Ординатура / Офтальмология / Английские материалы / Eye, Retina, and Visual System of the Mouse_Chalupa, Williams_2008

the only neurons that were found to elaborate neurites were those that did not express the alkaline phosphatase marker from the mutated Brn-3c locus, which suggests that, in RGCs that do not normally express Brn-3c, alternative transcription factors drive axon outgrowth. Thus, transcription factors involved in cell fate determination, which is often defined by cell morphology, may act directly via the control of axon and dendrite growth ability.

What are the genes downstream of Brn-3b that could effect changes in the intrinsic axon growth state of a neuron? Recently, Brn-3b−/− retinas were compared with wild-type retinas in microarray experiments to determine which genes depend on Brn-3b for their expression, and several genes linked to axon guidance were found to be downregulated in Brn-3b−/− retinas (Mu et al., 2004). Furthermore, the NGF receptor Trk-A was identified as a transcriptional target of Brn-3a (Ma et al., 2003) providing a direct link from transcription factor expression to trophic responsiveness. These experiments point to the power of using focused microarray experiments in relation to established phenotypic models.

Transcription factors are not the only master regulators in a cell. Levels of specific complements of proteins can be regulated posttranslationally by ubiquitination and degradation. Because there are hundreds of ubiquitin ligases in mammalian genomes, the potential for subcellular regulation of the levels of specific proteins is vast. Recently, exciting data have demonstrated that the ubiquitin ligase anaphasepromoting complex (APC) has a significant role in regulating the axon growth rate of cerebellar granule neurons. APC was originally found to be essential for mitosis in dividing cells, but it is also highly expressed in postmitotic neurons. Blocking APC or its activator Cdh1 with RNAi or dominant negative approaches led to a greater than twofold increase in axon length but not to a change in dendrite length, suggesting that Cdh1-APC complexes normally inhibit axon but not dendrite growth (Konishi et al., 2004). Furthermore, APC normally binds to and leads to the proteasomal degradation of the transcriptional corepressor SnoN, and knocking down SnoN itself led to a decrease in axon growth (Stegmuller et al., 2006). What are the targets of SnoN that regulate axon growth? Whether these specific proteins will be relevant for axon growth or regeneration in RGCs remains to be examined, but the principle of illuminating such regulatory pathways is very exciting.

How does the intrinsic state of neuron allow the same extrinsic cues to induce axon or dendrite growth preferentially (see figure 33.5)? Cytoplasmic kinases or phosphatases may provide an axonor dendrite-specific interpretation of extrinsic cues. For example, electrical activity seems to be crucial to potentiate both axon and dendrite growth in various neurons in response to trophic signals (Goldberg, 2003). The influx of Ca2+ during depolarization activates a family of Ca2+-dependent signaling kinases, calmodulin

kinases (CaMKs), which have now been implicated in both axonand dendrite-specific growth. For axon growth, overexpression of a dominant-negative variant of CaMKI in embryonic hippocampal and postnatal cerebellar granule neurons decreases axon outgrowth, whereas expression of cytosolic dominant-negative CaMKII or nuclear dominantnegative CaMKIV constructs has no effect on axon growth in these cells (Wayman et al., 2004). In other studies, CaMKII-β and nuclear CaMKIV have been found to stimulate dendrite but not axon growth in hippocampal or cortical cultures (Fink et al., 2003; Redmond et al., 2002). Thus, CaMK isoforms can differentially control axonand dendrite-specific outgrowth in hippocampal neurons. Further experiments should point to mechanisms of either developmental or subcellular regulation, or both, for these kinases.

The levels of second messengers such as cAMP and cGMP, if intrinsically maintained, could affect axon and dendrite growth differentially. As discussed earlier, cAMP is crucial for RGCs to extend axons in response to neurotrophic factors (Goldberg et al., 2002a), and a developmental decrease in cAMP levels has been proposed to underlie the developmental loss of axon regeneration in the presence of inhibitory myelin-associated cues (Cai et al., 2001). In a strongly trophic, noninhibitory environment, however, an increase in cAMP does not revert the slow axon growth of postnatal RGCs to the fast growth of their embryonic state (Goldberg et al., 2002b).

Thus, critical questions remain. What is the molecular basis for the developmental loss of intrinsic axon growth ability in RGCs? And how does such a loss relate to the failure of RGCs to regenerate in vivo?

