Development and Aging of the Vertebrate Visual System

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Development and Aging of the Vertebrate Visual System

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Dietrich E. Lorke, Al Ain

75 figures, 1 in color, and 4 tables, 2007

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Vol. 4, No. 1–2, 2006–07

Contents

5 Development and Aging of the Vertebrate Visual System. Introduction Lorke, D.E. (Al Ain/Hamburg) 8 Aqueous Humor Formation and Its Regulation by Nitric Oxide:

A Mini Review Do, C.W. (Hong Kong); Kong, C.W. (Hong Kong/New York, N.Y.); Chan, C.Y.; Lam, C.; To, C.H. (Hong Kong) 13 Revisiting the Trabecular Meshwork in the Eyes of the Developing Human Chan, C.Y.; Li, W.W.Y.; Pang, C.C.P.; Lam, D.S.C.; Yew, D.T. (Hong Kong) 19 Morphological and Neurochemical Development of the Human Neural

Retina Nag, T.C.; Wadhwa, S. (New Delhi) 31 Review of Development and Aging in the Eye of Teleost Fish Kunz, Y.W. (Dublin) 61 The Pathology of Mueller Cells under Different Disease Conditions Nishikawa, S. (Tokorozawa) 76 Disruption of Sonic Hedgehog Signaling Affects Axon Routing in the

Mouse Optic Chiasm Hao, Y. (Hong Kong/Guangdong); Wang, J. (Hong Kong/Beijing); Chan, C.K.; Chan, S.O. (Hong Kong) 85 Aging of the Mammalian Circadian Timing System: Changes in the

Central Pacemaker and Its Regulation by Photic and Nonphotic Signals Duncan, M.J. (Lexington, Ky.) 102 Extrapineal Melatonin: General Conception, Localization and Biological

Role in the Visual System Kvetnoy, I.M.; Smirnova, I.O.; Polyakova, V.O. (St. Petersburg) 112 Melanopsin in Lower Vertebrates: The Race Is On to Show that It Is Really

There Meyer-Rochow, V.B. (Bremen/Oulu) 115 Author Index Vol. 4, No. 1–2, 2006–07 116 Subject Index Vol. 4, No. 1–2, 2006–07

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Neuroembryol Aging 2006–07;4:5–7 DOI: 10.1159/000103580

Published online: June 5, 2007

Development and Aging of the Vertebrate Visual System Introduction

Dietrich E. Lorke Department of Anatomy, Faculty of Medicine and Health Sciences, United Arab Emirates University, Al Ain, United Arab Emirates; Institute of Anatomy II, Experimental Morphology, Research Group Neuroembryology, University Hospital Hamburg-Eppendorf, Hamburg, Germany

Key Words Aging
Development
Melatonin
Optic nerve
Retina

Abstract This special issue encompasses articles dealing with a number of unique features of the vertebrate visual system. These characteristics make it an established model to study the molecular and genetic control of axonal growth, neurogenesis, gliogenesis and regeneration in the central nervous system. The clinical relevance of developmental malformations and age-related diseases is briefly described. The possible effects of visual inputs on melatonin secretion and changes in this system during aging are summarized. Copyright © 2007 S. Karger AG, Basel

Developmental malformations and age-related diseases of the visual system, including cataract, age-related maculopathy, glaucoma and diabetic retinopathy, are very common and potentially devastating sight-threatening conditions. This special issue of Neuroembryology and Aging is devoted to several aspects of the development and aging of the vertebrate visual system. The first two articles deal with the posterior and anterior chambers of the eye, which are, at least partly, mesenchymal derivatives of the neural crest. In their review on the regulation of aqueous humor formation, Do et al. [pp 8–12] focus on the mechanisms whereby nitric oxide

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modulates the production of aqueous humor. Chan et al. [pp 13–18] describe changes in the collagen and elastic fiber distribution in the trabecular meshwork during the development of the human fetus. They show that collagen fibers are relatively early present, whereas elastic fibers appear much later in this part of the eye, which plays an important role in the drainage of aqueous humor. The clinical relevance of these two articles is obvious, because imbalance between production and drainage of aqueous humor may lead to elevated ocular pressure and glaucoma, which is a principal cause of blindness. However, the visual system not only performs essential sensory functions and plays an important clinical role, it also has several unique characteristics, which make it an ideal model system for neurobiological and neuroembryological studies [1–4]. The retina is composed of six distinct types of neurons and a unique type of glial cell, which can be easily identified by their position in well-defined layers and by their characteristic morphology. Light stimulates photoreceptors (rods and cones) which transmit their signals via bipolar cells to ganglion cells. Horizontal and amacrine cells modulate this direct signaling pathway. Ganglion cell axons transmit the result of all information processing along the optic nerve to various target areas in the brain. The retina and the optic nerve develop from the optic cup, an evagination of the forebrain vesicle, and therefore represent parts of the central nervous system (CNS). Simplicity and accessibility have made the retina a model for

Dietrich E. Lorke Department of Anatomy Faculty of Medicine and Health Sciences UAE University, Box 17666, Al Ain (United Arab Emirates) Tel. +971 3 7137 497, Fax +971 3 7672 033, E-mail [email protected]

the study of the genesis of neuronal diversity [4]. In addition to molecular and genetic control of the highly coordinated and precisely regulated cell proliferation in the germinative neuroepithelium, the mechanisms by which multipotent progenitor cells give rise to different neuronal and glial cell types in a characteristic order of birth have been analyzed in the retina as a model system [4– 10]. The development of the human retina is summarized in the article by Nag and Wadhwa [pp 19–30]. Histogenesis of the human retina is a relatively complex process, because, in addition to neurons and glia, it also contains blood vessels whose formation is closely related to neuronal maturation. This article describes the temporospatial sequence of neuronal differentiation, synaptogenesis and vascular development. In addition, it discusses the appearance of numerous neurotransmitters and neuromodulators during specific developmental stages. In contrast to the mammalian CNS, the retina and the optic nerve of teleost fish grow throughout life and retain the ability to regenerate. New ganglion cells, as well as all other cell types, are continuously added to the existing retina in a peripheral growth zone. Their axons grow towards the optic tectum along the already myelinated, functional nerve fibers. The teleost visual system is therefore an excellent tool to study axonal growth and regeneration [11]. In recent years, the zebrafish visual system has become the model system for the genetic analysis of vertebrate CNS development and function. Zebrafish are very fertile and develop rapidly; their embryos are transparent, and some of the genetic defects of the zebrafish retina resemble human retinopathies [12, 13]. Kunz [pp 31–60] outlines the development and aging of the eye of teleosts. She specifies developmental stages of the eye in different teleost species, including zebrafish, describes growth zones in the developing and fully differentiated teleostean retina and discusses the role of ultraviolet-sensitive cones for the protection of the retina during early life. Müller cells represent a unique type of glia present in the retina. Their cell bodies lie in the middle of the inner nuclear layer and their processes span its entire thickness, reminiscent of radial glia, which in most other parts of the CNS is only found during development. Müller cells play an important role in neurotransmitter metabolism due to their ability to take up and metabolize glutamate and GABA. Nishikawa [pp 61–75] describes the development of this particular cell type and its involvement in retinal degeneration and aging.

6

Neuroembryol Aging 2006–07;4:5–7

The optic nerve as a derivative of the optic stalk is an ideal system for studies of development and regeneration of CNS white matter. Being only composed of axons (from one specific source) and glia, it is devoid of neuronal cell bodies and therefore an invaluable model to study axonal growth, glial cell lineage, gliogenesis, myelination and regeneration in the CNS [14–21]. Using the optic nerve as a model of axonal path finding in the CNS, Hao et al. [pp 76–84] examine the importance of an axon guidance molecule, Sonic Hedgehog (Shh), in determining the axon course in the mouse optic chiasm. Shh, originally discovered as a protein whose mutation created spiny backs in fruit flies, was named after Sonic the Hedgehog, a very popular video game hero in the 1990s. It is now known to determine neuronal fate also during early mammalian CNS development and to affect axon routing in the chick visual system [22]. Using the technique of embryonic brain slices, Hao et al. demonstrated that blocking the signaling of this molecule leads to marked misrouting of axons, indicating that, also in mammals, Shh significantly contributes to path finding of axons crossing the midline. The task of the vertebrate visual system is not confined to the perception of a three-dimensional image of the surrounding environment, it also plays a central role in adjusting the internal circadian clock to the external lightdark cycle. Ocular light transmits time-of-day information to the hypothalamic suprachiasmatic nucleus, the brain’s biological clock, which in turn suppresses melatonin production by the pineal gland. Thereby, secretion of melatonin is adjusted to the external light-dark cycle. Melatonin is well known as a hormone which influences seasonal reproduction, induces sleep and potentially has antioxidant effects [23–26]. Duncan [pp 85–101] reviews age-related changes in the mammalian circadian timing system. Although there is little evidence for neurodegeneration in the suprachiasmatic nucleus, the neurons of this circadian pacemaker show numerous functional alterations with increasing age. In addition, expression of genes relevant to the circadian clock is decreased. The pineal gland is not the only source of melatonin. Kvetnoy et al. [pp 102–111] review these extrapineal sites of melatonin production and their physiological significance. Melatonin-producing cells are found in a central and a peripheral compartment. The central compartment consists of cells associated with the visual system and the pineal gland, whereas the peripheral compartment comprises almost all organ systems, including enterochromaffin cells of the gut. During aging, melatonin Lorke

production decreases both in the pineal gland and in extrapineal melatonin-producing enterochromaffin cells of the gut, and this may play a role in age-related circadian rhythm disorders. Melatonin biosynthesis must, at least in part, be controlled by photoreceptors other than the known rods and cones, because light-induced melatonin suppression is observed in humans and mice lacking these two receptors. A new photosensitive pigment chemically related to opsin, called ‘melanopsin’, has been detected in certain ganglion cells of the mammalian and avian retina, which may constitute a non-image-forming photosensitive system in the mammalian eye regulating circadian timing. In his commentary, Meyer-Rochow [pp 112–114] addresses the question of whether melanopsin also plays a role in lower vertebrates. He has examined the eyes of an Antarctic teleost fish, Trematomus bernacchii, by immunohistochemistry using an antibody directed against

rhodopsin and another one against a broad range of opsin molecules, including visinin. Binding of the latter antibody is observed not only in the rhodopsin-containing outer rods, but also in a few scattered cells in the ganglion cell layer. Based on the presence of melanopsin in a similar location in the mammalian retina, he concludes that melanopsin or another opsin-like photopigment may also be involved in regulating biorhythms in these teleost fish. In summary, these articles give insight into the broad range of studies dealing with the development and aging of the vertebrate visual system, which have made it the most thoroughly characterized part of the CNS [4]. This research elucidates fundamental mechanisms governing the complex cellular interactions during neurogenesis, differentiation and regeneration of the entire vertebrate CNS.

References 1 Choy KW, Baum L, Lam DSC, Pang CP: Molecular genetic control of retinal development. Neuroembryology 2002; 1:54–60. 2 Dyer MA, Cepko CL: Regulating proliferation during retinal development. Nat Rev Neurosci 2001;2:333–342. 3 Stone J, Maslim J, Valter-Kocsi K, Mervin K, Bowers F, Chu Y, Barnett N, Provis J, Lewis G, Fisher SK, Bisti S, Gargini C, Cervetto L, Merin S, Peer J: Mechanisms of photoreceptor death and survival in mammalian retina. Prog Retin Eye Res 1999;18:689–735. 4 Cepko CL, Austin CP, Yang X, Alexiades M, Ezzeddine D: Cell fate determination in the vertebrate retina. Proc Natl Acad Sci USA 1996;93:589–595. 5 Lorke DE, Winking H: Histogenesis of the retina in murine trisomy 19. Brain Res Bull 1986;16:845–851. 6 Lorke DE, Engelmann E: Quantitative and morphological studies of retinal development in mice with trisomy 19. Acta Anat (Basel) 1988;131:56–65. 7 Provis JM: Development of the primate retinal vasculature. Prog Retin Eye Res 2001;20: 799–821. 8 Mey J, Thanos S: Development of the visual system of the chick. I. Cell differentiation and histogenesis. Brain Res Brain Res Rev 2000;32:343–379.

Introduction

9 Wu Y, Lorke DE, Lai H, Wai SM, Kung LS, Chan WY, Yew DTW: Critical periods of eye development in vertebrates with special reference to humans. Neuroembryology 2003; 2:1–8. 10 Wai MS, Lorke DE, Kung LS, Yew DT: Morphogenesis of the different types of photoreceptors of the chicken (Gallus domesticus) retina and the effect of amblyopia in neonatal chicken. Microsc Res Tech 2006; 69: 99– 107. 11 Mack AF, Wolburg H: Growing axons in fish optic nerve are accompanied by astrocytes interconnected by tight junctions. Brain Res 2006;1103:25–31. 12 Neuhauss SC: Behavioral genetic approaches to visual system development and function in zebrafish. J Neurobiol 2003; 54:148–160. 13 Avanesov A, Malicki J: Approaches to study neurogenesis in the zebrafish retina. Methods Cell Biol 2004; 76:333–384. 14 Lorke DE, Lauer M: Gliogenesis and myelination in the optic nerve of trisomy 19 mice. A quantitative electron-microscopic study. Acta Anat (Basel) 1990;137:222–233. 15 Tsai HH, Miller RH: Glial cell migration directed by axon guidance cues. Trends Neurosci 2002;25:173–175. 16 Vaughn JE: An electron microscopic analysis of gliogenesis in rat optic nerves. Z Zellforsch 1969;94:293–324. 17 Valat J, Privat A, Fulcrand J: Multiplication and differentiation of glial cells in the optic nerve of the postnatal rat. A reassessment. Anat Embryol (Berl) 1983;167:335–346.

18 Skoff RP, Price DL, Stocks A: Electron microscopic autoradiographic studies of gliogenesis in rat optic nerve. I. Cell proliferation. J Comp Neurol 1976;169:291–312. 19 Raff MC, Abney ER, Miller RH: Two glial cell lineages diverge prenatally in rat optic nerve. Dev Biol 1984;106:53–60. 20 Raff MC, Durand B, Gao FB: Cell number control and timing in animal development: the oligodendrocyte cell lineage. Int J Dev Biol 1998;42:263–267. 21 Aguayo AJ, David S, Bray GM: Influences of the glial environment on the elongation of axons after injury: transplantation studies in adult rodents. J Exp Biol 1981;95:231–240. 22 Charron F, Tessier-Lavigne M: Novel brain wiring functions for classical morphogens: a role as graded positional cues in axon guidance. Development. 2005;132:2251–2262. 23 Arendt J: Melatonin: characteristics, concerns, and prospects. J Biol Rhythms 2005; 20:291–303. 24 Reiter RJ: Melatonin: clinical relevance. Best Pract Res Clin Endocrinol Metab 2003; 17: 273–285. 25 Cajochen C, Krauchi K, Wirz-Justice A: Role of melatonin in the regulation of human circadian rhythms and sleep. J Neuroendocrinol 2003;15:432–437. 26 Hannibal J: Roles of PACAP-containing retinal ganglion cells in circadian timing. Int Rev Cytol 2006;251:1–39.

Neuroembryol Aging 2006–07;4:5–7

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Neuroembryol Aging 2006–07;4:8–12 DOI: 10.1159/000103581

Published online: June 6, 2007

Aqueous Humor Formation and Its Regulation by Nitric Oxide: A Mini Review Chi Wai Do a Chi Wing Kong a, c Chu Yan Chan b Chuen Lam a Chi Ho To a a

Laboratory of Experimental Optometry, School of Optometry, Hong Kong Polytechnic University, and Department of Anatomy, Chinese University of Hong Kong, Hong Kong, SAR, China; c Department of Ophthalmology, Mount Sinai School of Medicine, New York, N.Y., USA b

Key Words cGMP
Chloride transport
Ciliary epithelium
Glaucoma
Intraocular pressure

Abstract Glaucoma is a common and severe aging eye disease which can cause permanent visual loss if untreated. One common strategy of glaucoma treatment is to lower the intraocular pressure (IOP) of the eye. So far, the pharmacologic reduction of IOP is the only proven intervention that can delay the progression of glaucomatous damage. Lowering the IOP can be achieved either by reducing the rate of aqueous humor secretion or by facilitating the rate of aqueous drainage. The formation of aqueous humor is driven primarily by vectorial ion transport, mainly Cl–, across the ciliary epithelium from ciliary stroma into the posterior chamber, resulting in the generation of an osmotic gradient which drives water movement. Many ion transporters and channels have been shown to participate in the process of aqueous secretion. Among several other signaling mechanisms, the nitric oxide (NO) signaling cascade is thought to regulate the ion transport across the ciliary epithelium, thereby reducing the rate of aqueous humor formation and IOP. Therefore, in this review, the potential significance of NO in the regulation of aqueous humor formation and IOP will be discussed.

Introduction

Vision is enormously important as it accounts for 38% of the total sensory input to the brain [1]. Vision can be impaired permanently by a number of conditions. Glaucoma is a severe, sight-threatening disease which is the leading cause of irreversible blindness in the world [2]. Glaucoma is an age-related disease, being more common in the elderly. The prevalence of glaucoma in people aged 40 and over is approximately 2% [2]. As the global population continues to age, it is estimated that the number of glaucoma patients will be increased by 50% by year 2020 [3]. People with glaucoma often have elevated intraocular pressure (IOP) of the eye and therefore the major current therapy is to lower the IOP. This is usually achieved initially by drugs but if the hypotensive response is not satisfactory, surgery is then considered. This hypotensive treatment is so far the only proven method to retard glaucomatous damage [4, 5]. However, due to the facts that some patients may eventually develop resistance to these drugs and these agents do have significant side effects, more potent and selective pharmacologic agents are urgently needed.

