID Number, Location NCT01920867, USA NCT03011541, USA NCT02638714, Jordan Treatment Delivery Method Condition Autologous Bone Marrow-derived Stem cells Combination of retrobulbar, subtenon, intravitreal, intraoptic nerve, subretinal, and intravenous injection Retinal disease Macular degeneration Hereditary retinal dystrophy Optic nerve disease Glaucoma Status Results/Findings Enrolling by invitation NAION: 73.6% of treated eyes gained vision with an average of 3.53 Snellen lines of vision improvement (72). LHON: Up to 35 letters of improvement by ETDRS (73) ETDRS, early treatment diabetic retinopathy study; LHON, Leber's Hereditary Optic Neuropathy; NAION, nonarteritic ischemic optic neuropathy. Gokoffski et al: J Neuro-Ophthalmol 2020; 40: 234-242 239 Copyright © North American Neuro-Ophthalmology Society. Unauthorized reproduction of this article is prohibited. State-of-the-Art Review success of cell replacement-based therapies will depend on the health of the residual supporting environment. In other words, if we manage to regenerate the optic nerve in patients with advanced glaucoma, will there be neurons in the lateral geniculate nucleus for them to synapse with? In considering the many endogenous barriers to axon regeneration in the CNS, it was interesting to learn that myelin, a lipoprotein well known for its insulating function, also robustly inhibits axon growth after injury. This finding raises the question of whether myelin serves to prevent axon growth in the nontraumatic state, assuring that neural networks, once established would not change? Prevention of axonogenesis in fully developed, healthy tissue may be important for preventing inappropriate rewiring and the formation of misguided, aberrant connections. Numerous examples of aberrant regeneration can be readily found in the PNS: reparative responses to severe damage to the oculomotor nerve such as from crush injury can result in miswiring between different muscle groups such as the levator palpebrae and inferior rectus, causing inappropriate co-contraction. Conversely, few examples of acquired CNS aberrancy have been reported in the clinical literature (74). Most examples of CNS synkinesis are found in congenital conditions (e.g., Joubert Disease) in which aberrant connections were likely made before myelination set in (75). Although myelination is associated with decreased CNS plasticity, does it in fact play a direct role in regulating the stability of neuronal networks? (76) Is poor clearance of myelin from the CNS after injury a protective response to guard against aberrant regeneration? Given the strong antiregenerative effect of myelin, successful CNS regeneration will likely require targeted blockade of these inhibitory signals. Although cell replacement strategies are becoming a viable option for restoration of vision in patients blinded by advanced optic neuropathies, there are still a number of significant challenges that must be overcome before they can be safely implemented in the clinical setting. Technical advances that merge basic science with biomedical engineering to control intrinsic and extrinsic regulators of neuroregeneration are essential to maintain the current momentum of translating RGC transplantation to the clinical arena. STATEMENT OF AUTHORSHIP Category 1: a. Conception and design: K. K. Gokoffski; b. Acquisition of data: K. K. Gokoffski, P. Lam, B. F. Alas, M. G. Peng, and H. R. R. Ansorge; c. Analysis and interpretation of data: K. K. Gokoffski, P. Lam, B. F. Alas, M. G. Peng, and H. R. R. Ansorge. Category 2: a. Drafting the manuscript: K. K. Gokoffski, P. Lam, B. F. Alas, M. G. Peng, and H. R. R. Ansorge; b. Revising it for intellectual content: K. K. Gokoffski. Category 3: a. Final approval of the completed manuscript: K. K. Gokoffski. REFERENCES 1. Richardson PM, McGuinness UM, Aguayo A. J. Axons from CNS neurons regenerate into PNS grafts. Nature. 