Axon regeneration in the mature visual system

Axons that get cut either by injury or disease in the PNS reinitiate the whole process of axon growth, elongating back to their peripheral targets and restoring sensorimotor function. Yet when axons in the adult mammalian CNS are severed, they largely fail to regenerate. This appears to occur independent of the mode of axon injury, whether traumatic, ischemic, immunologic, or degenerative: in all cases, RGCs fail to regenerate into the optic nerve. Furthermore, RGCs (and other CNS neurons) typically die after axon injury. After an optic nerve crush injury behind the eye, 95% of postnatal rodent RGCs die within 2–3 days, and a similar number die in the adult rodent within 2 weeks (Berkelaar et al., 1994; Villegas-Perez et al., 1993). Thus, there are multiple issues to address when considering approaches to improve RGC regeneration and survival and axon growth past the injury site first, and later guidance and target innervation.

Although it was once believed that adult CNS neurons intrinsically lacked any ability to regenerate, this view was

disproved 25 years ago by elegant experiments demonstrating that at least some RGCs could regenerate their axons through fragments of peripheral nerve grafts (So and Aguayo, 1985). The failure of RGCs and other CNS neurons to regenerate their axons is generally ascribed to an inhibitory glial environment; however, the lack of successful regeneration in experiments designed to overcome inhibitory CNS glial cues also hints at other extrinsic or intrinsic regulation of axon growth.

How Is Retinal Ganglion Cell Regeneration Studied? Although traumatic optic neuropathy is one of the least common human scenarios in which RGC axons are injured, it has been the best studied model in rodents by far. Typically, the optic nerve is surgically cut or crushed within the orbit behind the eye, although experiments crushing the nerve intracranially have also been studied. The regenerative response is assayed by labeling RGC axons and determining whether any extend past the lesion site into the optic nerve (figure 33.6), or into a peripheral nerve graft in experiments styled after Aguayo’s early work (So and Aguayo, 1985). Interestingly, although injuring the optic nerve at increasing distances from the eye delays the onset of RGC cell death (Berkelaar et al., 1994), there may be less regenerative response with more distal injury (You et al., 2000).

Other optic neuropathies are also beginning to be modeled in rodents. In optic neuritis, optic nerve oligodendrocytes demyelinate from RGC axons in concert with an autoimmune reaction. This has been modeled in rat and mouse models of experimental autoimmune encephalomyelitis, in which rodents’ immune system is stimulated to react against

Figure 33.6 After an optic nerve crush injury, RGC axons were labeled with an intravitreal injection of fluorophore-conjugated cholera toxin B. The bright axons failed to regrow across the lesion site (arrows) after 8 days in vivo. Optic nerve nuclei are counterstained with a nuclear dye, DAPI. See color plate 23. (Y. Duan and J. L. Goldberg. [2008]. Unpublished data.)

myelin-associated proteins. Immunization against myelinoligodendrocyte glycoprotein (MOG) induces multiple sclerosis-like demyelination throughout the brain, including in the optic nerve (Storch et al., 1998), but creating transgenic mice with T cells directed against MOG, or passively transferring anti-MOG T cells in rats, creates demyelinating disease largely confined to the optic nerve, mimicking optic neuritis (Bettelli et al., 2003; Shao et al., 2004). In such models, RGC axons are incidentally severed and fail to regenerate, and, as after optic nerve trauma, RGCs die with a 1- to 2-week delay (Guan et al., 2006).

Ischemic optic neuropathy—a stroke of the optic nerve— is exceedingly common in humans, and taking an old technology for creating strokes in the brain and applying it to the optic nerve has allowed the study of ischemic optic neuropathy in rodents. In this photothrombotic model, the photosensitizing dye rose bengal is injected into the tail vein and the surgically exposed optic nerve is illuminated with a 514 nm or 535 nm wavelength laser. Photoactivation of the intravascular rose bengal leads to free radical generation, endothelial damage, and platelet degranulation, creating a focal thrombosis in the optic nerve. This model has been studied recently to examine the RGC and optic nerve response to ischemic axon injury (Bernstein et al., 2003), and future work may be directed at enhancing RGC survival and regeneration in such models.