Copyright © 2007 S. Karger AG, Basel

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Dr. Chi Ho To School of Optometry, Hong Kong Polytechnic University Hung Hom, Kowloon Hong Kong, SAR (China) Tel. +852 2766 6102, Fax +852 2764 6051, E-Mail [email protected]

Aqueous Humor Formation

The level of IOP is maintained by the dynamic balance between the rate of secretion (inflow) and drainage (outflow) of aqueous humor. Aqueous humor is a transparent fluid that is formed by the ciliary processes of the ciliary body. After production, aqueous humor flows from the posterior chamber to the anterior chamber of the eye via the pupil. In the anterior chamber, the temperature difference between the warmer iris and the cooler cornea results in a convectional circulation of the aqueous fluid. Finally, aqueous humor exits the eye via one of the two principal outflow pathways, the pressure-dependent trabecular pathway and the pressure-independent uveoscleral pathway [6, 7]. The flow of aqueous humor serves a number of important functions in the eye [6, 8]: (1) to maintain adequate IOP for structural integrity and normal optical functioning of the eye; (2) to supply nutrients to as well as remove metabolic wastes from the avascular structures of the anterior segment; (3) to maintain a high concentration of ascorbate in most species, including humans (ascorbate may have many functions including an antioxidative role to combat oxidative challenges), and (4) to participate in cellular and humoral immune responses under adverse conditions, such as inflammation and infection. The secretion of aqueous humor demonstrates a circadian rhythm with the highest production rate in the morning right after awakening and lowest during sleeping [9]. On average, the rate of aqueous humor secretion ranges from 2.5 to 3.0
l
min–1. Since the total volume of the aqueous chamber is about 300
l, the entire volume of aqueous humor is recycled in every 100 min [7].

Regulation of Aqueous Humor Formation

The driving force of aqueous humor secretion is primarily derived from the energy-dependent ion transport (e.g. Cl– secretion) across the ciliary epithelium, thus creating an osmotic gradient to move water by osmosis. The transepithelial ion secretion proceeds in at least three major steps: ion uptake from the ciliary stroma into the pigmented ciliary epithelium (PE); ion diffusion from PE to non-pigmented ciliary epithelium (NPE), and finally ion efflux from NPE into the posterior chamber [10–16]. The rate of aqueous humor formation has been shown to be modulated by a number of well-known secondary messenger systems including cAMP and cyclic guanosine 3,5-monophosphate (cGMP) pathways [17–22]. Regulation of Aqueous Humor Formation by NO

Their effects are largely mediated by modulating the activities of ion transporters and channels in both PE and NPE in order to control the rate of ion secretion into the aqueous humor. Recently, attention has been given to the nitric oxide (NO) pathway. NO is a signaling molecule which has multiple physiological significances in relation to its protective, regulatory and deleterious functions [23]. In the biological system, NO is produced by the conversion of L-arginine into L-citrulline by NO synthase (NOS) [24]. There are three NOS isoforms [25]: 1 NOS I, previously called neuronal NOS (nNOS) – a low output NOS regulated by Ca2+ and calmodulin; 2 NOS II, previously called inducible NOS (iNOS) – a high output NOS and is Ca2+/calmodulin independent, and 3 NOS III, previously called endothelial NOS (eNOS) – also a low output NOS regulated by Ca2+ and calmodulin. NO is known to regulate several physiological functions. It is found to be the most powerful vasodilator. In addition, it is a free radical and capable of inflicting oxidative damage. NO plays important regulatory roles in various physiological functions, ranging from altering the cardiovascular system to modulating neuronal function [24, 26, 27]. The direct effect of NO is by binding to the ‘heme complexes’ in molecules such as guanylate cyclase (GC), cytochrome P450, NOS, and hemoglobin [28]. The most notable heme protein that forms an iron-nitric oxide adduct in vivo is the soluble GC (sGC) [29]. When NO binds to sGC, it changes the protein configuration and activates the sGC. The activated sGC can facilitate the conversion of guanosine triphosphate to cGMP [28]. The NO-sGC-cGMP signal transduction pathway is a major NO signaling pathway [30]. The cGMP produced by the activation of sGC is an important second messenger that acts on three main classes of effector proteins [31]: cGMP-dependent protein kinase, cGMP-gated ion channels, and cGMP-regulated phosphodiesterase. The biological effects of cGMP are terminated when it is broken down by the phosphodiesterase. NO has been suggested to modulate several physiological functions in the eye including retinal phototransduction [32], ocular immune responses [33], vascular tone [34] and aqueous humor dynamics [35]. Because of its potential pharmacological significance in glaucoma therapy, the effect of NO on IOP has been extensively studied. However, the result of these studies has not been conclusive because of the multiple actions of NO on both inflow and outflow pathways [33]. NO donors have been demonstrated to reduce IOP in normal and glaucomatous aniNeuroembryol Aging 2006–07;4:8–12

9

mals [36–38] and in humans [39], and the hypotensive effect is largely explained by the effects on vasodilation and/or relaxation of smooth muscle, such as the modulation of ocular blood flow [40], trabecular meshwork and ciliary muscle relaxation [41], and aqueous humor outflow [37]. However, conflicting data have been reported that NO may not contribute to ocular hypotension in humans [42]. Recent demonstration of NOS activities in pig [43] and bovine [44] ciliary processes, and the localization of NOS in pig ciliary epithelium [45] added support to the involvement of NO in regulating the aqueous humor formation. In an in vitro arterially perfused bovine eye study, a nitrovasodilator, sodium azide, was shown to lower IOP by acting on the ciliary epithelium directly but not by relaxing the vascular smooth muscles [46]. Consistent with this finding, it has been shown that L-arginine, a precursor of NO, produces a parallel reduction in aqueous humor formation and IOP in an arterially perfused porcine eye. The inhibition is blocked by NG-nitroL-arginine methyl ester (L-NAME), an NOS inhibitor [20]. Similarly, 8-pCPT-cGMP as well as sodium azide and sodium nitroprusside elicit a significant inhibition of aqueous humor formation and IOP. The inhibitory effects elicited by sodium azide and sodium nitroprusside can be blocked by 1H-[1,2,4]Oxadiazolo[4,3-alquinoxalin-1-one (ODQ), an sGC inhibitor [20]. These findings strongly suggest the inhibitory action of NO on aqueous humor formation and thereby IOP. In excised ciliary epithelium, NO donors such as sodium nitroprusside produce both the inhibition of short-circuit current (Isc) and net transepithelial Cl– secretion across the native porcine ciliary epithelium, supporting the functional significance of NO in mediating Cl– secretion into the posterior chamber [47]. Unlike the results of whole eye studies [20], 8-pCPT-cGMP produces a paradoxical stimulation of both Isc and net Cl– flux in porcine ciliary epithelium [47]. This finding is consistent with another independent study using the same tissue where the activation of the NO-sGC-cGMP pathway stimulates a transmembrane anionic current which is inhibited by Cl– channel blockers [48]. In parallel with this finding, activation of the sGC-cGMP-PKG pathway causes a transient stimulation of the Isc [49]. The exact reason for the discrepancy is not apparent because of the complexity of the system. It might suggest that NO has at least two opposing effects on Cl– secretion. It possibly stimulates the Cl– transport through cGMP-dependent pathways and inhibits Cl– secretion through cGMP-independent pathways simultaneously. Another possibility is that in the pig, aqueous humor for10

Neuroembryol Aging 2006–07;4:8–12

mation is not only driven by Cl– secretion, but also by other anionic players such as HCO3–. For example, depletion of HCO3– from the bathing solution abolished the Isc in porcine ciliary epithelium [47]. In addition, recent findings showed that the removal of HCO3– from the stromal bath significantly reduced the fluid transport into the aqueous humor by half [50]. In that case, NO might have exerted different effects on both electrogenic Cl– and HCO3– transport. The effect of NO on HCO3– transport is yet to be determined and the exact mechanism by which NO modulates the rate of aqueous humor formation and thereby IOP is unknown. Given the fact that NO alters the rate of Cl–, and possibly HCO3– secretion into the posterior chamber, it is reasonable to speculate that the activities of certain ion transporters and channels in both PE and NPE cells might be modulated by NO, hence affecting the inflow rate. As found in studies of bovine ciliary epithelium, the quantities and distribution of the Na+-K+-2Cl–, a major player responsible for secreting Cl– across the ciliary epithelium, might change with the age of the animal [10, 51], suggesting that the potential importance of NO in the regulation of aqueous humor formation might vary with age as well. Further studies are required to investigate the effect of NO on the activities of respective ion transporters and also the effect of aging on the distribution of ion transporters.

Conclusion

Although pharmacological treatment of glaucoma has been generally effective, the precise action of these agents on the regulation of aqueous humor formation is still largely unknown. It renders the modification of these agents so as to achieve better efficacy very difficult. The modulation of aqueous humor secretion by NO may represent a potential target for lowering IOP. Indeed, understanding of the pathophysiology of aqueous humor formation not only provides scientific information of how aqueous humor is secreted and regulated but also helps to devise a better treatment for glaucoma and save the vision of millions of people worldwide.

Acknowledgment The study was kindly supported by the PolyU Research Grants (A-PE59, A-PF22, G-T807, G-U028).

Do /Kong /Chan /Lam /To

References 1 Alward WL: Biomedicine. A new angle on ocular development. Science 2003;299:1527– 1528. 2 Quigley HA: Number of people with glaucoma worldwide. Br J Ophthalmol 1996; 80: 389–393. 3 Friedman DS, Wolfs RC, O’Colmain BJ, Klein BE, Taylor HR, West S, Leske MC, Mitchell P, Congdon N, Kempen J, Eye Diseases Prevalence Research Group: Prevalence of open-angle glaucoma among adults in the United States. Arch Ophthalmol 2004; 122:532–538. 4 Collaborative Normal-Tension Glaucoma Study Group: Comparison of glaucomatous progression between untreated patients with normal-tension glaucoma and patients with therapeutically reduced intraocular pressures. Am J Ophthalmol 1998;126:487–497. 5 The AGIS Investigators: The advanced glaucoma intervention study (AGIS). 7. The relationship between control of intraocular pressure and visual field deterioration. Am J Ophthalmol 2000;130:429–440. 6 Krupin T, Civan MM: Physiologic basis of aqueous humor formation; in Ritch R, Shields MB, Krupin T (eds): The Glaucomas: Basic Sciences. St. Louis, Mosby, 1995, vol 1, pp 251–280. 7 Freddo TF: Anatomy and pathophysiology of aqueous production and outflow; in Sassani JW (ed): Ophthalmic Fundamentals: Glaucoma. Thorofare, Slack, 1999, pp 33– 51. 8 Civan MM: Transport components of net secretion of the aqueous humor and their integrated regulation; in Civan MM (ed): Current Topics in Membranes; vol 45: The Eye’s Aqueous Humor: From Secretion to Glaucoma. San Diego, Academic Press, 1998, pp 1–24. 9 Brubaker RF: Flow of aqueous humor in humans [The Friedenwald Lecture]. Invest Ophthalmol Vis Sci 1991;32:3145–3166. 10 Crook RB, Takahashi K, Mead A, Dunn JJ, Sears ML: The role of NaKCl cotransport in blood-to-aqueous chloride fluxes across rabbit ciliary epithelium. Invest Ophthalmol Vis Sci 2000;41:2574–2583. 11 Wiederholt M, Helbig H, Korbmacher C: Ion transport across the ciliary epithelium: lessons from cultured cells and proposed role of the carbonic anhydrase; in Botré F, Gross G, Storey BT (eds): Carbonic Anhydrase. New York, VCH, 1991, pp 232–244. 12 Do CW, Peterson-Yantorno K, Civan MM: Swelling-activated Cl– channels support Cl– secretion by bovine ciliary epithelium. Invest Ophthalmol Vis Sci 2006; 47: 2576– 2582.

Regulation of Aqueous Humor Formation by NO

13 Do CW, Lu W, Mitchell CH, Civan MM: Inhibition of swelling-activated Cl– currents by functional anti-ClC-3 antibody in native bovine non-pigmented ciliary epithelial cells. Invest Ophthalmol Vis Sci 2005; 46: 948– 955. 14 Jacob TJ, Civan MM: Role of ion channels in aqueous humor formation. Am J Physiol 1996;271:C703–C720. 15 Do CW, To CH: Chloride secretion by bovine ciliary epithelium: a model of aqueous humor formation. Invest Ophthalmol Vis Sci 2000;41:1853–1860. 16 McLaughlin CW, Peart D, Purves RD, Carre DA, Macknight AD, Civan MM: Effects of HCO3– on cell composition of rabbit ciliary epithelium: a new model for aqueous humor secretion. Invest Ophthalmol Vis Sci 1998; 39:1631–1641. 17 Ni Y, Wu R, Xu W, Maecke H, Flammer J, Haefliger IO: Effect of cAMP on porcine ciliary transepithelial short-circuit current, sodium transport, and chloride transport. Invest Ophthalmol Vis Sci 2006; 47: 2065– 2074. 18 Do CW, Kong CW, To CH: cAMP inhibits transepithelial chloride secretion across bovine ciliary body/epithelium. Invest Ophthalmol Vis Sci 2004;45:3638–3643. 19 Do CW, Peterson-Yantorno K, Mitchell CH, Civan MM: cAMP-activated maxi-Cl– channels in native bovine pigmented ciliary epithelial cells. Am J Physiol Cell Physiol 2004; 287:C1003–C1011. 20 Shahidullah M, Yap M, To CH: Cyclic GMP, sodium nitroprusside and sodium azide reduce aqueous humour formation in the isolated arterially perfused pig eye. Br J Pharmacol 2005;145:84–92. 21 To CH, Kong CW, Chan CY, Shahidullah M, Do CW: The mechanism of aqueous humour formation. Clin Exp Optom 2002; 85: 335– 349. 22 Do CW, Civan MM: Basis of chloride transport in ciliary epithelium. J Membr Biol 2004;200:1–13. 23 Wink DA, Mitchell JB: Chemical biology of nitric oxide: insights into regulatory, cytotoxic, and cytoprotective mechanisms of nitric oxide. Free Radic Biol Med 1998;25:434– 456. 24 Moncada S, Palmer RM, Higgs EA: Nitric oxide: physiology, pathophysiology, and pharmacology. Pharmacol Rev 1991;43:109– 142. 25 Masters BS: Structural variations to accommodate functional themes of the isoforms of NO synthases; in Ignarro LJ (ed): Nitric Oxide: Biology and Pathobiology. San Diego, Academic Press, 2000, pp 91–104. 26 Ignarro LJ: Endothelium-derived nitric oxide: pharmacology and relationship to the actions of organic nitrate esters. Pharm Res 1989;6:651–659.

27 Culotta E, Koshland DE Jr: NO news is good news. Science 1992;258:1862–1865. 28 Miranda KM, Espey MG, Jourd’heuil D, Grisham MB, Fukuto JM, Feelisch M, Wink DA: The chemical biology of nitric oxide; in Ignarro LJ (ed): Nitric Oxide: Biology and Pathobiology. San Diego, Academic Press, 2000, pp 41–56. 29 Murad F: The nitric oxide-cyclic GMP signal transduction system for intracellular and intercellular communication. Recent Prog Horm Res 1994;49:239–248. 30 Ignarro LJ, Degnan JN, Baricos WH, Kadowitz PJ, Wolin MS: Activation of purified guanylate cyclase by nitric oxide requires heme. Comparison of heme-deficient, hemereconstituted and heme-containing forms of soluble enzyme from bovine lung. Biochim Biophys Acta 1982; 718:49–59. 31 Ignarro LJ: Nitric Oxide: Biology and Pathobiology. San Diego, Academic Press, 2000, pp 3–19. 32 Perez MT, Larsson B, Alm P, Andersson KE, Ehinger B: Localisation of neuronal nitric oxide synthase-immunoreactivity in rat and rabbit retinas. Exp Brain Res 1995;104: 207– 217. 33 Becquet F, Courtois Y, Goureau O: Nitric oxide in the eye: multifaceted roles and diverse outcomes. Surv Ophthalmol 1997;42:71–82. 34 Haefliger IO, Meyer P, Flammer J, Luscher TF: The vascular endothelium as a regulator of the ocular circulation: a new concept in ophthalmology? Surv Ophthalmol 1994; 39: 123–132. 35 Nathanson JA, McKee M: Identification of an extensive system of nitric oxide-producing cells in the ciliary muscle and outflow pathway of the human eye. Invest Ophthalmol Vis Sci 1995;36:1765–1773. 36 Nathanson JA: Nitrovasodilators as a new class of ocular hypotensive agents. J Pharmacol Exp Ther 1992;260:956–965. 37 Schuman JS, Erickson K, Nathanson JA: Nitrovasodilator effects on intraocular pressure and outflow facility in monkeys. Exp Eye Res 1994;58:99–105. 38 Wang RF, Podos SM: Effect of the topical application of nitroglycerin on intraocular pressure in normal and glaucomatous monkeys. Exp Eye Res 1995;60:337–339. 39 Chuman H, Chuman T, Nao-i N, Sawada A: The effect of L-arginine on intraocular pressure in the human eye. Curr Eye Res 2000;20: 511–516. 40 Schmetterer L, Polak K: Role of nitric oxide in the control of ocular blood flow. Prog Retin Eye Res 2001;20:823–847. 41 Wiederholt M, Sturm A, Lepple-Wienhues A: Relaxation of trabecular meshwork and ciliary muscle by release of nitric oxide. Invest Ophthalmol Vis Sci 1994; 35: 2515– 2520.

