Article

Strategies for Optic Nerve Regeneration

Where are we now?

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No treatment exists that can reverse the permanent vision loss that results from retinal ganglion cell (RGC) degeneration in primary open-angle glaucoma (POAG). Vision loss is permanent because RGCs have poor capacity to self-repair and no capacity for self-regeneration. Strategies to restore vision in patients with optic neuropathies like POAG will need to (1) stop RGC degeneration, (2) promote RGC axon regeneration, or (3) replace lost RGCs. This review aims to provide a succinct update on the current questions surrounding and the barriers to optic nerve regeneration.

Why Does Damage to the Axon Lead to RGC Degeneration?

Although the exact site of injury in POAG is unknown, much work has pointed to the biomechanics at the level of the optic nerve head as an inciting source.1 Why would damage to the axon lead to degeneration of the RGC soma itself? Part of the answer to this question lies in neuronal survival depending on growth factors secreted in the synaptic cleft. In the synaptic terminal, neurotrophic factors, such as nerve growth factor (NGF), are produced and secreted by downstream neurons. These neurotrophic factors bind receptors (eg, TrkB) on RGC axon terminals, and this complex is then internalized by the RGC and the vesicle and is transported retrograde within the axon back to the soma, where it activates transcriptional programs (eg, PI3K) that are necessary for neuron survival and growth.2 This dependence on downstream growth signals may serve as a mechanism through which the RGC soma in the retina “checks in” with its axon terminal, ensuring that the axon is still functional. This theory is supported by RGC degeneration occurring more rapidly when axotomy is performed closer to the soma than when it is performed more distally.

Why Can’t the Optic Nerve Regenerate?

Relative to the peripheral nervous system (PNS), the central nervous system (CNS) has poor capacity to self-repair. Neither system has the capacity to self-renew. Barriers to regeneration in the CNS are categorized as cell-intrinsic or cell-extrinsic barriers.

Cell-Intrinsic Barriers

Cell-intrinsic barriers to regeneration are established during development, when organs transition from a state of proliferation to differentiation to maturation. These steps are generally irreversible, with mature cells focusing on carrying out their intended function. While some organs (eg, the liver) maintain a reserve of stem cells to regenerate damaged cells, the CNS does not. An increased understanding of the signaling pathways responsible for directing brain and retinal development has led researchers to hypothesize that reactivation of these pathways may be able to revert RGCs to a proregenerative state. Indeed, different genetic tools used to activate mammalian target of rapamycin (mTOR) and JAK/STAT have led to increased RGC axon regeneration after optic nerve crush injury in rodent experiments.3-6 Researchers have also shown that controlling zinc levels7,8 and activation of intracellular inflammation (via puncturing the lens capsule)9 are associated with increased RGC axon regeneration after crush injury.

There are limitations associated with these genetic approaches. First, they have only been shown to be effective when activated before or at the same time as crush injury, limiting clinical application. Second, activation of these pathways carries the risk of neoplastic conversion. Misregulation of mTOR underlies the phacomatosis syndrome called tuberous sclerosis.10 Finally, regenerated axons have been shown to grow toward aberrant targets (eg, back on itself or across the chiasm to the other optic nerve, as opposed to the superior colliculus).3,11 This knowledge indicates that regenerative approaches require signals that not only promote axon growth, but also signals that will direct growth.

Cell-Extrinsic Barriers

Studies from the 1980s showing that crushed RGC axons grow into peripheral nerve grafts12,13 were pivotal in demonstrating that (1) the extracellular environment in the CNS plays a role in limiting axon regeneration, and (2) these cell-extrinsic signals can override cell-intrinsic inhibition. Since then, researchers have identified myelin basic protein and glial scarring as major cell-extrinsic inhibitors of regeneration in the CNS.

In the case of myelin, the composition of CNS myelin differs from that of the PNS. This difference is thought to account for some of the difference in regenerative capacity between the 2 systems. Specifically, myelin-associated glycoprotein (MAG), myelin oligodendrocyte glycoprotein (MOG), and neurite outgrowth inhibitor (Nogo) are unique to the CNS and have been found to inhibit axon outgrowth.14-17 When oligodendrocytes and myelin-associated proteins were inhibited by monoclonal antibody, there was enhanced regeneration after optic nerve injury.18 Another molecule, secretory leukocyte protease inhibitor (SLPI), has also been shown to overcome the inhibitory effect of myelin-associated proteins and to improve RGC regeneration in rats.19