Finally, the second most common cause of irreversible blindness in the world is glaucomatous optic neuropathy. Glaucoma is a neurodegenerative disease, and lowering intraocular pressure often slows its progression. Glaucomatous optic neuropathy has been extensively studied in rodent models (reviewed in Morrison et al., 2005; Whitmore et al., 2005), although the primary concern in these studies is usually RGC survival, not RGC axon regeneration, and as such is not considered in this chapter.

Intrinsic and Extrinsic Regulation of Adult Retinal

Ganglion Cell Regeneration Since RGCs do not extend axons in the absence of specific extracellular signals, regenerative failure might be explained in part by a relative inability of mature CNS astrocytes and oligodendrocytes to secrete trophic signals after injury. Indeed, ample evidence exists to support a role for neurotrophins in stimulating axon growth in vivo. The same peptide trophic factors that stimulate RGC axon growth in vitro enhance RGC survival and axon regeneration in vivo (Aguayo et al., 1996; Cui et al., 1999; Yip and So, 2000). Furthermore, as was seen for RGCs in vitro (Goldberg et al., 2002a), elevation of cAMP levels in RGCs enhances their regenerative ability after optic nerve injury in vivo (Cui et al., 2003; Monsul et al., 2004; Watanabe et al., 2003).

Our understanding of the mechanisms of axon growth has led to progress in understanding regenerative failure. For

example, understanding that the regulation of both repulsion and collapse of growth cones by inhibitory molecules relies on the same intracellular signaling pathways suggests that manipulating these common regulators can increase regeneration. Thus, blocking the growth cone collapsing activity of the small GTPase rho increases RGC regeneration in the inhibitory environment of the optic nerve in vivo (Lehmann et al., 1999), an effect that may be caused in part by blockade of inhibitory signaling at the growth cone, but also in part by increasing the intrinsic growth ability of the neurons themselves. Such manipulations may be combined with cAMP elevation and trophic factor delivery to enhance regeneration even further (Hu et al., 2006).

Does the loss of intrinsic axon growth ability by RGCs (Goldberg et al., 2002b) contribute to their failure to regenerate after injury in the adult? As mentioned earlier, a loss of intrinsic axon growth ability could explain why in many previous experiments, regeneration proceeded remarkably slowly, even when glial inhibitory cues were removed. For example, most RGCs take 2–3 months to regenerate through peripheral nerve grafts to the superior colliculus (Aguayo et al., 1987; Bray et al., 1987), although the fastest RGCs may extend 1–2 mm/day into peripheral nerve grafts. This is approximately the rate they extend axons in vitro (Goldberg et al., 2002b), and far slower than the 10 days they would take if they elongated their axons at an embryonic growth rate of 10 mm/day. It is not clear whether this developmental switch is reversible either: soluble signals from optic nerve glia or peripheral nerve glia, or from retinal or superior collicular cells, were not able to reverse the loss of rapid axon growth ability in RGCs in vitro, suggesting that the developmental switch may normally be permanent (Goldberg et al., 2002b). There remains the possibility, however, that other signals, or the discovery and manipulation of genes involved in this transition, may revert the postnatal neurons to their embryonic axon growth ability, and that this may be critical to increase regeneration in the CNS. Thus, a combined approach may be required—both the intrinsic neuronal growth state and the extrinsic environment may have to be optimized for successful regeneration after injury.

Conclusion

Our understanding of how RGC axons grow and the regulation of RGC axon regeneration is still in its infancy. Although a great deal of progress has been made recently in understanding the nature of the extracellular signals that induce axon growth, we still know relatively little about the intracellular molecular mechanisms by which these signals are transduced into the neuron and ultimately how they elicit growth. Fortunately, new molecular tools, including genomics, proteomics, and RNAi, should help us elucidate novel components of the axon growth machinery that couple

transmembrane signaling receptors at the growth cone to the axonal cytoskeleton. In particular, these methods should soon reveal the underlying transcriptional program elicited by peptide trophic factors that triggers axon growth. Determining the full roster of genes induced during axon growth, which genes are needed for axon elongation both during development and for regeneration, and whether developmental differences in intrinsic axon growth ability between embryonic and postnatal RGCs underly the failure of RGC regeneration should help in developing new approaches to enhancing RGC regeneration in the mouse and, ultimately, in the human visual system.

acknowledgments Portions of this chapter are reprinted with permission from earlier publications (Goldberg, 2003, 2004). Figures 33.1 to 33.5 and their accompanying captions are reprinted by permission of the publishers. Figure 33.6 was generated in experiments supported by a grant from the National Eye Institute.