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42 Kiss B, Dallinger S, Findl O, Rainer G, Eichler HG, Schmetterer L: Acetazolamideinduced cerebral and ocular vasodilation in humans is independent of nitric oxide. Am J Physiol 1999;276:R1661–R1667. 43 Haufschild T, Nava E, Meyer P, Flammer J, Luscher TF, Haefliger IO: Spontaneous calcium-independent nitric oxide synthase activity in porcine ciliary processes. Biochem Biophys Res Commun 1996; 222:786–789. 44 Geyer O, Podos SM, Mittag T: Nitric oxide synthase activity in tissues of the bovine eye. Graefes Arch Clin Exp Ophthalmol 1997; 235:786–793.

12

45 Meyer P, Champion C, Schlotzer-Schrehardt U, Flammer J, Haefliger IO: Localization of nitric oxide synthase isoforms in porcine ocular tissues. Curr Eye Res 1999;18:375–380. 46 Millar JC, Shahidullah M, Wilson WS: Intraocular pressure and vascular effects of sodium azide in bovine perfused eye. J Ocul Pharmacol Ther 2001;17:225–234. 47 Kong CW: Chloride (Cl–) transport and its regulation by nitric oxide (NO) in porcine ciliary body/epithelium (CBE); PhD thesis, Hong Kong, Hong Kong Polytechnic University, 2005, p 190. 48 Fleischhauer JC, Beny JL, Flammer J, Haefliger IO: NO/cGMP pathway activation and membrane potential depolarization in pig ciliary epithelium. Invest Ophthalmol Vis Sci 2000;41:1759–1763.

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49 Wu R, Yao K, Flammer J, Haefliger IO: Role of anions in nitric oxide-induced short-circuit current increase in isolated porcine ciliary processes. Invest Ophthalmol Vis Sci 2004;45:3213–3222. 50 Candia OA, To CH, Law CS: Fluid transport across the isolated porcine ciliary epithelium. Invest Ophthalmol Vis Sci, in press. 51 Dunn JJ, Lytle C, Crook RB: Immunolocalization of the Na-K-Cl cotransporter in bovine ciliary epithelium. Invest Ophthalmol Vis Sci 2001;42:343–353.

Do /Kong /Chan /Lam /To

Neuroembryol Aging 2006–07;4:13–18 DOI: 10.1159/000103734

Published online: June 8, 2007

Revisiting the Trabecular Meshwork in the Eyes of the Developing Human C.Y. Chan a Winnie Wai-Ying Li b Calvin C.P. Pang b Dennis S.C. Lam b David T. Yew a Departments of a Anatomy and b Ophthalmology and Visual Sciences, Chinese University of Hong Kong, Hong Kong Eye Hospital, Hong Kong, SAR, China

Key Words Anterior angle
Collagen fibers
Elastic fibers
Fetal eye

Abstract Background: The trabecular meshwork (TM) is a specialized tissue which is crucial to maintain ocular health (e.g. intraocular pressure) and to sustain optical clarity. Since collagen and elastic fibers are the primary structural components of the tissue, the characterization of these fibers in the fetal eye would advance our understanding of the basic morphology and further congenital studies related to the TM. Aims: To reveal the early development of the human fetal TM. Results: The results showed the presence of circular (radial) and longitudinal collagen fibers as well as trabecular cells in the TM by 12 weeks of gestation. By 26 weeks, the venous system connected with Schlemm’s canal and thin elastic fibers began to form. At birth, longitudinal collagen fibers linked with each other to form long strands. There was also evidence of an increase in circular (or radial) collagen fibers in the TM. Conclusion: The present study characterized the developmental and morphological changes in the genesis of the fetal TM in terms of growth in collagen and elastic fibers. The elastic fibers were found to exist in small quantity at birth, and, thus, any increase in these fibers to maturity is likely to occur after birth. Copyright © 2007 S. Karger AG, Basel

© 2007 S. Karger AG, Basel 1661–3406/07/0042–0013$23.50/0 Fax +41 61 306 12 34 E-Mail [email protected] www.karger.com

Accessible online at: www.karger.com/nba

Introduction

The trabecular meshwork (TM) is a specialized tissue located at the anterior angle of the eye. It is composed of connective tissues (mainly collagen and elastic fibers) covered with trabecular cells. Together with Schlemm’s canal (SC) and aqueous veins, it forms the major outflow/ drainage pathway for aqueous humor [1]. Since a normal intraocular pressure is crucial to maintain ocular health and to sustain optical clarity, blockage of the TM may cause severe pathological changes such as glaucoma. Interest in the development of the TM has been immense since the study of Reme and d’Epinay [2] in 1981. They documented the trabecular anlage formation already at 15 weeks of gestation (WG) [2]. By the 24th week, differentiation of the definitive structure, e.g. trabecular beams, was clearly observed and specialization was continued until 28 weeks [2]. At a similar gestational stage, SC was found to have first delineation [3], and the maturing meshwork is lined by uveal trabecular endothelial cells [4]. Such lining was perforated with discrete intercellular gaps. The gaps continue to increase in size and density until 18–30 WG. They also provided a communication pathway between the fetal anterior chamber and the widening intertrabecular TM space [5]. The outer collectors that parallel the SC, however, were not evident until 33 weeks [3]. On the other hand, interconnection between SC and outer collectors was visible at 36 WG via intercanal links, with evidence of the intrascleral plexus

Prof. David T. Yew Department of Anatomy, The Chinese University of Hong Kong Shatin, New Territories Hong Kong, SAR (China) Tel. +852 2609 6899, Fax +852 2603 5031, E-Mail [email protected]

at the growing stage [3]. Further molding and maturation of the anterior angle were attained at 1–8 years of age [2]. During the last trimester of gestation, a posterior migration of the ciliary body was seen. It was due to the differential growth of the corneoscleral layer, the ciliary ectoderm and the mesoderm in between [6]. Throughout the fetal development, no remarkable cleavage, atrophy [2] or cell death could be identified in the human TM [7]. In order to fulfill its physiological function, the TM thickens by inscribing more intertrabecular space during development. This space is part of the aqueous drainage pathway which belongs to the uveal outflow system. The progressive changes were observed and accompanied with a decrease in cellularity with no changes in trabeculae (both in terms of thickness and number) until 20 fetal weeks [8]. However, another study by McMenamin [7] revealed a 2- to 3-fold increase in the absolute number of cells per eye with the employment of stereological analysis. Considering the absolute volume, the extracellular matrix increased most significantly from 12–22 WG, while the increase in intertrabecular spaces was more variable. Another study on cat eyes also showed a 150fold increase in the volume of TM from birth to adulthood [9]. Towards maturation, the intertrabecular spaces also increased 24-fold, and there were 14-fold-elevated trabecular cell numbers, too [9]. Besides, elastic fibers also play an important role in the TM as it is the major component of the trabecular beams, which were responsible for elastic properties of the tissue [10]. Elastin was found to be present, which was thought to play a crucial role in the resistance of aqueous humor outflow [11]. In congenital glaucoma, trabecular beams are thicker than in normal subjects. Such a thickening is thought to hinder the normal migration of ciliary body and iris root, which results in the obstruction of aqueous humor drainage and leads to an increase in intraocular pressure [6]. Other researchers also found thickened subcanalicular tissues (composed of elastin or fibronectin fibrils) beneath the inner wall of SC in glaucomatous patients [11, 12]. Such a thickened membrane was proposed to be an immature stage of the TM, which was believed to lead to glaucoma development [13]. Besides, patients with congenital glaucoma also demonstrate primary anomalies, including a developmental defect in TM, persistent mesenchymal tissues in the anterior angle and excessive collagen fibrils in the trabecular matrix [14]. In this study, we evaluated and revisited the development of the TM in order to analyze (1) the incorporation of elastic and collagen fibers; (2) the appearance of TM cells, and (3) the perforation of endothelial cells together 14

Neuroembryol Aging 2006–07;4:13–18

with SC formation. This information would increase our understanding of the development of the human TM during gestation.

Materials and Methods Human fetuses were obtained after death with the consent of both the parents and the Kunming Hospital, China. The gestational age and numbers (in parentheses) of enucleated eyes adopted for the experiment were as follows: 12 weeks (1), 15 weeks (2), 20 weeks (1), 23 weeks (1), 26 weeks (1), 28 weeks (1), 30 weeks (1), 34 weeks (1), and a stillborn at 40 weeks (1). All of the eyes were fixed in 10% buffered formalin and dehydrated in graded alcohol series. They were cleared by immersion in xylene and finally embedded in paraffin. Sections (6
m) were obtained and stained with Weigert’s resorcin-fuchsin, a progressive staining technique for various fibers [15]. The elastic fibers would appear bluish-black, while the collagen fibers would be in pink after the staining. Finally, samples were observed and documented with a Nikon photomicroscope.

Results

As shown in figure 1, TM was already present at 12 WG, mainly at the region where the ciliary muscle and the scleral spur met (fig. 1). At the anterior pole, where one marked the corneal endothelium, a small condensation of collagen fibers was noted, which would likely become Schwalbe’s line at a later stage (fig. 2). On the other hand, the peripheral corneal endothelium appeared to detach from each other with gaps present intercellularly. At this stage, the major components of TM were collagen fibers, which existed mainly in two forms. The lightly stained collagen fibers were radial or circular, whilst the large fibers were short and longitudinally arranged (fig. 3). Gaps were found between the collagen fibers in the angular region as well as between the base of ciliary processes and the corneoscleral junction (fig. 3). As seen in figure 4, the whole TM was generally compact with little space. A flattened SC lined by endothelium could be marked, along with sclerotic vessels externally (fig. 4). Elongated and spindle-like trabecular cells, being similar to fibrocytes morphologically, were seen in the early gestational period (fig. 4). By 26 WG, the collagen fibers increased in Schwalbe’s lines (fig. 5). Both types of collagen fibers became prominent and formed stacks at the anterior and posterior TM with blood cells scattered in between (fig. 5, 6). These fibers originated mainly from the scleral spur region. Besides, thin elastic fibers penetrated the posterior region of Chan /Li /Pang /Lam /Yew

Fig. 1. TM at 12 WG (arrow) near to scleral spur (S) and ciliary muscles (M). !200.

Fig. 2. Condensation of collagen fibers (arrow) adjacent to the posterior epithelium of cornea forming Schwalbe’s region at 12 WG. !400.

Fig. 3. In the TM, deposits of longitudinal collagen fibers (solid

Fig. 4. Presence of SC (solid arrow) and trabecular cells (dotted

arrows) and radial collagen fibers (dotted arrows) began to appear at 12 WG. !400.

arrow) and scleral vein (S) at 12 WG. !200.

Trabecular Meshwork in the Human Fetal Eye

Neuroembryol Aging 2006–07;4:13–18

15

Fig. 5. Thickening of collagen fibers (arrow) at Schwalbe’s region

Fig. 6. Presence of short longitudinal collagen fibers (solid arrow)

at 26 WG. !400.

and radial collagen fibers (dotted arrows) at 26 WG. !400.

Fig. 7. Penetration of elastic fibers (arrows) in the posterior part of the TM by 26 WG. !400.

Fig. 8. Longitudinal short elastic fibers (arrows) now appear in the anterior areas of the meshwork at 26 WG. !400.

the TM (fig. 7) and thin longitudinal elastic fibers began to appear randomly towards the anterior TM region (fig. 8). They were mostly short elastic fibers which followed the beams of the collagen fibers. Moreover, a mem-

branous layer was found to cover the anterior angle, and gaps were present intercellularly (fig. 9). A definitive channel was formed and connected to both the aqueous veins and the SC (fig. 10).

16

Neuroembryol Aging 2006–07;4:13–18

Chan /Li /Pang /Lam /Yew

Fig. 9. Gaps (arrow) were evident in the surface of the TM guard-

Fig. 10. At 26 WG, the SC can be seen linking with the scleral vein

ing the angle at 26 WG. !200.

(arrow). !200.

Fig. 11. At 34 WG, some longitudinal collagen fibers were still

Fig. 12. At birth, the longitudinal collagen fibers linked together

short and not continuous (arrow). !400.

to form strands (solid arrow) where one could also see an increase in radial (circular) collagen fibers (dotted arrow). !400.

Trabecular Meshwork in the Human Fetal Eye

Neuroembryol Aging 2006–07;4:13–18

17

By the 34th week, short longitudinal collagen fibers were still active in forming layers (fig. 11). At 40 WG, long strands of collagen fibers were formed by stacking their short, longitudinal members together. Such a modification began at the region adjacent to the anterior angle (fig. 12) and the quantity of circular collagen fibers also increased (fig. 12). At this stage, the whole TM expanded further with more spacious pores inside the tissue layer.

Discussion

The present study agreed with the findings of Reme and d’Epinay [2] that the initial development of the TM is at 12–15 WG. Intercellular gaps were present in the posterior corneal endothelium at 12 WG, similar to a report by McMenamin [4]. The repositioning of the ciliary processes due to differential growth of the neuroepithelium and the corneoscleral mesoderm is noteworthy in all vertebrates including humans, calves and chicks [16]. The present study further showed that SC and the nearby sclerotic vessels were present at an earlier period (i.e. 12 WG) compared to that reported (i.e. 24 WG) by Ramirez et al. [3]. At a later period of gestation, SC even connected with the aqueous vein, which clearly revealed the initial basic framework as an aqueous outflow system. Collagen fibers play a very crucial role in the development of TM. It is the most dominant type of connective tissue in the TM during gestation, forming actively since 12 WG. These connective tissues formed the basic frame-

work and anchoring system of the TM [17, 18]. Elastic fibers, however, were less prominent during this period. They present as a thin connective structure until birth. Nevertheless, they are the prime connective components which offer resistance to aqueous flow and give the TM its elasticity at a later stage [10]. Since 26 weeks of the developmental stage, increase in cellular components (mainly collagen fibers) became very obvious. The increase in both the cellular content and intertrabecular space resulted in TM thickening, in agreement with McMenamin’s [7] study, but in contrast to another study, which suggested the thickening of TM was purely due to widening of the intertrabecular space [8]. In conclusion, the development of the human TM can be summarized in different stages: (1) the early presence of trabecular cells and collagen components; (2) the presence of SC along with sclerotic vessels; (3) an increase in collagen fibers and initial formation of radial or circular elastic fibers; (4) SC connected to the aqueous vein; (5) long strands of collagen fibers formed by stacking short fibers, and finally (6) the invasion of circular collagen fibers into the external layer and expansion of the intertrabecular space, resulting in increased TM thickness. In the present study, we presented a detailed description of the development of the human TM, emphasizing the changes in collagen and elastic fibers. This information advances our understanding of the basic morphology of the human TM and will benefit future pathological and congenital studies related to TM.

References 1

2

3

4

5

6

18

Hart W: Intraocular pressure; in Hart W (ed): Alder’s Physiology of the Eye. St. Louis, Mosby Year Book, 1992, pp 248–267. Reme C, d’Epinay SL: Periods of development of the normal human chamber angle. Doc Ophthalmol 1981; 51:241–268. Ramirez JM, Ramirez AI, Salazar JJ, Rojas B, De Hoz R, Trivino A: Schlemm’s canal and the collector channels at different developmental stages in the human eye. Cells Tissues Organs 2004;178:180–185. McMenamin PG: Human fetal iridocorneal angle: a light and scanning electron microscopic study. Br J Ophthalmol 1989;73: 871– 879. McMenamin PG: A morphological study of the inner surface of the anterior chamber angle in pre- and postnatal human eyes. Curr Eye Res 1989; 8:727–739. Anderson DR: The development of the trabecular meshwork and its abnormality in primary infantile glaucoma. Trans Am Ophthalmol Soc 1981;79: 458–485.

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McMenamin PG: A quantitative study of the prenatal development of the aqueous outflow system in the human eye. Exp Eye Res 1991; 53: 507–517. 8 Sugiura T, Mizokami K, Yamamoto M: The development of human trabecular meshwork. Nippon Ganka Gakkai Zasshi 1991; 95:1238– 1243. 9 Richardson TM, Marks MS, Ausprunk DH, Miller M: A morphologic and morphometric analysis of the aqueous outflow system of the developing cat eye. Exp Eye Res 1985; 41:31– 51. 10 Hamanaka T: Scleral spur and ciliary muscle in man and monkey. Jpn J Ophthalmol 1989; 33:221–236. 11 Segawa K: Trabecular meshwork and elastin. Nippon Ganka Gakkai Zasshi 1995;99:1291– 1302. 12 Umihira J: Ultrastructural immunohistochemical localization of elastin in the human trabecular meshwork. Nippon Ganka Gakkai Zasshi 1993;97:1143–1150.

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13 Tawara A, Inomata H: Developmental immaturity of the trabecular meshwork in congenital glaucoma. Am J Ophthalmol 1981; 92: 508– 525. 14 Lin J, Sun W, Li E, Wang S, Bai J, Yang H: Ultrastructural observations of the anterior chamber angle tissues in congenital glaucoma. Yan Ke Xue Bao 1994;10: 50–56. 15 Luna L: Manual of Histology Staining Methods of the Armed Forces Institute of Pathology, ed 3. New York, McGraw-Hill, 1968. 16 Barishak YR: The development of the angle of the anterior chamber in vertebrate eyes. Doc Ophthalmol 1978; 45:329–360. 17 Horstmann HJ, Rohen JW, Sames K: Age-related changes in the composition of proteins in the trabecular meshwork of the human eye. Mech Ageing Dev 1983;21:121–136. 18 Rittig M, Lutjen-Drecoll E, Rauterberg J, Jander R, Mollenhauer J: Type VI collagen in the human iris and ciliary body. Cell Tissue Res 1990;259:305–312.