Glial scar formation, mediated by reactive astrocytes, has also been shown to impede axonal regrowth,20 leading many to hypothesize that modulation of the astrocytic response may create a more conducive extracellular environment for neuroregenerative therapies.21 One molecule that has been implicated in the regulation of astrocyte activation is microRNA-21 (miR-21).22 Rats treated with an miR-21 inhibitor demonstrated a reduction in astrocyte activation and glial scar elaboration, as well as increased optic nerve regeneration after crush.23

Our group is investigating the potential for exogenous electric field (EF) application as an approach to directing RGC axon regeneration. Electric fields naturally occur in the body and have been shown to direct long-distance axon growth.24 Electric fields have been recorded in the developing retina and have been proposed to direct formation of the optic nerve head.25 Recently, we demonstrated that EFs direct RGC axon growth in tissue culture experiments,26 and there are promising preliminary in vivo data that suggest that EFs robustly direct RGC axon regeneration after optic nerve crush injury. Importantly, EF application appears to be effective even when applied weeks after axon injury. The mechanism through which EFs are thought to direct axon regeneration is by selectively activating voltage-sensitive calcium channels that subsequently initiate axon building pathways.27

Is Promoting RGC Survival Enough to Regenerate the Optic Nerve?

Given that RGC degeneration occurs as a result of growth factor deprivation (described above), would replacing these factors or preventing apoptosis be sufficient to promote axon regeneration? In other words, is the failure to regenerate an axon a result of RGCs not surviving long enough?

To test this theory, researchers have investigated the effect of neurotrophic factor supplementation on promoting axon regeneration. Both eyedrops and intravitreal applications of nerve growth factor (NGF) have been investigated in rodent models after optic nerve crush injury and have been shown to have a dose-dependent effect on counteracting RGC death. Unfortunately, NGF had minimal effects on promoting axon regrowth.28 Monotherapy with other neurotrophic factors, such as brain-derived neurotrophic factor and ciliary neurotrophic factor, demonstrated increased rates of neuronal survival after optic nerve crush injury; however, they failed to produce significant axon regeneration, even with the use of peripheral nerve grafts.11,18,29

Although failure of these approaches to demonstrate avid axon regeneration could be attributed, in part, to failure to sustain sufficient levels of neurotrophic factor signaling,30,31 other experiments have suggested that the problem stems from RGC survival and axon growth being regulated by independent signaling pathways. For example, nearly 100% of RGCs survived optic nerve crush injury in Bcl-2 transgenic mice, but virtually none of these cells regenerated their axons.32 This outcome suggests that, in addition to providing neurotrophic support, optic nerve regeneration will require approaches that also promote axon growth. The results of recent clinical trials testing the role of intravitreal delivery of neurotrophic factors in POAG and an antiapoptosis molecule for nonarteritic ischemic optic neuropathy are pending.

Can Stem Cells Be Used to Restore Vision in Patients With End-Stage POAG?

The advent of human embryonic stem cells (hESCs) and inducible pluripotent stem cells (iPSCs) has turned cell replacement-based strategies from science fiction to plausible. The overarching goal of these strategies is to transform stem cells into healthy RGCs, integrate them into the retina, direct them to sprout new axons that grow out of the eye and into the optic nerve and, upon reaching their synaptic target in the diencephalon, form new and appropriate synapses. Each of these steps is no small feat to overcome.

Development of Stem Cells and Their Transformation Into RGCs

In 2006, Yamanaka applied transcription factors to adult fibroblasts to reprogram them into iPSCs.33 This momentous breakthrough overcame obstacles of scarcity and ethical concerns with using hESCs or cadaveric retinas. Using a cocktail of growth factors, hESCs and iPSCs can be transformed into retinal organoids or self-organizing, 3D miniature retinas.34

Retinal organoids recapitulate many aspects of retinal development in vitro, including basic retinal organization and photoreceptor-driven action potentials. Although RGCs can be readily purified from these organoids, these procedures are still costly and not ready for high-volume production. A major advantage gained from transplanting iPSC-derived RGCs over hESC-derived RGCs or cadaveric donations is that they are less likely to be destroyed by the immune system because they are derived from the host and thus are immune compatible. A disadvantage, however, is that these RGCs would have the same genetic makeup and thus be disposed to the same degeneration that occurred in the host.

Retinal Integration

There are a number of physical and environmental obstacles that limit RGC integration into the retina, such as the inner limiting membrane, postinjection inflammatory response, and extracellular matrix. In rodents, RGC integration rates into the retina ranged from 1% to 7%, with 3% being the most common.35 Integration rates were increased when the inner limiting membrane was peeled.36 Fortunately, of the RGCs that managed to integrate, many appeared to form connections and functional synapses with native retinal cells, as well as to elicit action potentials, albeit weaker than native RGCs.