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Much of our present understanding of the mechanisms underlying the development of sensory connections is based on work done in the mammalian retinogeniculate pathway. In recent years, the mouse has come to the forefront as a model system in which to study visual system development, largely because modern molecular biology allows targeted genetic manipulation. The advent of transgenic mouse models has brought forth abundant new information about the molecular mechanisms involved in early pathfinding, visual map formation, and the subsequent activity-depen- dent refinement of connections. Although other vertebrate systems also allow genetic dissection, the mouse is particularly well suited for such study because its visual system has some of the rudimentary features found in higher mammals, including humans (see chapter 21).

With the emergence of mouse models, basic information about the structural and functional composition of the developing retinogeniculate pathway is needed. This chapter provides a detailed examination of the changes that occur during late prenatal and early postnatal life, when retinal axons innervate the lateral geniculate nucleus (LGN) and establish and then rearrange their connections with relay cells to form adult patterns of connectivity. This review focuses largely on studies done in a common pigmented strain (C57BL/6) and on the neural elements and related events associated specifically with the retinogeniculate pathway. Topics addressed include the pattern of retinal innervation in the LGN, the structural and functional composition of relay cells and interneurons, their associated patterns of connectivity, and the potential mechanisms underlying the remodeling of connections. The LGN also receives rich innervation from a variety of nonretinal sources, including the brainstem, thalamic reticular nucleus, and layer VI of visual cortex. Virtually nothing is known about how such nonretinal circuitry develops or how such input contributes to the maturation of retinogeniculate connections. Therefore, these aspects of LGN circuitry, while important, are not discussed.

The development of eye-specific segregation in the lateral geniculate nucleus

The topographic representation of visual fields in the retina and central visual targets is a hallmark feature of vision. Visuotopic maps are defined by an orderly series of connections that link neighboring retinal ganglion cells (RGCs) with neighboring neurons in their primary targets, such as the dorsal LGN of thalamus. For example, in the mouse, the nasal-temporal visual axis maps in a medial to lateral plane of the LGN, while upper to lower visual fields map in a dorsal to ventral direction (Grubb et al., 2003; Wagner et al., 2000). A defining feature of these maps is the segregation of inputs from the two eyes. In the LGN of carnivores and primates, retinal projections from the two eyes are divided by cytoarchitectonic laminae. LGN cells within each lamina receive monocular input from the contralateral or ipsilateral eye. However, as is the case with many nocturnal rodents, the LGN of the mouse lacks an obvious lamination pattern (figure 34.1A; Reese, 1988; Van Hooser and Nelson, 2006). Instead, retinal projections are organized into nonoverlapping territories called eye-specific domains that can be visualized only with anterograde labeling of RGCs (figure 34.1B; Godement et al., 1984; Jaubert-Miazza et al., 2005).

In the adult mouse, axons from nasal retina and most of temporal retina cross at the optic chiasm and project contralaterally to the lateral and ventral regions of the LGN. Since the mouse has laterally placed eyes and poor binocular vision, in pigmented strains, the majority of retinal fibers (95%) cross at the optic chiasm (Dräger and Olsen, 1980). Crossed projections representing the contralateral eye occupy as much as 85%–90% of the total area in LGN. A much smaller group of RGCs (5%) that arise from the ventrotemporal region, known as the temporal crescent, have axons that do not cross at the optic chiasm but instead project ipsilaterally and terminate in the anteromedial region of the LGN. These uncrossed projections representing the ipsilateral eye form a patchy cylinder that runs through the