Chan /Li /Pang /Lam /Yew

Neuroembryol Aging 2006–07;4:19–30 DOI: 10.1159/000103582

Published online: June 5, 2007

Morphological and Neurochemical Development of the Human Neural Retina T.C. Nag S. Wadhwa Department of Anatomy, All India Institute of Medical Sciences, New Delhi, India

Key Words Development
Differentiation
Neurochemical maturation
Retina

Abstract The human retina, a derivative of the diencephalon, differentiates into its components in sequential patterns following a spatiotemporal order. The neuronal differentiation in the fovea and central retina begins earlier than that occurring elsewhere, completes by midgestation (19–21 weeks) and continues in the periphery until 30 weeks of gestation. By birth, all retinal layers and neurons are mature over its entire extent, except at the fovea, where photoreceptor differentiation (outer segments) continues up to 5 months postnatally. During differentiation, the neurons express an array of marker molecules, such as amino acid neurotransmitters, neuropeptides, calcium-binding proteins, nitric oxide and proteins of phototransduction pathways. Although their precise involvement in retinal development is unclear, their patterns of expression indicate that many of them may be necessary for neuronal differentiation. In order to gain knowledge on this issue, future studies utilizing embryonic human retinal culture and transplantation of the tissue into suitable hosts and characterization of the role of neurochemicals involved in retinal development appears to be worthwhile. This approach may highlight novel information about the specific neurochemical requirements for the adult retina in normal and pathological states. Copyright © 2007 S. Karger AG, Basel

© 2007 S. Karger AG, Basel 1661–3406/07/0042–0019$23.50/0 Fax +41 61 306 12 34 E-Mail [email protected] www.karger.com

Accessible online at: www.karger.com/nba

Introduction

The retina develops as an outgrowth of the diencephalon during early embryonic life. Its neurons originate from two progenitor cell layers, the outer and inner neuroblastic zones, of the embryonic neural retina from 6 weeks of gestation (WG). Cell proliferation is initially rapid in the inner neuroblastic zone, and then the waves spread to the outer neuroblastic zone. The human retina is characterized by the possession of a specialized anatomical region (about 500
m in diameter), called the fovea, where the density of the cones and neurons of the inner retinal layers is exceedingly higher than elsewhere in the retina. Studies have indicated that the onset of differentiation of most neurons starts in the incipient fovea and than progresses outside the fovea. In vitro studies with embryonic animal retinas indicate that the generation of various retinal neurons is under the control of various neuroactive substances. Subsequently, there have been initiatives to study the development of various markers of neurons and their maturation. Characterization of properties of the markers reveals that many of them are apparently involved in cellular proliferation and differentiation at various stages of development to shape the future retina. Therefore, knowledge of the role of various molecules in retinal neurogenesis would be of help to understand the mechanism of normal retinal development. The present review aims to collate the current state of knowledge of the various molecular markers and their spatiotemporal expressions during development and differentiation of the human neural retina.

Prof. Shashi Wadhwa Department of Anatomy All India Institute of Medical Sciences New Delhi 110029 (India) Tel. +91 11 2659 4875, Fax +91 11 2658 8663, E-Mail [email protected]

Components of the Adult Human Retina

The adult human neural retina consists of three main types of cells, viz photoreceptors, interneurons and ganglion cells (GCs), arranged in three major layers, outer nuclear layer (ONL), inner nuclear layer (INL) and ganglion cell layer (GCL), respectively (fig. 1). Photoreceptors synapse in the outer plexiform layer (OPL), while the interneurons synapse in OPL and inner plexiform layer (IPL). The principal outflow GC axons form the nerve fiber layer to exit the optic nerve at the optic papilla. In addition to these cell types, specialized radial glia, known as Müller cells, are interspersed amongst the cells, spanning the entire depth of the retina from the inner to the outer limiting membrane. These limiting membranes provide structural support to the retina.

Formation of Retinal Layers and Neurons

O’Rahilly [1] reviewed the early development of the human eye during the embryonic period and showed the appearance of the retinal disk at 28 days of gestation. At 6 WG, the retina begins to differentiate through the formation of its inner neuroblastic zone and GCs. Also, at this period, the axons of GCs enter the optic nerve [2]. Müller cells appear at 7–8 WG and with GCs they coexist in the inner neuroblastic zone [1–3]. Rhodes [3] described the formation of the layer of Chievitz at 8 WG due to inward migration of GCs and Müller cells. At 9 WG, the outer neuroblastic zone appears and is separated from the inner neuroblastic zone by the layer of Chievitz. The latter becomes the definitive IPL, as seen in the retina at 11 WG (fig. 2a). Meanwhile, the GCL has been formed from the inner neuroblastic zone, while cell proliferation continues in the outer neuroblastic zone. Firm data on the generation of the various neurons in the human retina are not yet available. Tritiated thymidine labeling in monkeys shows that horizontal cells are generated after GCs and just before cones, and that bipolar cells are born late with rods and Müller cells [4]. In the human retina, probably the horizontal cells are born in the incipient fovea at 7–8 WG. Before 10 WG, the outer neuroblastic zone consists of a homogeneous population of undifferentiated neuroblasts. Ki-67 immunolabeling indicates that the central retina appears more mature than the peripheral retina before 9 WG [5]. At around 11 WG, the INL begins to differentiate by the formation of amacrine cells from the innermost row of the outer neuroblastic zone [3]. Bipolar cells arise from this layer at a late gestational period 20

Neuroembryol Aging 2006–07;4:19–30

Fig. 1. Photomicrograph of the adult human retina at the macula. The different retinal layers are indicated. PL = Photoreceptor layer; RPE = retinal pigment epithelium. Scale bar: 50
m.

(15–16 weeks). The photoreceptors differentiate from the outer rows of this zone, cones appear first at 10 WG and rods appear by 12 WG [3, 6, 7]. The OPL begins to differentiate in the central retina by 11–12 WG. By 16 WG, this layer becomes prominent in the central (fig. 2b) to midperipheral part of fetal retina. Except for the narrow peripheral zone, the differentiation of most retinal neurons is attained by midgestation (20–21 WG; fig. 2c), which proceeds in a central to peripheral direction. Also, connections to visual centers via retinal GC axons are partially formed at this time point [8, 9]. Using antigenic markers, immunohistochemical studies addressed the issue of specific cell types present in early neuroblastic layers. Walcott and Provis [10] used markers to identify progenitor cells and Müller cells in the human fetal retina. They found nestin (an intermediate filament protein expressed in neural progenitor cells) expression to colocalize with vimentin (a marker of Müller cells) in cells over extensive regions of human fetal retina. Nestin and vimentin each colocalized with Ki-67positive proliferating cells. These interesting findings led Nag/Wadhwa

the authors to suggest that Müller cells are a differentiated phenotype of retinal progenitor cells. Immunoreactivity (IR) for other markers such as GABA [11], tyrosine hydroxylase [12, 13], calretinin [14], Trk B and Trk C [15] is localized in the inner neuroblastic cells (GCs) from 10 to 11 WG onward (table 1). The progenitor cells destined to form neurons express IR to Bcl-2 and Bcl-X from 13 WG [16]. A study showed differential expressions of Drosophila optomotor blind-related T-box genes (TBX2, TBX3 and TBX5) in the neuroblastic zones of embryonic human retina [17] and suggested these genes to play some role in providing positional information and cell differentiation. The immature neural cells show high levels of N-myc and c-src mRNAs at 12 WG, though N-myc disappears with their differentiation [18]. Their specific role in the early human retina is unknown.

Growth of the Retinal Area

Using postmortem specimens, Robb [19] showed a rapid increase in the retinal area in the last trimester with a gradual increase until the 6th postnatal year. Retinal growth, determined by measuring the surface area, shows a steady growth rate of 10–15 mm2/week throughout 10– 40 WG [20]. Our data show a 12-fold increase in retinal area during 7–24 WG, with maximum increase (188%) occurring between 10 and 16 WG. This early increase in the retinal area is due to active generation of cells by mitosis seen up to 16–17 WG in the central retina and 24 WG in the peripheral retina [21]. Although immunomarkers suggest that progenitor cells remain in the postnatal retina, the increase in retinal area beyond this period occurs mainly by growth of individual cells.

The GCL and Establishment of a Cell Density Gradient across the Retina

GCs differentiate at 8–9 WG and arrange into a distinct layer (GCL) in the central retina at 13–14 WG. The early GCs possess few dendrites between 7 and 11 WG. At 10–11 WG, some of them exhibit dendritic filiform processes and spines. They show progressive growth and

Fig. 2. Light micrographs of the human fetal retina at three gestational ages. At 11 WG (a), the retina consists of inner (INZ) and

outer neuroblastic zones (ONZ), separated by the IPL. RPE = Retinal pigment epithelium. At 16 WG (b), the GCL and the OPL are

Development of the Human Neural Retina

prominent in the central retina. The INL neurons are still in the process of differentiation. At 20 WG (c), the retinal layers are well established. Scale bar: 50
m.

Neuroembryol Aging 2006–07;4:19–30

21

Fig. 3. Camera lucida drawing of DiI-impregnated GCs at 19–20 WG. Using morphological characteristics, three classes of GCs can be distinguished: small parvocellular cells (a), large magnocellular cells (b), and cells with mixed characteristics (c), physiologically classified as W cells. Scale bar: 100
m.

Table 1. Neurochemical detection of GCL neurons in the human fetal retina

Markers

Gestational age, WG

Retinal location

References

Tyrosine hydroxylase Calretinin TULP Trk A, B, Cb GABAa Leucine-enkephalinb Bcl-2 Parvalbuminc Recoverin Glutamateb Somatostatinb Calbindina Taurineb Nitric oxide synthasea Neuropeptide Y Substance Pb C-fos

10–21 11–25 11 11–21 11–25 12–30 13–22 13–25 13–24 14–21 15–40 15–25 16–17 16–21 17–40 20–30 26–40

midperipheral central, peripheral central, midperipheral central to peripheral central to peripheral midperipheral, foveal entire retina central, midperipheral central, midperipheral midperipheral central, midperipheral central, midperipheral central to peripheral central to peripheral midperipheral central, foveal central, peripheral

12, 13 14 28 15 11 12, 29 16 30, 31 32 33 34, 35 30, 31 36 37 12, 35 12, 29 38

a

Present in some cells. b Present in many cells. c Present in most cells. Gestational age range denotes the period of IR expression studied, by one or several workers.

22

Neuroembryol Aging 2006–07;4:19–30

Nag/Wadhwa

dendritic elaborations at 17–18 WG. By 20 WG, three major cell types (fig. 3) – parvocellular, magnocellular and cells with mixed features resembling physiological X, Y and W classes of GCs (or ,  and  classes of GCs in the cat) – are identifiable in the human retina [22]. Provis et al. [23–25] described the development of the human retinal GC topography. Mitosis ceases in the central retina at 14 WG to form the putative fovea, wherefrom the GCs later migrate to the peripheral region, which shows the highest cell density. At 13–14 WG, cell distribution in the GCL is almost uniform, but at term, a cell density gradient of about 10:1 in the central to peripheral retina is established. This is mainly due to a high rate of cell death in the peripheral GCL occurring between 18 and 30 WG [24–26]. The estimated number of GCs between 18 and 30 WG is 2.2–2.5 million, which then declines to 1.5 million cells due to apoptosis [23]. GCs mature in the central retina by 24 WG. On extrapolation from the data of La Vail et al. [4] on monkey retina, however, it would appear that the last GC should be born in the peripheral human retina by 20 or 22 WG at the latest. Thus a central to peripheral sequence of the GC maturation is evident [25]. Their maturation parallels the establishment of the retinal circulation. In adults, the GC density peaks around the fovea and declines sharply towards the periphery [27]. The GCL neurons express a number of characteristic markers right from the period they are generated. Table 1 shows the various markers labeled thus far and their spatiotemporal course of expression in the human fetal retina.

Photoreceptor Differentiation

As seen by scanning electron microscopy, at 15 WG, the cilium develops as a small protrusion from the apical ends of cone inner segments. The latter become arranged into mosaic patterns by 18–19 WG [39], wherein few large cone inner segments (putative blue cones) stand out prominently from the mosaic of many small cone inner segments (prospective red/green cones). Between 19 and 25 WG, they elongate and change from a spherical to an oval shape. The rod inner segments are identified at 18– 19 WG and show ciliary outgrowths. At 24 WG, the rod outer segments begin to develop from the distal ends of cilia [39]. Cell death in the ONL peaks at around 23–24 WG [40]. Fovea cones differentiate in the putative fovea at 13 WG [41]. Rods are absent until 15 WG [42], but present Development of the Human Neural Retina

in the region surrounding the foveal cone mosaic, where their density at 20–21 WG is 59,200/mm2. By 24 WG, foveal cones show a packing density of approximately 38,000/mm2 [43]. Their density gradually decreases towards the peripheral retinal region. Analysis of photoreceptor topography from 13 to 24 WG indicated that from the period of differentiation, photoreceptors migrate centripetally, i.e. towards the center of the developing fovea [41, 43]. Using cone opsin labeling, Xiao and Hendrickson [44] showed that foveal red/green cones have short outer segments at 15 WG and parafoveal blue cones (S cones) have outer segments at 12 WG. In the parafovea, both cone types (red/green and blue cones) have long outer segments by 20 WG. The photoreceptors in the midperipheral retina develop earlier than those in the fovea, which suggests that the neonates utilize more the peripheral retina before foveal cone vision becomes dominant [45]. A shallow fovea is present at birth [46]. Before birth, the fovea has a diameter of 1,000
m, whose size decreases to 650–700
m after birth. This is attributed to the migration of cones to the central region. The adult diameter of the fovea is noted at a postnatal age of about 45 months. From the time of proliferation, cones express many marker molecules in a spatiotemporal pattern in the human fetal retina. These molecules may be critical in their differentiation and maturation. These are tubby-like protein-1 (TULP) [28, 47], pigment epithelium-derived factor (secreted from the retinal pigment epithelium [48]), the Leber congenital amaurosis protein AIPL 1 [49] and cone opsins [44, 47, 50, 51]. TULP labeling in the cones precedes detection of any IR for cone opsins, suggesting that TULP may serve an important role for cone development [28]. This is consistent with a photoreceptor defect seen in patients with retinitis pigmentosa caused by mutations in the TULP gene. A good marker of cones in the adult retina is calbindin, a member of EF-family calciumbinding proteins (calbindin, parvalbumin and calretinin [14, 30]). Foveal cones show calbindin IR at 24 WG [31] and the central cones show calretinin IR at 20–21 WG [14]. In adults, all cones (except for foveolar cones) are brilliantly calbindin positive, but show weak calretinin IR. This suggests that calretinin plays a limited role for Ca2+ buffering in the cones of the human retina. [14]. Infant cones show IR to parvalbumin, although it does not express in the fetal retina till 25 WG [30]. In adults, only few cones of the macula show parvalbumin IR [14, 30]. In the adult human retina, cones are characterized by their possession of short, medium or long wavelengthspecific opsins. During development, cone opsins appear Neuroembryol Aging 2006–07;4:19–30

23

Table 2. Photoreceptor markers in the

developing human retina

Markers

Gestational age, WG

Retinal location

References

Cones IRBP TULP Phosphodiesterase  AIPL 1 Short opsin Neuron-specific enolase Synaptophysin Long/medium opsin C-fos Trk A, B, C Taurine Calretinin Heat shock protein 70 PEDF Calbindin

9–19 9–19 10 11.8 11–38 14–16 14–25 14–38 15–40 16–17, 21–21 18–25 20–25 20–33 21.5 24

foveal, peripheral foveal, peripheral foveal foveal foveal RT foveal to peripheral, RT foveal central central to peripheral central to peripheral central, midperipheral midperipheral foveal foveal

47 47 47 48 44, 47, 50, 51 56 56a; 40, 57 44, 47, 50, 51 38 15 36 14 58 48 31

Rods Crx Nrl S-antigen Rod opsin -Transducin

10 10–19 13–18 15, 20 20

foveal edge foveal edge, peripheral central, RT RT, foveal RT

47, 54 47 56a, 59 47, 56a 56a

IRBP = Interphotoreceptor retinal binding protein; PEDF = pigment epithelium-derived factor; RT = Retinal transplant. Gestational age range denotes the period of IR expression studied by one or several workers. a Observation from a transplant study.

early in the human fetal retina. Short opsin expresses earlier (11 WG) than the long/medium opsins (14 WG) across the fetal retina, and cones containing both short and long/medium opsins appear around the fovea shortly after long/medium opsin is expressed [44, 51]. The short and long/medium cones decrease in number after birth [51]. In addition, cones destined to express long or medium opsin may initially and transiently express short opsin [51]. Short cones show a spatial density of around 2,000/mm2 near the optic disk at 17 WG, which drops to 50% by midgestation and stabilizes at around 500 cells/ mm2 by birth [50, 51]. These density changes are associated with their displacement towards the incipient fovea [51]. The transcription factors Crx and Nrl are reported to express in early rod photoreceptors [52, 53]. In human fetal retina, Crx expression is detected by RT-PCR at 10– 10.5 WG [47, 54], and Nrl expression in rod nuclei at the foveal edge at 10–11 WG [47]. Both express in parallel with rod generation, indicating a critical functional in24

Neuroembryol Aging 2006–07;4:19–30

teraction operative between the two factors for rod development [55]. Rod opsin is first detected between 15 and 16 WG in the rod inner segments [47]. Since photoreceptor outer segments appear later, the significance of this early expression of opsins in the fetal human retina remains unknown. The various markers expressed by the developing photoreceptors are shown in table 2.