1980;284:264-265. 240 2. Ramón y Cajal S, DeFelipe J, Jones EG. Cajal's Degeneration and Regeneration of the Nervous System. Oxford, UK: Oxford University Press, 1991. 3. Aguayo AJ, Dickson R, Trecarten J, Attiwell M, Bray GM, Richardson P. Ensheathment and myelination of regenerating PNS fibres by transplanted optic nerve glia. Neurosci Lett. 1978;9:97-104. 4. Yang P, Yang Z. Enhancing intrinsic growth capacity promotes adult CNS regeneration. J Neurol Sci. 2012;312:1-6. 5. Sun F, He Z. Neuronal intrinsic barriers for axon regeneration in the adult CNS. Curr Opin Neurobiol. 2010;20:510-518. 6. Benowitz LI, He Z, Goldberg JL. Reaching the brain: advances in optic nerve regeneration. Exp Neurol. 2017;287:365-373. 7. Yiu G, He Z. Glial inhibition of CNS axon regeneration. Nat Rev Neurosci. 2006;7:617-627. 8. Schwab ME. Repairing the injured spinal cord. Science. 2002;295:1029-1031. 9. Goldberg JL, Vargas ME, Wang JT, Mandemakers W, Oster SF, Sretavan DW, Barres BA. An oligodendrocyte lineage-specific semaphorin, Sema5A, inhibits axon growth by retinal ganglion cells. J Neurosci. 2004;24:4989-4999. 10. Fabes J, Anderson P, Yáñez-Muñoz RJ, Thrasher A, Brennan C, Bolsover S. Accumulation of the inhibitory receptor EphA4 may prevent regeneration of corticospinal tract axons following lesion. Eur J Neurosci. 2006;23:1721-1730. 11. Takahashi K, Yamanaka S. Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell. 2006;126:663-676. 12. Yu J, Vodyanik MA, Smuga-Otto K, Antosiewicz-Bourget J, Frane JL, Tian S, Nie J, Jonsdottir GA, Ruotti V, Stewart R, Slukvin II, Thomson JA. Induced pluripotent stem cell lines derived from human somatic cells. Science. 2007;318:1917-1920. 13. Kuwahara A, Ozone C, Nakano T, Saito K, Eiraku M, Sasai Y. Generation of a ciliary margin-like stem cell niche from selforganizing human retinal tissue. Nat Commun. 2015;6:6286. 14. Völkner M, Zschätzsch M, Rostovskaya M, Overall RW, Busskamp V, Anastassiadis K, Karl MO. Retinal organoids from pluripotent stem cells efficiently recapitulate retinogenesis. Stem Cell Rep. 2016;6:525-538. 15. Fligor CM, Langer KB, Sridhar A, Ren Y, Shields PK, Edler MC, Ohlemacher SK, Sluch VM, Zack DJ, Zhang C, Suter DM, Meyer JS. Three-dimensional retinal organoids facilitate the investigation of retinal ganglion cell development, organization and neurite outgrowth from human pluripotent stem cells. Sci Rep. 2018;8:14520. 16. Aparicio JG, Hopp H, Choi A, Mandayam Comar J, Liao VC, Harutyunyan N, Lee TC. Temporal expression of CD184(CXCR4) and CD171(L1CAM) identifies distinct early developmental stages of human retinal ganglion cells in embryonic stem cell derived retina. Exp Eye Res. 2017;154:177-189. 17. Meyer JS, Howden SE, Wallace KA, Verhoeven AD, Wright LS, Capowski EE, Pinilla I, Martin JM, Tian S, Stewart R, Pattnaik B, Thomson JA, Gamm DM. Optic vesicle-like structures derived from human pluripotent stem cells facilitate a customized approach to retinal disease treatment. Stem Cells. 2011;29:1206-1218. 18. Hallam D, Hilgen G, Dorgau B, Zhu L, Yu M, Bojic S, Hewitt P, Schmitt M, Uteng M, Kustermann S, Steel D, Nicholds M, Thomas R, Treumann A, Porter A, Sernagor E, Armstrong L, Lako M. Human-induced pluripotent stem cells generate light responsive retinal organoids with variable and nutrientdependent efficiency. Stem Cells. 2018;36:1535-1551. 19. Lamba DA, Karl MO, Ware CB, Reh TA. Efficient generation of retinal progenitor cells from human embryonic stem cells. Proc Natl Acad Sci U S A. 2006;103:12769-12774. 20. Sluch VM, Davis CH, Ranganathan V, Kerr JM, Krick K, Martin R, Berlinicke CA, Marsh-Armstrong N, Diamond JS, Mao HQ, Zack DJ. Differentiation of human ESCs to retinal ganglion cells using a CRISPR engineered reporter cell line. Sci Rep. 2015;5:16595. 