Directing Axon Growth

Technologies being developed to direct RGC axon regeneration after axon injury may be applicable for cell-transplantation strategies for optic nerve regeneration. In addition to the approaches discussed above, other groups are investigating the use of printable nanoscaffolds,37 as well as peripheral nerve graft transplantation,38,39 as alternative strategies for directing axon growth.

Synapse Formation

Successful optic nerve regeneration is contingent on long-distance axon regeneration and new synapse formation, which recapitulate the intrinsic retinotopic map. Researchers have demonstrated that neuroactivation is necessary for new synapse formation. During development, the retina is primed by “retinal waves” to establish functional connections with higher brain structures and prepare the eye for light stimulation.40 Postnatal light exposure is critical for establishing and strengthening these visual pathways. Leveraging this knowledge, researchers have demonstrated that high-contrast light stimulation in conjunction with mTOR activation led to partial recovery of visual function after optic nerve injury in rodents.

Discussion

Ultimately, the development of effective neuroregenerative therapies for POAG will require a combinatorial approach, incorporating strategies that modulate both extrinsic and intrinsic barriers to regeneration. As the community continues to develop regenerative and cell-transplantation approaches for the optic nerve, it is important to consider which patients will constitute good candidates for treatment. With the slow, progressive degeneration that characterizes glaucoma, early diagnosis and treatment may yield improved responses to eyedrops or intravitreal administration of neurotrophic factors. Conversely, patients presenting at a more advanced stage of disease may require more significant intervention in the form of cell-replacement therapies. Regardless of approach, the success of any therapy will be limited by the health of the surrounding structures. In other words, successful regeneration of the optic nerve will depend on there being a healthy lateral geniculate nucleus to receive and connect with regenerating RGC axons.

One advantage gained from not having the ability to regenerate is that the CNS mitigates the risk of having newly regenerated axons disrupt established circuits if they regrow and reconnect incorrectly. This advantage contrasts with the PNS, which has modest regenerative capacity and numerous examples of aberrant regeneration. In light of this finding, regenerative approaches will need not only to promote axon growth but also to direct their growth to maintain the retinotopic map of the visual pathway. GP