Retinal Vascular Development: Association with Neuronal Maturation

The development of retinal vasculature begins at around 15 WG. Early vessels, reported to be formed by spindle-shaped cells (presumably vascular precursor cells), appear first in the nerve fiber layer/GCL and then proceed towards the INL, where vessel formation occurs by sprouting from the preexisting vessels [60]. The contribution and presence of vascular precursor cells in vesNag/Wadhwa

sel formation in the fetal human retina have been debated (reviewed by Provis [61]). Hughes et al. [62] reported a prior invasion of vascular precursor cells in human retinal vascular development. According to Chan-Ling et al. [63], before 15 WG, vascular precursor cells, identified by adenosine diphosphatase enzyme histochemistry, migrate from the optic disk to the retina. They then proliferate and differentiate into endothelial cells characterized by their IR to CD34 and major histocompatibility complex class II antigens. These cells aggregate to form primordial vessels in the central, inner retinal layers. It is, however, clear that by 23–24 WG, the neural retina is almost vascularized, except for a narrow zone of the incipient fovea [64]. The outer plexus and the peripapillary capillaries are formed by extension of buds from the existing inner vessels, which occurs around the fovea between 25 and 26 WG. Evidence available thus far indicates that in the human retina, vasculogenesis is responsible for the formation of early vessels and the inner vascular plexus, whereas angiogenesis contributes to increasing vascular density and vascularization of the peripheral outer retina up to the ONL [60–65]. Asymmetry in vascular development is also reported, with temporal and superior quadrants showing advanced vascular development compared to the nasal-inferior quadrants. This differential vascular development pattern is perhaps linked to the more advanced neuronal maturation in the superior than in the inferior quadrant of the human retina [66]. Astrocytes seem to play a crucial role in vascular development in the fetal retina [61–64, 66–69]. The precursors of astrocytes, which are Pax 2 positive, migrate from the optic nerve head into the retina at around 8 WG [69]. They proliferate, accompany the developing vessels [64] and lie ahead of most peripheral vasculature [61]. In response to physiological hypoxia [70], they presumably release vascular endothelial growth factor, which promotes proliferation and differentiation of endothelial cells [61]. This seems likely, since its signal is reported to be absent before 15 WG, when vessel formation begins [66]. Studies have also shown microglia and macrophages [67, 71, 72] to associate with the developing human retinal vasculature. According to Diaz-Araya et al. [71, 72], microglia enter the fetal retina from the optic nerve head and retinal margin, and they as well as the macrophages intimately associate with the developing vasculature. Even before vascularization, at 10 WG, microglia are aligned in the IPL and OPL of the far peripheral retina [72]. Their actual contribution in retinal vascular development remains elusive. The immunophenotypical markers expressed by glia in human fetal retina are listed

Table 3. Glial markers in the human fetal retina

Development of the Human Neural Retina

Neuroembryol Aging 2006–07;4:19–30

Markers

Gestational age WG

References

Müller cells Bcl-2 Glutamine synthetase Vimentin Nestin CRALBP Nitric oxide synthase CD44 GFAP

13–22 14–22 14–21 15 16, 19–20 20–25 21 25

16 16 10, 56a, 74 10 10, 56a 37 74 56a

Astrocytes CD44 GFAP Interleukin-8 receptor

16–21 24 16

74 75 76

Microglia MHC class I antigen MHC class II antigen CD45

10–25 10–25 10–25

71 71, 72 71, 72

Macrophage Mac S22

14–25

71

CRALBP = Cellular retinaldehyde binding protein; GFAP = glial fibrillary acidic protein; MHC = major histocompatibility complex. Gestational age range denotes the period of IR expression studied by one or several workers. a Observation from a transplant study.

in table 3. The adult pattern of the human retinal vasculature is attained around the 8th fetal month [73], though maturation occurs only after birth, a process that is different from the monkey retina showing prenatal vascular maturation [60].

Synaptic Development

The IPL forms at 9 WG in the central retina [77] and synapses appear in it at around 12 WG. By 11 WG, a thin OPL forms in the macula, separating the INL from a single row of cone precursors [78]. Synaptic ribbons and vesicles differentiate and become prominent in their basal cytoplasm at around 15 WG [7]. By labeling with markers of synaptogenesis (synaptophysin and syntaxin-1), Nag and Wadhwa [57] showed that IR to both markers is present in the central IPL at 11–12 WG. This indicates that both proteins are incorporated into respective synaptic 25

Fig. 4. Localization of syntaxin-1 and syn-

aptophysin IR in the fetal human retina at 20 WG. a, b Syntaxin-1 IR in the central (a) and peripheral part (b) of the retina, in the IPL and OPL of the central retina, and in the IPL of the peripheral retina. In the latter, the OPL is yet to differentiate. c, d Synaptophysin IR in the central (c) and peripheral part (d) of the retina. In both, IR in the IPL is indicated. Cones with their terminals are immunopositive in the central OPL (c) whereas in the peripheral retina, immature photoreceptor processes show fine granular staining (asterisk). v = Vitreal side. Scale bar: 50
m. (From [57], J Biosci 2001;26:179–191, reproduced with permission of the publisher)

elements in parallel with morphological differentiation of the synapses at this time point. In the OPL, however, IR becomes prominent by 16 WG, consistent with electron-microscopic data of appearance of synaptic vesicles and ribbons in photoreceptors. While synaptophysin is likely to be present in synaptic vesicle membranes (amacrine, bipolar cell axon and photoreceptor cell terminals), syntaxin-1 is presumably present along the presynaptic membranes at chemical synapses, but reportedly absent from the ribbon synapses of mammalian retina [79, 80]. In the OPL, the cone pedicles, which make conventional synapses with dendrites of flat bipolar cells [81, 82], may contain syntaxin-1 along their basal presynaptic membranes. Immunolabeling with marker synaptic proteins suggests that IPL matures before OPL in the human retina (fig. 4), as in the primates. The marker expressions in both plexiform layers follow a central to peripheral gradient of synaptic development (fig. 4), as reported for the cat and monkey retina [83, 84]. In the human fetal retina, at 15 WG, the IPL consists of well-differentiated synapses of amacrine cells and immature, monad bipolar cell synapses. Dyad bipolar synapses are few and perhaps appear late in fetal life [42]. Puncta adherentia and other intercellular junctions are also present. Synaptogenesis of the 26

Neuroembryol Aging 2006–07;4:19–30

human retina is not studied beyond this period by electron microscopy, and the entire sequence of synaptic development till midgestation, when bipolar and horizontal cells are well differentiated, remains to be looked for.

Neuroactive Substances and Neuronal Differentiation

Calcium-Binding Proteins The distribution of three calcium-binding proteins, viz calbindin, parvalbumin and calretinin, was examined in human fetal retinas [14, 30, 31]. Calbindin IR is present in some horizontal cells and GCs during differentiation and migration (fig. 5), while parvalbumin IR occurs in most GCs, all horizontal cells and few amacrine cells from 13 to 15 WG onward [30, 31]. At 20–21 WG, calretinin IR is mainly localized in most differentiated amacrine and horizontal cells. The calretinin-positive horizontal cells are smaller and less frequent in distribution than the calbindin- and parvalbumin-positive counterparts. No horizontal cells are calretinin positive in the postnatal or adult retinas [14]. These facts suggest a critical role for calretinin in the maturation of a type of horNag/Wadhwa

izontal cells, whose identity is unknown at present. On the other hand, the expression of parvalbumin IR in most differentiating horizontal cells suggests a possible involvement of this protein in their differentiation. Yan and Wiechmann [32] showed small populations of recoverin-positive cells in the GCL and nerve fiber layer. According to them, in the fetal retina recoverin may play a role in determining the polarity of the differentiating GC and axonal growth in a centrifugal spatiotemporal order. Some of these recoverin-positive cells may also be photoreceptors that have migrated incorrectly. GABA GABA is expressed at 12 WG in many cells of the inner neuroblastic layer (GCs) and in a few axons of the nerve fiber layer, which suggests that some of the stained cells could be prospective GCs. From 16 to 25 WG, intense IR is found in most amacrine and horizontal cells in the central to midperipheral retina. In the postnatal retina, some horizontal cells are moderately labeled, but no IR is found in the adult human retina [11], consistent with the known patterns in mammals [85]. Thus, there is a possibility that GABA plays a role in horizontal cell differentiation in the human retina. Furthermore, whereas in the human fetal retina many GCs and amacrine cells are GABA positive, in the postnatal and adult retina few of these cells show GABA IR. The role of GABA in early neuronal development as a neurotrophic factor for maturation of the neurons as well as for the generation and maintenance of spontaneous activity waves in the period of functional network formation is becoming evident [86].

Fig. 5. Immunohistochemical demonstration of calbindin in the fetal retina at 20 WG. In the central part (a), calbindin IR is pres-

ent in the differentiated horizontal cells (arrows) and GCs (arrowheads). In the peripheral retina (b), the migrating horizontal cells show IR (arrowheads). Scale bar: 50
m.

Nitric Oxide Synthase and Glutamate A transient increase in neuronal nitric oxide synthase IR in GCs is reported to occur between 18 and 25 WG of fetal human retina [37]. This coincides with the period of naturally occurring cell death in the human fetal GCL [26]. GC loss is substantiated by an excessive reduction in the number of their axons between 16 and 20 WG and a gradual loss up to 26 WG of prenatal life [20, 87]. At 16–17 WG, IR to glutamate and parvalbumin (that binds glutamate-released Ca2+) increases in the GCL of fetal human retina [30, 33]. Thus, cell death in the GCL is perhaps linked to excessive glutamate and nitric oxide activity, as proposed earlier [88]. In adult human retina, glutamate is present in photoreceptors, bipolar cells and GCs [89]. In fetal human retina, additionally, glutamate IR appears in horizontal and amacrine cells. During development, glutamate exerts a

variety of effects such as neuronal migration, neurite sprouting and maturation, cell death and synapse elimination [90–92]. In fetal human retina, glutamate may serve a developmental role for amacrine and horizontal cells, as was reported for the rabbit retina [93].

Development of the Human Neural Retina

Neuroembryol Aging 2006–07;4:19–30

Taurine In the human fetal retina, the GCs and some of their axons show taurine IR at 16–17 WG. From 20 to 21 WG, IR develops in Müller cell endfeet and in some amacrine, bipolar and horizontal cells, which increases with progressive fetal age (24–25 WG [36]). In postnatal infant retina, taurine IR is present in some amacrine cells and strongly in photoreceptors, but absent in GCs and horizontal cells. In human fetal retina, expression of taurine IR follows a sequence that parallels with neuronal differentiation in an inner (vitreal) to outer (scleral) direction. 27

Table 4. Interneuron markers in the human fetal retina

Markers

Gestational age, WG

Retinal location

References

GABA (A, H) Parvalbumin (Aa, Hc) Synaptophysin (Bc) Trk A, B, C (A, H) Calbindin (Aa, Hb, Ba) nNOS (Aa) Calretinin (Ab, Ha) Glutamate (Ab, Ha, Bb) TH (Ab) TH (Ab) C-fos (Ab) Neuropeptide Y Somatostatin Substance P Leucine-enkephalin

12–25 14–25 14–25 16–21 16–25 16–25 16–25 20–25 10–21 24 15–40 14–40 14–40 14–21 16–21

central to peripheral central to peripheral foveal to peripheral central to peripheral central, peripheral central, midperipheral central to peripheral midperipheral midperipheral central, midperipheral central, midperipheral midperipheral central, midperipheral midperipheral central, peripheral

11 30, 31 40, 57 15 30, 31 37 14 34 12 13 38 12, 35 34, 35 12, 29 12, 29

A = Amacrine; B = bipolar; H = horizontal; nNOS = neuronal nitric oxide synthase; TH = tyrosine hydroxylase. Wherever indicated, the superscripts indicate IR detected in some cells (a), many cells (b) and in most cells (c). Gestational age range denotes the period of IR expression studied by one or several workers.

It is likely that this amino acid is vital for retinal cell differentiation, as was shown in cultures [94]. Neuropeptides and Aminergic Transmitters The earliest fetal age to show neuropeptide IR in the human retina is 10 WG, when substance P, enkephalin and serotonin-positive cells are seen in the outer neuroblastic layer and tyrosine hydroxylase-immunopositive amacrine cells in the INL [12, 13]. During 17–24 WG, IR appears in the ONL, INL and IPL. By 26–30 WG, neurons in the GCL and the fovea are positive for substance P and enkephalin [29]. The IR to all neuropeptides occurs in a sequence from outer to inner layers of the developing human retina. Neuropeptide Y-immunopositive cells and fibers are seen in the INL and IPL from 15 WG till term [35]. All these reports indicate that neurotransmitters (tyrosine hydroxylase and serotonin) and neuropeptides (substance P, neuropeptide Y and enkephalin) play some role in neuronal differentiation. Colocalization studies of neurotransmitters with neuropeptides during human retinal development are not carried out. This aspect may shed light on the possible interactions between transmitter candidates and neuropeptides and their role in neuronal development. The expression of various markers of interneurons in the human fetal retina is summarized in table 4.

28

Neuroembryol Aging 2006–07;4:19–30

Conclusion

The availability of antigenic markers has provided a fresh impetus and opportunity to retinal researchers, to learn about the neurochemical development of the human retina. The review of data demonstrates a changing IR pattern of the amino acids and their derivatives, neurotransmitter candidates and neuropeptides in the developing human retina, which is relevant to comprehend the neurochemical differentiation in human retinal transplants [95] before use for therapeutic purposes. More indepth studies are, however, needed to understand the involvement of various neurochemical substances in retinal cell differentiation.

Nag/Wadhwa

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Development of the Human Neural Retina

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49 van der Spuy J, Kim JH, Yu YS, Szel A, Luthert PJ, Clark BJ, Cheetham ME: The expression of the Leber congenital amaurosis protein AIPL1 coincides with rod and cone photoreceptor development. Invest Ophthalmol Vis Sci 2003;44:5396–5403. 50 Cornish EE, Hendrickson AE, Provis JM: Distribution of short-wavelength-sensitive cones in human fetal and postnatal retina: early development of spatial order and density profiles. Vision Res 2004;44:2019–2026. 51 Cornish EE, Xiao M, Yang Z, Provis JM, Hendrickson AE: The role of opsin expression and apoptosis in determination of cone types in human retina. Exp Eye Res 2004;78: 1143–1154. 52 Furukawa T, Morrow EM, Cepko CL: Crx, a novel otx-like homeobox gene, shows photoreceptor-specific expression and regulates photoreceptor differentiation. Cell 1997; 91: 531–541. 53 Swain PK, Hicks D, Mears AJ, Apel IJ, Smith JE, Hendrickson AE, Milam AH, Swaroop A: Multiple phosphorylated isoforms of NRL are expressed in rod photoreceptors. J Biol Chem 2001;276:36824–36830. 54 Bibb LC, Holt JK, Tarttelin EE, Hodges MD, Gregory-Evans K, Rutherford A, Lucas RJ, Sowden JC, Gregory-Evans CY: Temporal and spatial expression patterns of the CRX transcription factor and its downstream targets. Critical differences during human and mouse eye development. Hum Mol Genet 2001;10:1571–1579. 55 Mears AJ, Kondo M, Swain PK, Takada Y, Bush RA, Saunders TL, Sieving PA, Swaroop A: Nrl is required for rod photoreceptor development. Nat Genet 2001;29:447–452. 56 Seiler MJ, Aramant RB: Photoreceptor and glial markers in human embryonic retina and in human embryonic retinal transplants to rat retina. Dev Brain Res 1994;80:81–95. 57 Nag TC, Wadhwa S: Differential expression of syntaxin-1 and synaptophysin in the developing and adult human retina. J Biosci 2001;26:179–191. 58 Kim JH, Yu YS, Kim JH, Kim YJ, Heo JW, Kim CJ: Immunoreactivity of constitutive and inducible heat shock protein 70 in human fetal retina. Korean J Ophthalmol 2003; 17:14–18. 59 Ni M, Yamaki K, Kikuchi T, Ferrick M, Shinohara T, Nussenblatt RB, Chan CC: Developmental expression of S-antigen in fetal human and rat eye. Curr Eye Res 1992; 11: 219–229. 60 Gariano RF, Iruela-Arispe ML, Hendrickson AE: Vascular development in primate retina: comparison of laminar plexus formation in monkey and human. Invest Ophthalmol Vis Sci 1994;35:3442–3455. 61 Provis JM: Development of the primate retinal vasculature. Prog Ret Eye Res 2001; 20: 799–821. 62 Hughes S, Yang H, Chan-Ling T: Vascularization of the human fetal retina: roles of vasculogenesis and angiogenesis. Invest Ophthalmol Vis Sci 2000;41:1217–1228.