21. Ohlemacher SK, Sridhar A, Xiao Y, Hochstetler AE, Sarfarazi M, Cummins TR, Meyer JS. Stepwise differentiation of retinal Gokoffski et al: J Neuro-Ophthalmol 2020; 40: 234-242 Copyright © North American Neuro-Ophthalmology Society. Unauthorized reproduction of this article is prohibited. State-of-the-Art Review 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 40. ganglion cells from human pluripotent stem cells enables analysis of glaucomatous neurodegeneration. Stem Cells. 2016;34:1553-1562. Sanes JR, Masland RH. The types of retinal ganglion cells: current status and implications for neuronal classification. Annu Rev Neurosci. 2015;38:221-246. Duan X, Qiao M, Bei F, Kim IJ, He Z, Sanes JR. Subtype-specific regeneration of retinal ganglion cells following axotomy: effects of osteopontin and mTOR signaling. Neuron. 2015;85:1244- 1256. Daniel S, Clark AF, McDowell CM. Subtype-specific response of retinal ganglion cells to optic nerve crush. Cell Death Discov. 2018;4:7. Ou Y, Jo RE, Ullian EM, Wong RO, Della Santina L. Selective vulnerability of specific retinal ganglion cell types and synapses after transient ocular hypertension. J Neurosci. 2016;36:9240-9252. Feng L, Zhao Y, Yoshida M, Chen H, Yang JF, Kim TS, Cang J, Troy JB, Liu X. Sustained ocular hypertension induces dendritic degeneration of mouse retinal ganglion cells that depends on cell type and location. Invest Ophthalmol Vis Sci. 2013;54:1106-1117. Langer KB, Ohlemacher SK, Phillips MJ, Fligor CM, Jiang P, Gamm DM, Meyer JS. Retinal ganglion cell diversity and subtype specification from human pluripotent stem cells. Stem Cell Rep. 2018;10:1282-1293. Venugopalan P, Wang Y, Nguyen T, Huang A, Muller KJ, Goldberg JL. Transplanted neurons integrate into adult retinas and respond to light. Nat Commun. 2016;7:10472. Wu S, Chang KC, Nahmou M, Goldberg JL. Induced pluripotent stem cells promote retinal ganglion cell survival after transplant. Invest Ophthalmol Vis Sci. 2018;59:1571-1576. Hertz J, Qu B, Hu Y, Patel RD, Valenzuela DA, Goldberg JL. Survival and integration of developing and progenitor-derived retinal ganglion cells following transplantation. Cell Transpl. 2014;23:855-872. Johnson LN, Cashman SM, Kumar-Singh R. Cell-penetrating peptide for enhanced delivery of nucleic acids and drugs to ocular tissues including retina and cornea. Mol Ther. 2008;16:107-114. Johnson TV, Bull ND, Martin KR. Transplantation prospects for the inner retina. Eye. 2009;23:1980-1984. Kinouchi R, Takeda M, Yang L, Wilhelmsson U, Lundkvist A, Pekny M, Chen DF. Robust neural integration from retinal transplants in mice deficient in GFAP and vimentin. Nat Neurosci. 2003;6:863-868. Johnson TV, Bull ND, Martin KR Identification of barriers to retinal engraftment of transplanted stem cells. Invest Ophthalmol Vis Sci. 2010;51:960-970. Singhal S, Bhatia B, Jayaram H, Becker S, Jones MF, Cottrill PB, Khaw PT, Salt TE, Limb GA. Human Müller glia with stem cell characteristics differentiate into retinal ganglion cell (RGC) precursors in vitro and partially restore RGC function in vivo following transplantation. Stem Cells Transl Med. 2012;1:188-199. Nishina S, Hirakata A, Hida T, Sawa H, Azuma N. CD44 expression in the developing human retina. Graefes Arch Clin Exp Ophthalmol. 1997;235:92-96. Inatani M, Honjo M, Otori Y, Oohira A, Kido N, Tano Y, Honda Y, Tanihara H. Inhibitory effects of neurocan and phosphacan on neurite outgrowth from retinal ganglion cells in culture. Invest Ophthalmol Vis Sci. 2001;42:1930-1938. Tucker B, Klassen H, Yang L, Chen DF, Young MJ. Elevated MMP expression in the MRL mouse retina creates a permissive environment for retinal regeneration. Invest Ophthalmol Vis Sci. 2008;49:1686-1695. Singhal S, Lawrence JM, Bhatia B, Ellis JS, Kwan AS, Macneil A, Luthert PJ, Fawcett JW, Perez MT, Khaw PT, Limb GA. Chondroitin sulfate proteoglycans and microglia prevent migration and integration of grafted Müller stem cells into degenerating retina. Stem Cells. 