References

  1. Downs JC, Girkin CA. Lamina cribrosa in glaucoma. Curr Opin Ophthalmol. 2017;28(2):113-119.
  2. Harrington AW, Ginty DD. Long-distance retrograde neurotrophic factor signalling in neurons. Nat Rev Neurosci. 2013;14(3):177-187.
  3. Benowitz LI, He Z, Goldberg JL. Reaching the brain: Advances in optic nerve regeneration. Exp Neurol. 2017;287(Pt 3):365-373.
  4. Park KK, Liu K, Hu Y, et al. Promoting axon regeneration in the adult CNS by modulation of the PTEN/mTOR pathway. Science. 2008;322(5903):963-966.
  5. Bei F, Lee HHC, Liu X, et al. Restoration of visual function by enhancing conduction in regenerated axons. Cell. 2016;164(1-2):219-232.
  6. de Lima S, Koriyama Y, Kurimoto T, et al. Full-length axon regeneration in the adult mouse optic nerve and partial recovery of simple visual behaviors. Proc Natl Acad Sci U S A. 2012;109(23):9149-9154.
  7. Trakhtenberg EF, Li Y, Feng Q, et al. Zinc chelation and Klf9 knockdown cooperatively promote axon regeneration after optic nerve injury. Exp Neurol. 2018;300:22-29.
  8. Li Y, Andereggen L, Yuki K, et al. Mobile zinc increases rapidly in the retina after optic nerve injury and regulates ganglion cell survival and optic nerve regeneration. Proc Natl Acad Sci U S A. 2017;114(2):E209-E218.
  9. Kurimoto T, Yin Y, Omura K, et al. Long-distance axon regeneration in the mature optic nerve: contributions of oncomodulin, cAMP, and pten gene deletion. J Neurosci. 2010;30(46):15654-15663.
  10. Curatolo P, Moavero R. mTOR inhibitors in tuberous sclerosis complex. Curr Neuropharmacol. 2012;10(4):404-415.
  11. Pernet V, Joly S, Dalkara D, et al. 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.
  12. Richardson PM, McGuinness UM, Aguayo AJ. Axons from CNS neurons regenerate into PNS grafts. Nature. 1980;284(5753):264-265.
  13. Ramón y Cajal S, DeFelipe J, Jones EG. Cajal’s Degeneration and Regeneration of the Nervous System. New York: Oxford University Press; 1991.
  14. Wong EV, David S, Jacob MH, Jay DG. Inactivation of myelin-associated glycoprotein enhances optic nerve regeneration. J Neurosci. 2003;23(8):3112-3117.
  15. McKerracher L, David S, Jackson DL, Kottis V, Dunn RJ, Braun PE. Identification of myelin-associated glycoprotein as a major myelin-derived inhibitor of neurite growth. Neuron. 1994;13(4):805-811.
  16. Vajda F, Jordi N, Dalkara D, et al. Cell type-specific Nogo-A gene ablation promotes axonal regeneration in the injured adult optic nerve. Cell Death Differ. 2015;22(2):323-335.
  17. Wang KC, Koprivica V, Kim JA, et al. Oligodendrocyte-myelin glycoprotein is a Nogo receptor ligand that inhibits neurite outgrowth. Nature. 2002;417(6892):941-944.
  18. Weibel D, Kreutzberg GW, Schwab ME. Brain-derived neurotrophic factor (BDNF) prevents lesion-induced axonal die-back in young rat optic nerve. Brain Res. 1995;679(2):249-254.
  19. Hannila SS, Siddiq MM, Carmel JB, et al. Secretory leukocyte protease inhibitor reverses inhibition by CNS myelin, promotes regeneration in the optic nerve, and suppresses expression of the transforming growth factor-beta signaling protein Smad2. J Neurosci. 2013;33(12):5138-5151.
  20. Silver J, Miller JH. Regeneration beyond the glial scar. Nat Rev Neurosci. 2004;5(2):146-156.
  21. Pearson CS, Mencio CP, Barber AC, Martin KR, Geller HM. Identification of a critical sulfation in chondroitin that inhibits axonal regeneration. Elife. 2018;7.
  22. Sahni V, Mukhopadhyay A, Tysseling V, et al. BMPR1a and BMPR1b signaling exert opposing effects on gliosis after spinal cord injury. J Neurosci. 2010;30(5):1839-1855.
  23. Li HJ, Pan YB, Sun ZL, Sun YY, Yang XT, Feng DF. Inhibition of miR-21 ameliorates excessive astrocyte activation and promotes axon regeneration following optic nerve crush. Neuropharmacology. 2018;137:33-49.
  24. Patel N, Poo MM. Orientation of neurite growth by extracellular electric fields. J Neurosci. 1982;2(4):483-496.
  25. Yamashita M. Electric axon guidance in embryonic retina: galvanotropism revisited. Biochem Biophys Res Commun. 2013;431(2):280-283.
  26. 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(10):3659-3668.
  27. McCaig CD. Studies on the mechanism of embryonic frog nerve orientation in a small applied electric field. J Cell Sci. 1989;93(Pt 4):723-730.
  28. Mesentier-Louro LA, Rosso P, Carito V, et al. 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(2):1056-1069.
  29. Mansour-Robaey S, Clarke DB, Wang YC, Bray GM, Aguayo AJ. Effects of ocular injury and administration of brain-derived neurotrophic factor on survival and regrowth of axotomized retinal ganglion cells. Proc Natl Acad Sci U S A. 1994;91(5):1632-1636.
  30. Feng L, Puyang Z, Chen H, Liang P, Troy JB, Liu X. Overexpression of brain-derived neurotrophic factor protects large retinal ganglion cells after optic nerve crush in mice. eNeuro. 2017;4(1).
  31. Giannaccini M, Pedicini L, De Matienzo G, Chiellini F, Dente L, Raffa V. Magnetic nanoparticles: a strategy to target the choroidal layer in the posterior segment of the eye. Sci Rep. 2017;7:43092.
  32. Chierzi S, Strettoi E, Cenni MC, Maffei L. Optic nerve crush: axonal responses in wild-type and bcl-2 transgenic mice. J Neurosci. 1999;19(19):8367-8376.
  33. Takahashi K, Yamanaka S. Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell. 2006;126(4):663-676.
  34. Aparicio JG, Hopp H, Choi A, et al. 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.
  35. Venugopalan P, Wang Y, Nguyen T, Huang A, Muller KJ, Goldberg JL. Transplanted neurons integrate into adult retinas and respond to light. Nature communications. 2016;7:10472.
  36. Johnson TV, Bull ND, Martin KR. Transplantation prospects for the inner retina. Eye (Lond). 2009;23(10):1980-1984.
  37. Yang TC, Chuang JH, Buddhakosai W, et al. Elongation of axon extension for human iPSC-derived retinal ganglion cells by a nano-imprinted scaffold. Int J Mol Sci. 2017;18(9).
  38. Fang Y, Mo X, Guo W, et al. A new type of Schwann cell graft transplantation to promote optic nerve regeneration in adult rats. J Tissue Eng Regen Med. 2010;4(8):581-589.
  39. 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(11):692-697.
  40. Ackman JB, Crair MC. Role of emergent neural activity in visual map development. Curr Opin Neurobiol. 2014;24(1):166-175.