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63 Chan-Ling T, McLeod DS, Hughes S, Baxter L, Chu Y, Hasegawe T, Lutty GA: Astrocyteendothelial cell relationships during human retinal vascular development. Invest Ophthalmol Vis Sci 2004;45:2020–2032. 64 Sandercoe TM, Madigan MC, Billson FA, Penfold PL, Provis JM: Astrocyte proliferation during development of the human retinal vasculature. Exp Eye Res 1999; 69: 511– 523. 65 Ashton N: Retinal angiogenesis in the human embryo. Br Med Bull 1970;26:103–106. 66 Provis JM, Leech J, Diaz CM, Penfold PL, Stone J, Keshet E: Development of the human retinal vasculature: cellular relations and VEGF expression. Exp Eye Res 1997;65:555– 568. 67 Penfold PL, Provis JM, Madigan MC, van Driel D, Billson FA: Angiogenesis in normal human retinal development: the role of astrocytes and macrophages. Graefes Arch Clin Exp Ophthalmol 1990;228:255–263. 68 Gogat K, Le Gat L, Van Den Berghe L, Marchant D, Kobetz A, Gadin S, Gasser B, Quere I, Abitbol M, Menasche M: VEGF and KDR gene expression during human embryonic and fetal eye development. Invest Ophthalmol Vis Sci 2004;45:7–14. 69 Chu Y, Hughes S, Chan-Ling T: Differentiation and migration of astrocyte precursor cells and astrocytes in human fetal retina: relevance to optic nerve coloboma. FASEB J 2001;15:2013–2015. 70 Chan-Ling T, Gock B, Stone J: The effect of oxygen on vasoformative cell division: evidence that ‘physiological hypoxia’ is the stimulus for normal retinal vasculogenesis. Invest Ophthalmol Vis Sci 1995;36:1201–1214. 71 Diaz-Araya CM, Provis JM, Penfold PL: Ontogeny and cellular expression of MHC and leucocyte antigens in human retina. Glia 1995;15:458–470. 72 Diaz-Araya CM, Provis JM, Penfold PL, Billson FA: Development of microglial topography in the human retina. J Comp Neurol 1995;363:53–68. 73 Fedorov AA: Prenatal development of human retinal vessels (in Russian). Vestn Oftalmol 2003;119:59–63. 74 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. 75 Karim MM, Fujisawa K, Kanomata N, Ito H: Glial fibrillary acidic protein expression in the developing human retina. Kobe J Med Sci 1996;42:111–117. 76 Dame JB, Juul SE: The distribution of receptors for the pro-inflammatory cytokines interleukin (IL)-6 and IL-8 in the developing human fetus. Early Hum Dev 2000;58:25–39. 77 Spira AW, Hollenberg MJ: Human retinal development: ultrastructure of the inner retinal layers. Dev Biol 1973; 1:1–21. 78 Linberg KA, Fisher SK: A burst of differentiation in the outer posterior retina of the eleven-week human fetus: an ultrastructural study. Vis Neurosci 1990;5:43–60.

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79 Brandstätter JH, Lohrke S, Morgans CW, Wässle H: Distributions of two homologous synaptic vesicle proteins, synaptoporin and synaptophysin, in the mammalian retina. J Comp Neurol 1996;370:1–10. 80 Morgans CW: Presynaptic proteins of ribbon synapses in the retina. Microsc Res Tech 2000;50:141–150. 81 Dowling JE: The Retina: An Approachable Part of the Brain. Cambridge, Harvard University Press, 1987. 82 Missotten L: The Ultrastructure of the Human Retina. Brussels, Arscia, 1965. 83 Maslim J, Stone J: Synaptogenesis in the retina of the cat. Brain Res 1986;373:35–48. 84 Okada M, Erickson A, Hendrickson A: Light and electron microscopic analysis of synaptic development in Macaca monkey retina as detected by immunocytochemical labeling for the synaptic vesicle protein, SV2. J Comp Neurol 1994;339:535–558. 85 Johnson DA: GABA in the retina; in Martin DL, Olsen RW (eds): GABA in the Nervous System. The View at Fifty Years. Philadelphia, Lippincott Williams & Wilkins, 2000, pp 459–470. 86 Jelitai M, Madarasz E: The role of GABA in early neuronal development; in Dhossche DM (ed): GABA in Autism and Related Disorders. London, Elsevier Academic Press, 2005, pp 28–52. 87 FitzGibbon T: The human fetal retinal nerve fiber layer and optic nerve head: a DiI and DiA tracing study. Vis Neurosci 1997; 14: 433–447. 88 Huxlin KR, Bennett MR: NADPH diaphorase expression in the rat retina after axotomy – a supportive role for nitric oxide? Eur J Neurosci 1995;7:2226–2239. 89 Crooks J, Kolb H: Localization of GABA, glycine, glutamate and tyrosine hydroxylase in the human retina. J Comp Neurol 1992;315: 287–302. 90 Lipton SA, Kater SB: Neurotransmitter regulation of neuronal outgrowth, plasticity and survival. Trends Neurosci 1989;12:265–270. 91 Rabbachi S, Bailly Y, Delhaye-Bouchard N, Mariani J: Involvement of NMDA receptor in synapse elimination during cerebellar development. Science 1992;256:1823–1825. 92 Rakic P, Cameron RS, Komuro H: Recognition, adhesion, transmembrane signaling and cell motility in guided neuronal migration. Curr Opin Neurobiol 1994; 4:63–69. 93 Redburn DA, Agarwal SH, Messersmith EK, Mitchell CK: Development of the glutamate system in rabbit retina. Neurochem Res 1992;17:61–66. 94 Altshuler D, LoTurco JJ, Rush J, Cepko C: Taurine promotes the differentiation of a vertebrate retinal cell type in vitro. Development 1993;119:1317–1328. 95 Yang P, Seiler MJ, Aramant RB, Whittemore SR: In vitro isolation and expansion of human retinal progenitor cells. Exp Neurol 2002;177:326–331.

Nag/Wadhwa

Neuroembryol Aging 2006–07;4:31–60 DOI: 10.1159/000103451

Published online: June 4, 2007

Review of Development and Aging in the Eye of Teleost Fish Yvette W. Kunz Maurice Kennedy Research Center for Emeritus Staff, University College Dublin, University of Ireland, Dublin, Ireland

Key Words Circadian rhythms
Color vision
Cone apoptosis
Embryonic fissure
Peripheral growth zones
Photoreceptor disk shedding
Retinomotor movements
Ultraviolet vision

Abstract The teleost eye is an excellent organ for studying retinal development since teleosts and their eyes continue to grow throughout life. The retinal growth zones are found at the periphery of the eye and in some species also along the ventral embryonic fissure. The retinal cones are arranged into regular mosaic patterns, which is an advantage for microspectrophotometrical analyses of visual pigments. They revealed that teleost eyes are di-, tri- and tetrachromatic. The last type includes ultraviolet-sensitive cones. They appear in early life over the whole retina and later disappear in this region although they continue to be formed in the peripheral growth zones, but die there once the retina is differentiated. The melanin granules in the pigment epithelium and the cones and rods undergo circadian retinomotor movements. Another circadian rhythm is displayed by the tips of the photoreceptor outer segments, which are shed by cones at night and by rods during the day. The embryonic development of the retina is described in detail for the viviparous guppy, the salmonids and the zebrafish, which display direct development. This is contrasted by results of teleosts with indirect development (involving metamorphosis). Copyright © 2007 S. Karger AG, Basel

© 2007 S. Karger AG, Basel 1661–3406/07/0042–0031$23.50/0 Fax +41 61 306 12 34 E-Mail [email protected] www.karger.com

Accessible online at: www.karger.com/nba

Introduction

This publication presents an overview of earlier comprehensive analyses of the development of the fish eye [1–3] updated with relevant results of recent publications. The isolated retina of the teleost eye, the ‘forgotten part of the CNS’, is an excellent model for studying neural mechanisms because these fish, and their eyes, grow throughout life by stretching of the retina and by peripheral cell addition. In other words, the retinal growth zones (peripheral growth zones, PGZ) recapitulate retinal genesis. They are also engaged in retinal repair. Damaged cells undergo apoptosis accompanied by removal of cell debris and the simultaneous creation of new neurons. Continuous peripheral cell addition allows the ontogeny of the retina to be followed at any age of the fish by analyzing the PGZ. However, due to the continual growth of the fish, the aging process has been largely unexplored. Bielmaier et al. [4] observed that in ‘adult’ zebrafish ‘practically no apoptosis was present in the central retina’. A recent publication of functional aging and gradual senescence in zebrafish shows that this species displays very gradual senescence [5]. Genade et al. [6] suggest that the annual fish Nothobranchius furzeri, with a lifespan of only 3 months, shows accelerated growth and expression of aging biomarkers at the level of histology and behavior. The last two publications did not deal with an analysis of the eye. Astyanax fasciatus is used for studies on teleost eye development because populations living in surface streams

Yvette W. Kunz ‘Festina lente’, Ballybawn, Kilmacanogue Co. Wicklow (Ireland) Tel. +353 1 286 7413, E-Mail [email protected]

A

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Fig. 1. Diagram of a teleostean eye (P. reticulata) in a dorsoventral

section through the optic nerve (ON). A–C = Differently structured layers of the sclera; BP = black patch, sclera devoid of iridophores; C’ = cornea; CG = choroid gland; L = lens; R = retina; RL = retractor lentis muscle; SC = sclera; VH = vitreous humour. (From Ennis and Kunz [12], with kind permission of Elsevier.)

GCL

Fig. 2. Radially sectioned retina of P. reticulata. C = Cones; R =

have large eyes and pigmentation whereas those adapted to caves have lost their eyes and pigmentation. In the cave fish embryo, eye development is initiated but the eye arrests later and degenerates. Their embryonic lens cells suffer extensive programmed cell death, which affects the development of the eye. Transplantation experiments have shown that a lens obtained from a surface fish donor is sufficient to rescue eye development in the host [7]. Another functional, though non-image-forming, photodetector in teleosts is the pineal organ, lying on the dorsal surface of the brain. In many species, it differentiates earlier than the retina. It contains photoreceptor inner (IS) and outer segments (OS), with the latter protruding into its lumen. The overlying region of the cranium may lack melanophores, thereby forming the so-called ‘pineal window’ [8–10]. The pineal organs of the Atlantic halibut Hippoglossus hippoglossus appear to have the potential for both ultraviolet (UV) and green photosensitivity at the embryonic stage [11]. 32

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rods. (From Kunz [13], with kind permission of the Editor of the Revue Suisse de Zoologie.)

The Differentiated Teleostean Eye

In order to understand the development of the teleostean eye, it is essential to give a short description of the differentiated eye (fig. 1). Since teleosts grow throughout life it is misleading to refer to eyes of ‘adult’ fish; the Latin word adultus means grown up. In fish with indirect development, e.g. flatfish and eels, one should distinguish between larval, metamorphic and mature stages. When describing the differentiated eye of fish with direct development (e.g. salmonids), the age, weight or length of the fish should be given. (Some authors of studies on salmonids mistakenly refer to ‘metamorphosis’ when they describe smoltification.) Kunz

The spherical lens is used only for focusing since the refractive index of the cornea is identical to that of water. At rest the eye is set for near vision and accommodation occurs for distant vision. Accommodation is achieved not by changing the shape of the spherical lens but by its backward movement in response to contraction of the ventrally attached retractor muscle (fig. 1). The teleostean retina is avascular, i.e. not traversed by blood vessels. However, a high-pressure dissolved oxygen is available for its high metabolic demand by a series of vascular connections. In the trout, the choroid rete mirabile is supplied with doubly oxygenated blood by the ophthalmic artery which is the efferent blood vessel of the pseudobranch. The pseudobranch (a hemigill) in turn is fed by the efferent artery of the first gill arch. Not all fishes possess a pseudobranch [e.g. ‘adult’ eels (in contrast to larval eels) and catfish lack them]. In addition, some teleosts possess a vascular falciform process which receives its blood from the lentiform body (another countercurrent system, which in turn receives its blood from the choriocapillaris). If the falciform process is absent or reduced, the hyaloid artery supplies a capillary network expanding over the internal surface of the retina. A radial section through the differentiated retina of Poecilia reticulata displays the different retinal layers (fig. 2). Adjacent to Bruch’s membrane is the retinal pigment epithelium (RPE). Its granules take part in lightand dark-adaptation (so-called retinomotor movements). The visual cells (photoreceptors) are made up of rods and different types of cones. Rods are prevalent in fish with dim vision (nocturnal, deep-sea and cave fish). In some of these fish, the rods are arranged in tiers. The photopigment of rods is either rhodopsin or porphyropsin or both. Karl von Frisch [14] was the first to maintain in several publications that fish see color to contradict the ophthalmologist Hess [15, 16], who had continuously stressed that fish are color blind. Four morphologically different cone types can be identified: short, intermediary, long single cones and double cones. Sporadically, triple and quadruple cones are observed. Microspectrophotometry (MSP), the direct measurement of visual pigment in individual photoreceptors, has shown fish with di-, tri- and tetrachromatic vision. It revealed that double cones are sensitive to long wavelengths while single cones usually contain shorterwavelength-sensitive visual pigments. However, no general correlation between single cone morphology and spectral sensitivity can be demonstrated. Many fish pos-

sess UV-sensitive single cones (UVS cones) with a peak sensitivity at about 360 nm (fig. 3a). In the 1-year-old brown trout and the young salmon they occupy the corners of the cone square mosaic (figs. 3b, 8). The cones of teleosts are usually arranged in a row or square mosaic pattern with the rods distributed at random. Triple cones tend to occupy the same position as double cones within the mosaic. In P. reticulata, the square mosaic changes into a row mosaic during dark adaptation. The mechanism of pattern formation in teleosts has been analyzed using mathematical models [19]. Some of the cone-IS of the guppy P. reticulata show intracellular color filters called ellipsosomes. They are of mitochondrial origin and have a function very similar to the colored oil droplets of birds and reptiles. However, instead of oil they contain the heme pigment cytochrome C and are thought to enhance color contrast in the blueviolet portion of the spectrum (fig. 4). A pure-cone fovea (shallow depression to pronounced deepening) in the temporal region of the retina has been reported. However, most fish have instead an ‘area’, i.e. a site of high cone density in the absence of rods. The guppy has three areae, one each in the temporal, ventral and fundic region. In some deep-sea fish, a pure rod fovea has been described. Areae with maximum cone density show an increased thickness of the retina and a maximum number of cells in the inner nuclear and ganglion cell layers. Retinomotor movements, a substitute for pupillary movements, occur during light and dark adaptation. Rods and cones undergo extension and contraction in a daily circadian rhythm. During the day, the melanin granules of the RPE move vitreally and rods extend into it; during the night, the melanin granules move sclerally, cones extend and rods move vitreally (fig. 5). A further circadian rhythm involves shedding of the tips of the photoreceptor OS. Rods shed during the day and cones at night (fig. 6). The outer nuclear layer (ONL) contains the photoreceptor nuclei; the large oval cone nuclei are situated more sclerally than the small round rod nuclei. The UVS cone nuclei lie almost at the level of the rod nuclei (fig. 3c). The terminals of the photoreceptors synapse with bipolar neurons in the outer plexiform layer (OPL). The inner nuclear layer (INL) consists of somata of the horizontal, bipolar, Mueller, interplexiform and amacrine cells. In the inner plexiform layer (IPL) synapses between the amacrine and ganglion cells are found. The axons of the ganglion cell layer (GCL) leave the eye as the optic nerve, which can be pleated as in Trachinus vipera (fig. 1).

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33

Absorbance (%)

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my d

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Fig. 3. a Mean absorbance spectra for the four cone pigments in the 1-year-old brown trout, S. trutta. Maximum absorbance values (
max): single corner cone, 355 nm (n = 6), single central cone, 441 nm (n = 16), green-sensitive members of double cone, 535 nm (n = 8); red-sensitive members of double cone, 600 nm (n = 5). b–d S. trutta photoreceptors. b Diagram of a section through the IS region in a yearling trout to show the square mosaic with dou-

34

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ble cones forming the sides, and single cones at the center (CE) and corners (CO). c Diagram of radially sectioned visual cells showing from right to left: rod (R), double cone (DC), two types of single cones (CE, CO) with OS, IS separated into ellipsoid (el) and myoid (my), nucleus (N), synapse (SY). d Diagram from a 2year old trout showing that the corner cones are absent. (From Bowmaker and Kunz [17], with kind permission of Elsevier.)

Kunz

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Fig. 4. TEM of double cones of P. reticulata showing a highly dense ellipsosome in a double cone of the ventral area. !7,000. E = ellipsosome; M = mitochondria; SS = subsurface membranes. Inset showing diagram of a dorsoventral section through the optic nerve (ON). (From Kunz and Wise [18], with kind permission of Birkhäuser Verlag, Basel.)

Lactate Dehydrogenase Pattern of Trout and Guppy Eye In teleosts, the glycolytic enzyme lactate dehydrogenase (LDH) presents three loci (A, B and C), with the C4-

isozyme being primarily expressed in retinal photoreceptors and optical parts of the brain. It may play an important role in the regeneration of the visual pigment (fig. 7).

Development and Aging in the Eye of Teleost Fish

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Fig. 7. Isozyme pattern of LDH in different organs of sexually ma-

ture P. reticulata. Arrow denotes point of application of the sample. # = Bands unique to eye and brain; – = cathode; + = anode. (From Kunz [13], with kind permission of the Editor of the Revue Suisse de Zoologie.)

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Fig. 6. Developmental pattern of phagosome count (i.e. shedding rhythm of OS tips) in the brown trout (S. Trutta). a Stage D. b stage E. c Ambient light conditions for a and b. d Stage F. e Ambient light conditions for d. f One-year-old trout. Broken line re-

fers to small and solid line to large phagosomes. ROS shed their disks during the day and COS at night. (From McCormack et al. [20], with kind permission of S. Karger AG, Basel.)

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37

UV

100

Blue

Green

Red

Relative OD

80

60

40

20

Fig. 8. Absorbance spectra obtained by MSP of four cone types in the young salmon. Above: diagram of square mosaic obtained by section through the ellipsoid region of the cones. OD = Optical density [from Loew and Kunz, unpubl.].