2008;26:1074-1082. Bull ND, Limb GA, Martin KR. Human Muller stem cell (MIO-M1) transplantation in a rat model of glaucoma: survival, Gokoffski et al: J Neuro-Ophthalmol 2020; 40: 234-242 41. 42. 43. 44. 45. 46. 47. 48. 49. 50. 51. 52. 53. 54. 55. 56. 57. 58. 59. differentiation, and integration. Invest Ophthalmol Vis Sci. 2008;49:3449-3456. Santos-Ferreira T, Llonch S, Borsch O, Postel K, Haas J, Ader M. Retinal transplantation of photoreceptors results in donor-host cytoplasmic exchange. Nat Commun. 2016;7:13028. Ortin-Martinez A, Tsai EL, Nickerson PE, Bergeret M, Lu Y, Smiley S, Comanita L, Wallace VA. A reinterpretation of cell transplantation: GFP transfer from donor to host photoreceptors. Stem Cells. 2017;35:932-939. McLaughlin T, O'Leary DD. Molecular gradients and development of retinotopic maps. Annu Rev Neurosci. 2005;28:327-355. Symonds AC, King CE, Bartlett CA, Sauvé Y, Lund RD, Beazley LD, Dunlop SA, Rodger J. EphA5 and ephrin-A2 expression during optic nerve regeneration: a "two-edged sword". Eur J Neurosci. 2007;25:744-752. Giger RJ, Pasterkamp RJ, Holtmaat AJ, VerhaagenSemaphorin J III. Role in neuronal development and structural plasticity. Prog Brain Res. 1998;117:133-149. Gokoffski KK PM, Alas B, Lam P. Neuro-protection and neuroregeneration of the optic nerve: recent advances and future directions. Curr Opin Neurol. 2020;33:93-105. Harrington AW, Ginty DD. Long-distance retrograde neurotrophic factor signalling in neurons. Nat Rev Neurosci. 2013;14:177-187. Goldberg JL, Espinosa JS, Xu Y, Davidson N, Kovacs GT, Barres BA. Retinal ganglion cells do not extend axons by default: promotion by neurotrophic signaling and electrical activity. Neuron. 2002;33:689-702. Mesentier-Louro LA, Rosso P, Carito V, Mendez-Otero R, Santiago MF, Rama P, Lambiase A, Tirassa P. Nerve growth factor role on retinal ganglion cell survival and axon regrowth: effects of ocular administration in experimental model of optic nerve injury. Mol Neurobiol. 2019;56:1056-1069. Sun F, Park KK, Belin S, Wang D, Lu T, Chen G, Zhang K, Yeung C, Feng G, Yankner BA, He Z. Sustained axon regeneration induced by co-deletion of PTEN and SOCS3. Nature. 2011;480:372-375. Bei F, Lee HHC, Liu X, Gunner G, Jin H, Ma L, Wang C, Hou L, Hensch TK, Frank E, Sanes JR, Chen C, Fagiolini M, He Z. Restoration of visual function by enhancing conduction in regenerated axons. Cell. 2016;164:219-232. Park KK, Liu K, Hu Y, Smith PD, Wang C, Cai B, Xu B, Connolly L, Kramvis I, Sahin M, He Z. Promoting axon regeneration in the adult CNS by modulation of the PTEN/mTOR pathway. Science. 2008;322:963-966. Lim JH, Stafford BK, Nguyen PL, Lien BV, Wang C, Zukor K, He Z, Huberman AD. Neural activity promotes long-distance, target-specific regeneration of adult retinal axons. Nat Neurosci. 2016;19:1073-1084. Li Y, Schlamp CL, Nickells RW. Experimental induction of retinal ganglion cell death in adult mice. Invest Ophthalmol Vis Sci. 1999;40:1004-1008. Luo X, Salgueiro Y, Beckerman SR, Lemmon VP, Tsoulfas P, Park KK. Three-dimensional evaluation of retinal ganglion cell axon regeneration and pathfinding in whole mouse tissue after injury. Exp Neurol. 2013;247:653-662. Pernet V, Joly S, Dalkara D, Jordi N, Schwarz O, Christ F, Schaffer DV, Flannery JG, Schwab ME. Long-distance axonal regeneration induced by CNTF gene transfer is impaired by axonal misguidance in the injured adult optic nerve. Neurobiol Dis. 2013;51:202-213. McCaig CD, Rajnicek AM, Song B, Zhao M. Controlling cell behavior electrically: current views and future potential. Physiol Rev. 2005;85:943-978. Song B, Gu Y, Pu J, Reid B, Zhao Z, Zhao M. Application of direct current electric fields to cells and tissues in vitro and modulation of wound electric field in vivo. Nat Protoc. 