Figure 34.1 Anatomical organization of the developing LGN in the C57BL/6 mouse. Neonatal and adult features are shown. A, Coronal sections through the LGN with a Nissl stain. At early postnatal ages (P7), the LGN can be distinguished from the intrageniculate leaflet (IGL) and ventral geniculate nuclei (VLG). The cytoarchitecture of the LGN lacks an obvious eye-specific laminar pattern. Scale bar = 100 μm. B, Anterograde labeling of retinal projections with fluorescent conjugates of cholera toxin β subunit reveals eye-specific patterning in LGN. Shown are coronal sections through the same section of LGN. The injection of two different fluorescent conjugates reveals both the contralateral eye (crossed, images at top) and ipsilateral eye (uncrossed, images at bottom) projections. At early ages, projections show substantial overlap. In the adult, uncrossed projections form a nonoverlapping patch that is confined to the anteromedial region of LGN. Scale bars = 100 μm. C, Camera lucida reconstructions of biocytin-filled LGN cells.

rostrocaudal axis of LGN and occupies about 10%–15% of the nucleus. This form of eye-specific patterning is not apparent during the early stages of target innervation and visual map formation but emerges sometime near the end of the first postnatal week (figures 34.1B and 34.2A; see also figure 34.5; Godement et al., 1984; Jaubert-Miazza et al., 2005). Initially, crossed and uncrossed fibers innervate the LGN at different times, with crossed projections arriving

Shown are two filled relay cells (P7 and adult) and one adult interneuron. Relay cells and interneurons are readily distinguished. Developing relay cells have smaller somata and fewer arbors with shorter branches. Scale bar = 20 μm. D, EM micrographs showing the synaptic structure in LGN. At P7 (left), synapses can be identified (arrows), but it is not possible to categorize their composite terminals on the basis of their ultrastructure. At older ages (>P14), the ultrastructure of retinal terminals (RLP), the dendrites (D) of relay cells, and the dendritic terminals (F2) of interneurons are distinct. Note the triadic arrangement of these elements. Axon terminals (F1) of interneurons also make synaptic contact with the dendrites of relay cells. F1 and F2 GABAergic profiles are immunostained with gold particles. Scale bars = 1 μm. (A, B, and C, Adapted from Jaubert-Miazza et al., 2005. D, Adapted from Guido et al., 2008.)

earlier (at E16, about 5 days before birth) than uncrossed ones (P0). At these perinatal ages, crossed retinal projections span almost the entire LGN. Uncrossed projections also begin to fill the LGN in a widespread manner, but by P2 a rudimentary patch of terminal arbors is evident in the anteromedial sector. Between P2 and P5, the inputs from the two eyes still share a substantial amount of terminal space. By P7, retinal projections from the two eyes begin to

A

contra ipsi superimposed

Figure 34.2 Retinogeniculate axon segregation in the developing mouse. A, Anterograde transport of CTβ conjugated to Alexa Fluor 594 (red) labels contralateral eye projections, and anterograde transport of CTβ conjugated to Alexa Fluor 488 (green) labels ipsilateral eye projections. Panels from left to right depict red and green fluorescence labeling of the same section of LGN. Adjacent to these are the superimposed fluorescent pattern and corresponding pseudo-colored image, where pixel intensity is assigned a single value above a defined threshold. Pixels that contain both red and green fluorescence are considered areas of overlap and are represented in yellow. Scale bar = 100 μm. B, Pixel intensity analysis reveals degree of eye-specific segregation in the developing LGN. Top, Scatterplots of pixel intensity for a single section through LGN at P3 and P28. Each point represents a pixel in which the fluorescence intensity of the ipsilateral projection is plotted against the intensity of the contralateral projection. At P3, when projections overlap, pixel intensities show a positive correlation. At P28, pro-

jections are segregated and pixel intensities show a negative correlation. Bottom, Corresponding R distributions of pixel intensity. For each pixel the logarithm of the intensity ratio (R = log10 FI/FC) is plotted as a frequency histogram (bin size = 0.1 log units). A narrow R distribution (P3) shows an unsegregated pattern; a wide one (P28) shows a segregated pattern. C, Summary plots showing the spatial extent of retinal projections (left) and the variance of R distributions (right) at different ages. Left, Percent area in LGN occupied by contralateral, ipsilateral, and overlapping terminal fields at different ages. Each point represents the mean and SEM for a group of same-aged animals. Note that ipsilateral projections and the degree of overlap recede between P3 and P12. Right, Mean and SEM variance values obtained from R distributions. Changes in spatial extent are accompanied by a parallel increase in variance and reflect a progressive increase in the degree of eye-specific segregation between P3 and P12. See color plate 24. (Adapted from Jaubert-Miazza et al., 2005.)