0 350

400

Age-Related Loss (by Apoptosis) of Ultraviolet-Sensitive Cones in the Main Retina

The yearling trout possesses over the whole retina a square mosaic with UVS cones occupying the corners of the square mosaic. On the basis of a publication by Fürst [21] in 1904, which mentions that corner cones are absent in older salmon, a 2-year-old trout was analyzed. Histological results showed that the corner cones had disappeared, which coincided with an absence of UV sensitivity demonstrated by MSP (fig. 3). Behavioral experiments in the rainbow trout confirmed a similar loss. Trout that lose UV vision with age show a parallel increase in absorption of UV light by their lens, thus protecting their retina. Similarly, the Atlantic salmon, Salmo salar, at the age of up to 120 days contains in its eye square mosaics with a full complement of cones. MSP analysis confirmed the presence of UVS cones in these young salmon, while they were absent in older fish (fig. 8). A 38

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450

500

550 600 Wavelength (nm)

650

700

750

short-wavelength-sensitive cone mechanism was also found in juvenile goldfish, yellow perch, Perca flavescens and Lepomis macrochirus, while it disappeared in all three adult fish. Electron-microscopical (EM) analyses revealed that in S. salar older than 120 days (length 1 4.5 cm) the UVS cones begin to disappear as a result of apoptosis. Pyknotic nuclei and densely clumped cytoplasm are indicative of degenerating cells. When the fish are 230 days old (length 9 cm), the corner cones are absent over the whole retina and the vacated spaces have been taken over by newly formed rods (fig. 9). It has been suggested that this loss of UV sensitivity might reflect either a change in feeding behavior or a change in polarization sensitivity. However, the disappearance of UVS cones is associated with a change in habitat, i.e. the salmon parr move to deeper waters in natural conditions. The effect of this change is demonstrated by the fact that UV light induces severe epidermal necrosis in hatchery-reared salmon Kunz

Fig. 9. EM showing that rods have taken up the sites of former corner cones in the retina of the salmon. CE =

Central cone; DC = double cone; R = rod; R’ = rod occupying the position of a former corner cone; V = sites vacated by corner cones. (From Kunzet et al. [22], with kind permission of Elsevier.)

moved too late from their hatchery trays to deeper outside ponds. In the retina of Oncorhynchus mykiss, UVS cones disappear during smoltification, which prepares them for deeper native waters. Near sexual maturation and the return migration to natal streams, UVS cones reappear [23]. A reduction in the number of UVS cones in older rainbow trouts was shown to be consistent with decreased sensitivity to UV light by electroretinograms [24]. The role of thyroid hormones in the migratory Atlantic salmon showed that elevated thyroxine levels imme-

diately precede changes in the digestive tract and habitat of the fish. The first peak precedes the transition from the yolk-dependent stage to the predatory stage and the second peak is correlated with smoltification. However, the above results on the retina of S. salar show that the disappearance of the UVS cones starts 3 months after the onset of exogenous food uptake and finishes 4 months before smoltification. The disappearance of one third of the cones in the main retina of Salmo trutta (aged 12–20 months) and in the eye of 4- to 7.5-month-old Atlantic salmon raises the question as to how the loss of synaptic contact with, pre-

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39

about 200 g [28]. Different studies on Oncorhynchus nerka have produced conflicting results with regard to the loss of UV sensitivity [29]. However, in situ labeling of the UVS cones confirms the putative identity of the disappearing corner cells. UVS opsin mRNA was localized to single corner cones in the PGZ of Oncorhynchus mykiss [24]. A review of the UV visual world of fishes was given by Losey et al. [30] in 1999. Teleost fish can also regenerate neural tissue after damage. Damaged cells undergo apoptosis accompanied by removal of cell debris and the simultaneous creation of new neurons. Fig. 10. Diagram of embryonic (chorionic) fissure. The ophthalmic artery enters the eye through the fissure. The eyecup has induced the overlying ectoderm to form a solid lens. 1 = Lens; 5 = fissure; 6 = ophthalmic artery. (From Portmann [31], with kind permission of Schwabe Verlag, Basel.)

sumably, horizontal and bipolar cells affects the retinal circuitry and finally the retinal mapping in the optic tectum. As mentioned above, the spaces vacated by the degenerated UVS cones are taken up by rods (fig. 9). Their progenitor cells are located in the INL and have migrated to the ONL where they divide and differentiate into rod precursors (fig. 13). In the cichlid Haplochromis burtoni it was shown that the insulin-like growth factor 1 produced by cone photoreceptors regulates rod progenitor proliferation in the teleost retina in a diurnal rhythm (rod progenitor cell division is threefold greater at night than during the day) [25]. Apart from the rim (PGZ), new rods are continuously inserted over the whole retina, which stretches during the life span of the fish. Contrary to the above results, in the anadromous and non-anadromous salmonid fish O. mykiss the UVS cones in the ventral retina disappear during smoltification while UV sensitivity remains intact in the dorsal retina. However, it may also be explained by a complete loss of UVS cones followed by cone regeneration in the dorsal retina. Since UV light reaches the ventral part of the eye, the question as to why UVS cones should remain in the dorsal part of the retina arises. Thyroxine treatment of juvenile O. mykiss mimics smoltification and causes a reduction in UV sensitivity [24, 26, 27]. Electrophysiological recordings from the optic nerve showed that while UV sensitivity is greatly diminished at the smolt stage it reappears in ‘adult’ fish weighing 40

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The Growth Zones in the Fully Differentiated Teleostean Eye

As mentioned above, the eyes of teleosts grow throughout life by stretching and cell addition along the rim (ora serrata), also referred to as PGZ. In species with an open ventral embryonic (choroid) fissure (coloboma), e.g. salmonids, this area persists as an additional site of neurogenesis (fig. 10). However, in other fishes the fissure can be more or less closed. For instance in Tilapia mossambica it is lined with PE only and in the guppy P. reticulata the fissure closes completely during early development. Surface viewing of the developing photoreceptors in the rim (PGZ) and in the open fissure of the brown trout (S. trutta) and the Atlantic salmon (S. salar) reveals the youngest (most peripheral) area of the growth zone with only cone inner segments (CIS), followed by cones with a connecting cilium (CC) and, finally, cones with OS (COS). Rods develop at a lower (more vitreal) level than the cones and are not seen by surface viewing. However, when fully developed, i.e. bordering the main retina, the rods move sclerally during daytime and their long outer segments (ROS) now obscure COS (fig. 11a, b). In the most peripheral part of the growth zones, the cones are arranged in a row mosaic, i.e. the double cones with their subsurface membranes and single cones (alternating future central and corner cones) are arranged in a straight line (row mosaic; figs. 12a, 13). In the adjacent zone, i.e. nearer the main retina, this mosaic changes into a square mosaic (figs. 12b, 13). Further centrally, but still in the growth zone, the corner cones begin to disappear and are replaced by newly developing rods (fig. 13). In other words, UVS cones are generated, differentiated and disappear within the PGZ and along the embryonic fissure in a conveyor-belt fashion. A diagrammatic representation of these developmental areas is given in figure 13. Kunz

Fig. 11. a Scanning EM of the growth zone

along the ventral embryonic fissure of the eye of the brown trout (S. trutta) and the Atlantic salmon (S. salar). Bar = 10 m. b Scanning EM of the growth zone in the rim (PGZ) of the eye of S. trutta and S. salar. RM = Row mosaic; SM = square mosaic. (From Kunz [32], with kind permission of Birkhäuser Verlag, Basel.)

Transmission electron microscopy (TEM) clearly displays apoptosis of the various parts of the UVS cones, which is always heralded by a densification of the parts involved. A dying OS of a UVS cone is shown in figure 14a and a degenerating inner segment (IS) filled with mito-

chondria in figure 14b. Electron-dense decaying myoid, nucleus and pedicle are illustrated in figure 15. Parts of the pyknotic nuclei are taken up by macrophages or Mueller cells (fig. 16).

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Fig. 12. Diagrams and Scanning EM of figure 11 at higher magnification to show the arrangements of row and square cone mosaics. a Row mosaic (RM). b Square mosaic (SM). Bar = 1 m. CE = Central cone; CO = corner cone; DC = double cone; R = rod; SC = Single cone. (From Kunz [32], with kind permission of Birkhäuser Verlag, Basel.)

It remains to be shown by MSP if the corner cones in the growth zones absorb UV light. However, in situ labeling of UVS cones confirms the putative identity of the disappearing corner cells. UVS opsin mRNA was localized to single cones with fully elaborated OS in the PGZ of O. mykiss [24].

The Developing Teleostean Eye

Over the years it has been maintained that the CNS in teleost embryos does not arise as an infolding but is solid with a cavity appearing only later (called ‘secondary neurulation’). However, due credit should be given to Goette 42

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[33, 34] who was the first to stress that the neural chord displays a ‘closed neural fold’ in the trout (fig. 17c). Some other early investigators reported the same observations but most were ignored until recent reports revealed, following the study of morphogenetic movements of single neural cells by labeling them with fluoresceinated dextran, that in the zebrafish neurulation also resembles the ‘primary neurulation’ of the other vertebrates (fig. 17e, f) [35, 36]. However, it differs from primary neurulation of other taxa in that neurocoele formation occurs secondarily. In teleosts, a lumen, as a result of cavitation, is observed in the future eye before it appears in the future brain (fig. 18). Equally, the anlage of the lens, induced by Kunz

SM

SM

Some CoC New rods missing added

SM NO CoC

All CoC RM All CoC

PE RL ONL OPL

Neuroblasts INL

IPL GCL R:C 1:1.5

R:C 1:1.5 to 1:1.25

R:C 1:1 to 1:0.75

R:C 1:2

Fig. 13. Diagrammatic representation of the developmental regions in the rim (PGZ) of the eye of a mature S. salar. Darkened patches and black arrows represent areas of rod development and their movements to the sites of former UVS corner cones of the square mosaic. Note changes of row mosaics (RM) into square mosaics (SM). C = Cone; CoC = corner cone; R = rod; RL = receptor layer. (From Kunz et al. [22], with kind permission of Elsevier.)

the double-walled eyecup, is also solid and acquires a lumen only later. The ophthalmic and retinal arteries enter the eye via the embryonic fissure (fig. 10). The inner layer of the eyecup differentiates into the sensory retina while the outer layer forms the RPE, which remains as a single layer. The cells of the PE are the only chromatophores not emanating from the neural crest. The rim (PGZ) of the eyecup becomes the edge of the future pupil. The retinal development of three species, the rainbow trout Salmo gairdneri, the guppy P. reticulata and the zebrafish Danio (Brachydanio) rerio is dealt with below. Development and Aging in the Eye of Teleost Fish

Developmental Sequence in the Retina of Salmonids and the Guppy

Retinal Pigment Epithelium, Visual Cell Layer and Outer Nuclear Layer At stage A (hatching) of the trout and stage 5 of the guppy, the RPE is largely differentiated and contains spherical granules. This stage is often referred to as ‘eyed ova’. The cell junctions display alternating zonulae adhaerentes and zonulae occludentes (fig. 19a). Invaginations of the basal membrane continuous with the endoplasmic reticulum (ER), pinocytotic (coated) vesicles and numerous mitochondria are indicative of the passage of Neuroembryol Aging 2006–07;4:31–60

43

Fig. 14. a Dying OS of a UVS cone in the

PGZ. Arrow points to OS; CE = central cone; DC = double cone. b Degenerating IS of a UVS cone in the PGZ. (From Kunz et al. [22], with kind permission of Elsevier.)

44

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Kunz

material. The pale oval nuclei lie with their long axis parallel to the plane of the RPE. Rods develop vitreally to cones. Their nuclei are smaller and more electron dense than those of the cones situated above them while the nuclei of the corner cones lie at the level of the rod nuclei (fig. 19b). The rod IS (RIS) and CIS contain abundant rough ER (RER), free ribosomes, an elaborate Golgi body, microtubules and lamellated bodies. The CIS stain more deeply than RIS and their mitochondria are elongated. CC of both are developed but lie at different levels. Similarly, the CIS project through the external limiting membrane (ELM) while the RIS lie vitreally to it (fig. 19b). Prospective long single and double cones can be distinguished, but other investigators maintain that double cones in fish appear much later than single cones. ELM is represented by zonulae adherentes between IS and Mueller cells (fig. 19b). At stage B of the trout and stage 6 of the guppy, photoreceptor nuclei become radially oriented and show marginal heterochromatin (fig. 19b). Myeloid bodies (parallel stacks of smooth flattened cisternae) are continuous with the RER (and in the guppy with lipid bodies). Besides the spherical granules, rod-shaped granules are now evident (fig. 19c). RPE microvilli interdigitate with the calycal processes of the CIS (fig. 19b). Microvilli, the future calycal processes of the IS, extend into the ventricular space and CC have enlarged into an accessory OS (AOS). The OS have developed disks. While in the prospective OS of the cones the infoldings persist, in the rods the infoldings remain at the base of a column of free-floating disks in packages of two (fig. 20a, b). Stage C of the trout is characterized by membranebound clusters of small melanin granules in the RPE (fig. 19d). The development of rods precedes that of cones (fig. 20c). The ROS display packages of 9 disks (fig. 21a). IS are divided into myoids (with Golgi body and ER) and ellipsoids (with accumulated mitochondria, elongated in rods and round in cones). Double cones display subsurface membranes, which begin to fuse into cisterns, along their apposing IS. At stage D of the trout and stage 7 of the guppy, the rod-shaped pigment granules are concentrated in the processes of the RPE (fig. 22). Phagosomes (shed photoreceptor OS) associated with myeloid bodies and RER are evident. They display a circadian rhythm with a sharp peak and subsequent fall after sunset (due to shedding of Development and Aging in the Eye of Teleost Fish

Fig. 15. Radial section of fissure showing dying nucleus, myoid, pedicle (arrow) of UVS and dying nucleus of horizontal cell. Hz = Horizontal cell; My = myoid; N = nucleus; Pc = pedicle. (From Kunz et al. [22], with kind permission of Elsevier.)

the cones) and a second peak after sunrise (shedding of the rods) as shown in the trout (figs. 6, 23). As a result of the retinomotor movements, which start at stage D (fig. 5), the junctions between the RPE cells become progressively restricted to the basal region. Neuroembryol Aging 2006–07;4:31–60

45

Fig. 16. Apoptosis of UVS nucleus. Note Mueller cells engulfing pyknotic nucleus in d. Mü = Mueller cell (note glycogen granules). (From Kunz et al. [22], with kind permission of Elsevier.)

Rods as well as all types of cones are differentiated and have attained a light-adapted position (fig. 22). The coneto-rod ratio in both species is 2:1, and the rod disks in the trout now number 800 and the infoldings in the cones 340. Scattered sparse large green-sensitive single cones in the mosaic position of double cones are observed 46

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(fig. 8). Occasional triple cones have also been spotted. Putative ellipsosomes (modified scleral mitochondria in the accessory member of the double cones) are observed (fig. 22). At stage E of the trout (swim-up stage) and stage 8 of the guppy, retinomotor movements of the pigment granKunz

Fig. 17. a–c Formation of the neural chord in trout embryo as a ‘closed fold’. a Transverse histological sections through the trunk regions of the trout embryo Neural keel (ax I) is evident. b Section of trout embryo older than a. The side plates of the neural keel have ‘shrunk’ and a pseudostratified structure of the neural keel (r) is evident. c Section of a trout embryo older than b. The formerly pseudostratified neural keel has acquired a lumen. ax I = Median keel of neural plate (axial plate); ax II = lateral parts of neural plate; ch = notochord; d = periderm; db = entoderm;

f = dorsal transient ‘dip’; g = future epidermis; ob = epidermis; r = neurochord; rr = lumen of neurochord; sp = continuation of neural chord (sensory plate); st = somite. (From Goette, 1873, 1878 [33, 34].) d–f Morphogenesis of the neural chord in the zebrafish. Diagrammatic transverse sections. d Neural plate, e Neural keel (closed neural fold), f Cylindrical neural rod (closed neural fold). (From Papan and Campos-Ortega [35], modified by Kimmel et al. [36], with kind permission of Wiley-Liss.)

ules have started. Only the rod-shaped granules are involved. They move vitread during the day and sclerad at night. The spherical granules remain at the base of the RPE cells, which in the trout now houses membranebound lipid droplets in contact with the nuclear membrane and/or myeloid bodies (fig. 24). In the guppy, lipid

droplets were already evident at stage 7 (fig. 22). Guppies have started to shed the tips of their rods and cones into the RPE (fig. 23a, b). Differentiation in the guppy retina has extended from the fundus to the dorsal and upper one third of the ventral region. The mosaic reveals that along the contact

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18

Fig. 18. Appearance of lumina in primordia of the eye and neural chord of the trout. a Transverse section of stage G (14 somites). b Transverse section of stage H (end of epiboly). d = diencephalon; cr = lens; e´ = ectoderm; e = epidermis; ms = mesoderm; n = nerve; p = periblast; vo = optical vesicle. (From Henneguy [36].) Fig. 19. a RPE of the newly hatched trout (stage A) displays at the base of its cells junctional complexes with alternating zonulae adhaerentes (ZA) and zonulae occludentes (ZO). !45,000. (From Schmitt and Kunz [38], with kind permission of S. Karger AG, Basel.) b EM showing radially oriented photoreceptor nuclei and budding CIS and RIS of the guppy at stage 6. Bar = 2 m. (From Kunz et al. [39], with kind permission of S. Karger AG, Basel.) c RPE of the guppy at stage 6 contains round and elongated pigment granules. Myeloid bodies are connected with the nuclear membrane and are continuous with RER. Bar = 1 m. (From Ennis and Kunz [40], with kind permission of Elsevier.) d Membrane-bound composite of small pigment granules in the RPE of a trout at stage C. !20,000. M = Mitochondrion; MB = myeloid body; N = nucleus in RPE; PG = pigment granule; RN = rod nucleus; SYN = synapse. (From Schmitt and Kunz [38], with kind permission of S. Karger AG, Basel.)