2007;2:1479-1489. Gokoffski KK, Jia X, Shvarts D, Xia G, Zhao M. Physiologic electrical fields direct retinal ganglion cell axon growth in vitro. Invest Ophthalmol Vis Sci. 2019;60:3659-3668. 241 Copyright © North American Neuro-Ophthalmology Society. Unauthorized reproduction of this article is prohibited. State-of-the-Art Review 60. Borgens RB, Roederer E, Cohen MJ. Enhanced spinal cord regeneration in lamprey by applied electric fields. Science. 1981;213:611-617. 61. Borgens RB, Bohnert DM. The responses of mammalian spinal axons to an applied DC voltage gradient. Exp Neurol. 1997;145:376-389. 62. Graves MS, Hassell T, Beier BL, Albors GO, Irazoqui PP. Electrically mediated neuronal guidance with applied alternating current electric fields. Ann Biomed Eng. 2011;39:1759-1767. 63. Merrill DR, Bikson M, Jefferys JG. Electrical stimulation of excitable tissue: design of efficacious and safe protocols. J Neurosci Methods. 2005;141:171-198. 64. Feng JF, Liu J, Zhang L, Jiang JY, Russell M, Lyeth BG, Nolta JA, Zhao M. Electrical guidance of human stem cells in the rat brain. Stem Cell Rep. 2017;9:177-189. 65. Johansson F, Carlberg P, Danielsen N, Montelius L, Kanje M. Axonal outgrowth on nano-imprinted patterns. Biomaterials. 2006;27:1251-1258. 66. Yang TC, Chuang JH, Buddhakosai W, Wu WJ, Lee CJ, Chen WS, Yang YP, Li MC, Peng CH, Chen SJ. Elongation of axon extension for human iPSC-derived retinal ganglion cells by a nano-imprinted scaffold. Int J Mol Sci. 2017;18:E2013. 67. Cen LP, Luo JM, Geng Y, Zhang M, Pang CP, Cui Q. Long-term survival and axonal regeneration of retinal ganglion cells after optic nerve transection and a peripheral nerve graft. Neuroreport. 2012;23:692-697. 68. Ackman JB, Crair MC. Role of emergent neural activity in visual map development. Curr Opin Neurobiol. 2014;24:166-175. 69. Hooks BM, Chen C. Distinct roles for spontaneous and visual activity in remodeling of the retinogeniculate synapse. Neuron. 2006;52:281-291. 242 70. Kuriyan AE, Albini TA, Townsend JH, Rodriguez M, Pandya HK, Leonard RE II, Parrott MB, Rosenfeld PJ, Flynn HW Jr, Goldberg JL. Vision loss after intravitreal injection of autologous "stem cells" for AMD. New Engl J Med. 2017;376:1047-1053. 71. Gupta N, Greenberg G, de Tilly LN, Gray B, Polemidiotis M, Yücel YH. Atrophy of the lateral geniculate nucleus in human glaucoma detected by magnetic resonance imaging. Br J Ophthalmol. 2009;93:56-60. 72. Dinkin M. Trans-synaptic retrograde degeneration in the human visual system: slow, silent, and real. Curr Neurol Neurosci Rep. 2017;17:16. 73. Mitchell JR, Oliveira C, Tsiouris AJ, Dinkin MJ. Corresponding ganglion cell atrophy in patients with postgeniculate homonymous visual field loss. J Neuroophthalmol. 2015;35:353-359. 74. Messe SR, Shin RK, Liu GT, Galetta SL, Volpe NJ. Oculomotor synkinesis following a midbrain stroke. Neurology. 2001;57:1106-1107. 75. Lee SK, Kim DI, Kim J, Kim DJ, Kim HD, Kim DS, Mori S. Diffusion-tensor MR imaging and fiber tractography: a new method of describing aberrant fiber connections in developmental CNS anomalies. Radiographics. 2005;25:53- 65; discussion 66-58. 76. Kapfhammer JP, Schwab ME. Inverse patterns of myelination and GAP-43 expression in the adult CNS: neurite growth inhibitors as regulators of neuronal plasticity? J Comp Neurol. 1994;340:194-206. 77. Li M, Yu JSL, Tilgner K, Ong SH, Koike-Yusa H, Yusa K. Genome-wide CRISPR-KO screen uncovers mTORC1-mediated Gsk3 regulation in naive pluripotency maintenance and dissolution. Cell Rep. 2018;24:489-502. Gokoffski et al: J Neuro-Ophthalmol 2020; 40: 234-242 Copyright © North American Neuro-Ophthalmology Society. Unauthorized reproduction of this article is prohibited.