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19

Kunz

Fig. 20. a Formation of OS in a cone of guppy at stage 6. Arrow points to tubular formation of future disks. Bar = 0.4 m. b Formation of OS in a rod of guppy at stage 6. Some disks (D) are pinched off from the OS plasma membrane. Bar = 0.1 m. (From Kunz et al. [39], with kind permission of Springer, Berlin/Heidelberg.) c RPE and visual cell layer in a trout at stage C. COS are ensheathed by RPE and rods have extended sclerally. RIS and CIS are divided into ellipsoid (el) and myoid (my). AOS = Accessory outer segment; B = bleb, CF = connecting cilium; DC = double cone; M = mitochondrion; RN = rod nucleus; SER = smooth endoplasmatic reticulum; SS = subsurface membrane. Bar = 2 m. (From Schmitt and Kunz [38], with kind permission of S. Karger AG, Basel.)

zones of the IS of double cones and long single cones, fenestrated cisterns are evident; however, they are absent in the contact zones of double and short single cones (fig. 25). Additionally, the rod myoids display fenestrated subsurface cisterns in all contact zones (with double, long single and short single cones). At stage F of the trout and stage 9 of the guppy, the RPE processes now contact apical microvilli of Mueller cells. In both trout and guppy, a two-peaked pattern of shedding (phagosome count) is observed (fig. 6). Full retinomotor movements are now evident (fig. 5). The OS of the rods now contain packages of 22 free-floating disks and are indented by the accessory OS (incisure; fig. 21b). The CIS display lateral fins interlocking with microvilli of Mueller cells in the trout, whereas fins are rudimentary and restricted to the myoid region in the

guppy. Nuclei of cones, with the exception of those of short single (UVS) cones, protrude through the ELM. The short single cones in the guppy and both types of single cones in the trout do not undergo retinomotor movements. As mentioned above, the cones of the guppy change from a square into a row mosaic during dark adaptation. In the dark, the COS of the trout do not penetrate into the RPE but bend. At Stage 10 and following stages of the guppy, the eye of the prepartum guppy shows an absence of short (blue) cones in the lower two thirds of the ventral part; they are replaced by long single cones. The mosaic pattern reveals zig-zag arrangement of double cones (probably red/red) alternating with rows of exclusively long single cones (probably green).

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traversing Mueller fibers and bipolar cells. At stage E, the horizontal cells are seen to be connected by gap junctions, and in the vitreal part nuclei of bipolar, Mueller and amacrine cells can be differentiated (fig. 26a–d). IPL. The number and type of synapses increase from stage C onwards. At stage C, conventional synapses are established followed by bipolar synapses, as well as ribbon and branched synapses at stage D.

Ontogeny of the Lactate Dehydrogenase Pattern in the Eye of Trout and Guppy

Fig. 21. a At stage C of the trout the ROS disks are separated into

packages of 9 as a result of the invagination of the outer plasma membrane (arrows). Bar = 0.25 m. b At stage F the number of disks in OS packages has increased to 22 (arrows). Bar = 0.1 m. cp = Calycal process. (From Schmitt and Kunz [38], with kind permission of S. Karger AG, Basel.)

The glycolytic enzyme LDH (EC 1.1.1.27) is a multilocus enzyme and displays a tissue-specific pattern (fig. 7). Two loci have been observed in most vertebrates: the Alocus coding for the A4 enzyme predominantly expressed in anaerobic tissue, e.g. skeletal muscles, and the B-locus coding for the B4 enzyme occurring predominantly in aerobic tissue, e.g. heart muscles. In teleosts, a third locus, C, was found to be expressed in the liver and retina as two different isozymes, which are thought to be encoded by two different genes (fig. 7). The retina-specific pattern is expressed in the guppy at stage 7 and in the trout shortly after hatching (fig. 27). In the early postembryonic phase, the eye bands move faster than at the end of the postembryonic phase.

Development of the Zebrafish Eye and Retina

In the neonate and 5- and 14-day-old guppy, the ellipsosome during light and dark adaptation reveals cristae with a dark patch. Outer Plexiform Layer, Inner Nuclear Layer and Inner Plexiform Layer in the Trout OPL. At stage C, cone terminals (pedicles), rod terminals (spherules) and growth cones of bipolar cells are observed. At stage E cone terminals exhibit three synaptic ribbons and rod terminals up to five dendritic invaginations. Telodendrial contacts between rod and cone terminals are observed. Spinules (fingerlike processes) present only in the light between horizontal cell dendrites and synaptic ribbons of pedicles become evident. By stage F, the network of above contacts has dramatically increased (fig. 26e, f). INL. At stage A, the horizontal cells form two layers (external and internal) followed by the layer of presumptive bipolar cells and spindle-shaped neuroblasts. By stage B, the midregion has been differentiated revealing 50

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The expression and regulation of ath5, a zebrafish homolog of Drosophila atonal, is exclusively transcribed in the developing retina in a pattern closely linked to neuronal differentiation. It has been suggested that a series of inductive events, initiated from the prechordal plate and progressing from the optic stalks, regulates the spread of neuronal differentiation across the zebrafish retina [42]. The ocular anterior segment consists of the cornea, lens, iris, ciliary body and highly specialized tissue at the iridocorneal angle. This segment is critical for focusing and for the regulation of intraocular pressure. Its morphogenesis in the developing zebrafish has been described by Soules and Link [43]. Developmental stages of the zebrafish are expressed as numbers of somites and/or hours postfertilization (hpf). Morphogenesis of the eye commences at 11.5 hpf. At 13 hpf (8–9 somites), the eye anlage has taken up a wing-like shape. The ventricle within the forebrain and the optic Kunz

Fig. 22. RPE and visual cell layer of the guppy at stage 7 (fundic region). Rods and cones are differentiated. Bar = 3 m. DC = Double cone; E = ellipsosome; L = lipid body; LS = long single cone; MB = myeloid body; MU = Mueller cell; MY = myoid; N = cone nucleus; RN = rod nucleus; SC = short single cone; SSC = subsurface cisternae; SY = synapse; W = lamellated body. (From Kunz et al. [39], with kind permission of Springer-Verlag, Heidelberg.)

lumina arise separately and fuse by the 12-somite stage (15 hpf). From 14–15 hpf, the optic primordia bend ventrally and rotate slightly in anterior direction, independently of the early stage of the cephalic flexure. Invagination of the eye primordium commences at 15–16 hpf and the choroid fissure becomes apparent. It increases in depth as the eyecup invaginates further and is completely formed by 24 hpf. A further rotation of the eye in relation to the embryonic axis takes place between 24 and 36 hpf so that the choroid fissure is now shifted into a ventral position, as is typical for all other teleost embryos so far

analyzed. This movement occurs concomitantly with the late stages of cephalic flexure. At 36 hpf the first axons exit.

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Retinal Pigment Epithelium Black round granules are first observed at 32 hpf. Additionally, rod-shaped granules are distinguished at 50 hpf. At 60 hpf RPE processes begin to extend between the visual cells. Often miniature pigment granules, enclosed by a membrane, are observed (fig. 28 ; as described for the trout in fig. 19d). 51

Fig. 23. Shedding of tips of photoreceptor OS in the guppy at stage 8. a ROS shed during the day. Bar = 1 m. b COS shed at night. Bar = 1 m. C = Choroid gland; P = cone phagosome; RP = rod phagosome. (From Kunz

and Ennis [41], with kind permission of Elsevier.)

Visual Cell and Outer Nuclear Layer While in most teleosts differentiation of the retina begins in the fundic region, retinal development in the zebrafish starts at 32 hpf in the ventral retina, from the nasal region to the optic nerve (referred to as ‘ventronasal patch’). At 50 hpf EM revealed in the ventronasal patch and in the fundic region photoreceptor IS with RER and abundant free ribosomes, as well as microtubules extending from the basal body into the axoneme of the CC (fig. 29b and inset). At 54–55 hpf a small number of photoreceptor OS was observed. In the ventronasal patch, large oval putative cone nuclei are interspersed with small round putative rod nuclei. Contrary to the situation in the guppy and trout, there is no distinct separation of these two types of nuclei into two layers. The ‘adult type cone mosaic’ is already established at this stage. At 60 hpf EM disclosed distinct ROS and COS in the ventronasal patch. ROS measure 3 m in length (fig. 28). A separation into myoids and ellipsoids can be clearly detected. Instead of spreading temporad as rods do 52

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(fig. 30a–d), cones and ganglion cells spread nasally. Their number increases rapidly within the ventral region while they are observed only sporadically in the dorsal part. A topographical pattern of rod differentiation from 50 to 72 hpf within the nasal and temporal regions of the retina in embryonic whole mounts (obtained with Fret 114 staining) is described in fig. 30a–d. By 72 hpf – shortly before postembryos become free swimming and start feeding – the retina has become functional. By 74 hpf an ‘area’ (site of acute vision) has arisen in the ventrotemporal region. Rods are rarely found in this region and are not encountered in the fundic region until 9–10 days after fertilization (fig. 31a, b). Outer Plexiform Layer At 50 hpf this layer is less than 2 m wide in the central and ventral retina and not yet evident in the dorsal region. Photoreceptor terminals are not yet encountered. Putative processes of horizontal cells, identified by the presence of glycogen granules, are observed. At 62 hpf the first photoreceptor synaptic ribbons are evident in the Kunz

Fig. 24. Light-adapted trout retina at stage E. ROS reach the midline of the RPE. Inset = Accumulation of lipid droplets. Bar = 5 m. aSC = Corner cone (UVS); CN = cone nucleus; cSC = central single cone; DC = double cone; f = fins; L = lipid droplet; my = myeloid body; ss = subsurface membrane. (From Schmitt and Kunz, 1989 [38], with kind permission of S. Karger AG, Basel.) Fig. 25. Dorsal square mosaic pattern during the light phase in the guppy (late stage 8). Zig-zagging subsurface membranes (SSM) separating the partners of the double cones (DC), indicative of a square mosaic pattern. SSM are also observed in the contact zones between DC and long single cones (LS), DC and rods, and LS and rods. Bar = 1 m. Inset upper right: short single cones (SS), sectioned at the level of their OS, show an intercellular space (SP) surrounding it. SS in contact-zones of DC and LS are also evident. Inset upper left: diagram of square mosaics showing rows of alternating SS and LS and a zig-zag arrangement of DC. (Kunz and Ennis [41], with kind permission of Elsevier.)

24

25

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Fig. 26. TEM of the inner retina of the trout: stage A (hatching). a Differentiating bipolar cells sclerad to spindle-shaped neuro-

blasts. Pyknotic nuclei are surrounded by Mueller cell fibers. Bar = 5 m. b Differentiating amacrine cells, characterized by large electron-lucent nuclei. Perikarya contain mitochondria, RER, microtubules and a Golgi body. Bar = 5 m. Stage E (swimup stage). c Nuclei of bipolar, Mueller and amacrine cells (arrows) are easily distinguished. Dendrites of bipolar and amacrine cells and a broad Mueller fiber extend towards the IPL. Bar = 12 m. d Gap junctions (arrow) between two somata of external horizontal cells. Bar = 0.1 m. Stage F (external food uptake). e Electrondense zone (arrow) between two telodendria. !100,000. f Gap junctions (arrow) between two telodendria. !33,000. am = Amacrine cell; bp = bipolar cell; M = Mueller cell fiber; mn = Mueller cell nucleus; nb = neuroblast; pk = pyknotic nuclei; r = ribosome; sv = synaptic vesicle. (From Schmitt and Kunz [38], with kind permission of S. Karger AG, Basel.)

ventronasal patch. At 70–74 hpf photoreceptor synapses form triads (consisting of two lateral horizontal cell processes and a central bipolar cell process opposing the ribbon; fig. 32a). Inner Nuclear Layer At 50 hpf this layer is 25 m wide. Horizontal cells are small and irregularly shaped. In the midregion spindleshaped neuroblasts (presumptive bipolar cells) are observed. Lightly staining presumptive nuclei of amacrine cells are evident. The difference between these cells is most advanced in the ventronasal patch (fig. 29a). At 60 hpf the horizontal cell nuclei are elongated and lie parallel to the retinal surface. Bipolar cells are distinguished 54

Neuroembryol Aging 2006–07;4:31–60

in the midregion. The width of the INL has increased to 28–30 m. Inner Plexiform Layer At 50 hpf the layer includes putative displaced amacrine cells which will be later incorporated into the ganglion cell layer. At 60 hpf presumptive bipolar cell axons and terminals become distinguishable. A conventional synapse (small cluster of synaptic vesicles at the presynaptic membrane of an amacrine cell) is evident. Conventional synapses are now present both in the ventral and dorsal regions. At 70–74 hpf presumptive synaptic ribbons within a bipolar cell axon terminal, opposed by two postsynaptic processes, are shown (fig. 32b). Kunz

PE R/C

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Fig. 27. Development of LDH isozyme pattern in the guppy embryo. a Starch gel electrophoresis of the eye in a stage 7 embryo showing adult pattern (compare with fig. 7). b Histological section of a stage 7 embryo showing that the rods and all three types of cones are present and have assumed a light-adapted position. c Histological section of a stage 6 embryo showing that rods, cones and RPE are present but not yet fully differentiated. d Electropho-

resis reveals that eye bands are not yet present. Arrow indicates site of application of sample. ^ = Bands unique to the eye and brain; – = cathode; + = anode. IC = Short single cones; MC = long single cones; OC = double cones; OLM = outer limiting membrane; R = rods; R/C = rods and cones. (Kunz [13], with kind permission of the Editor of the Revue Suisse de Zoologie.)

Ganglion Cell Layer At 32 hpf a small number of ganglion cells is observed in the ventronasal patch. The narrow optic lumen in the stalk is continuous with the ventricle of the forebrain. By 40 hpf ganglion cells form a large patch within the nasal retina and send two layers temporad into the optic nerve.

At 50 hpf displaced amacrine cells have arrived in the ganglion cell layer. After the zebrafish has hatched, it attains vision just before the ‘swimming-up’ stage to start exogenous food uptake.

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Fig. 28. TEM of zebrafish eye showing devel-

oping IS and OS in the ventronasal patch at 60 hpf. Bar = 1.5 m. White arrows point to zonulae adherentes between Mueller cells and ellipsoids which form the outer limiting membrane. Black arrows point to pigment granules made up of small granules. Inset: IS is now divided into ellipsoid and myoid. Black arrows point to developing CC. e = Ellipsoid; M = Mueller cells; my = myoid; n = nucleus; RN = rod nucleus. (From Schmitt and Dowling, 1999 [44], with kind permission of WileyLiss.)

Fig. 29. Zebrafish eye at 50 hpf. a Magnified ventral part, to the right of optic nerve. Arrowheads point to pigment granules. Thin arrow indicates presumptive rod nuclei. Bar = 14 m. b Two types of pigment granules in the PE are shown as well as differentiation of photoreceptor IS with large mitochondria (asterisks) extending towards the PE into the interphotoreceptor space (IPS). Mueller cells project between cone nuclei and form zonulae adherentes (arrows) with the developing IS. Bar = 1 m. Inset: Microtubules

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(small arrows) extend from the basal body into the CC of the photoreceptor. Bar = 0.5 m. A = Amacrine cell; BB = basal body; CN = cone nucleus; G = ganglion cell; GCL = ganglion cell layer; H = horizontal cell; IS = photoreceptor IS; M = Mueller cell; NB = neuroblast; NE = undifferentiated neuroepithelial cells; ON = optic nerve. (From Schmitt and Dowling [44], with kind permission of Wiley-Liss.)

Kunz

Fig. 30. Topographical pattern of rod differentiation within the

nasal and temporal regions of the retina in whole mounts of zebrafish embryos (Fret 114 staining). a 50 hpf: initial rods along the nasal side of the choroids fissure (small arrows). b 57 hpf: increased number of rods (small arrows). Stained rods now extend temporally (asterisk). c 72 hpf: rods extend equally into both nasal and temporal regions (white arrows). d 72 hpf: rods densely ag-

Development of the Eye of Teleosts with Indirect Development

The bilaterally symmetrical larvae of the Atlantic halibut H. hippoglossus have one eye on each side of the body. During metamorphosis the larva changes to a juvenile adapted to a benthic lifestyle. This involves changes in the general body plan including the position of the Development and Aging in the Eye of Teleost Fish

gregated within the ventral patch (large arrow) and randomly scattered through nasal and temporal regions within the dorsal periphery (small arrows). D = Dorsal; N = nasal; R = rods; T = temporal; V = ventral; VP = ventral patch. Bar = 75 m (a–c) and 100 m (d). (From Schmitt and Dowling [45], with kind permission of Wiley-Liss.)

eyes. One eye migrates to the contralateral side of the head adjacent to the other eye. As a result the fish can now scan binocularly the surrounding above them [46]. The retina of the pelagic, surface-dwelling larva, at the phase of first feeding, consists of only one type of single cone cells, which are morphologically identical and packed in rows. Four different cone opsins – UVS, blue-, Neuroembryol Aging 2006–07;4:31–60

57

green- and red-sensitive cones – are expressed at the end of the yolk sac period (40 days after hatching) by in situ hybridization using RNA probes. UVS opsin is expressed only in the ventral retina. 90% of the visual cells express green-sensitive opsins, without any regional differences. In contrast, the remaining percentage of blue- and redsensitive opsins is distributed over the whole retina. No rod-opsin-expressing cells are found. However, they are present in the retina of the halibut after metamorphosis. It was hypothesized that the single cone retina of the juvenile halibut indicates some form of mosaic expression reminiscent of that found in halibuts after metamorphosi. This implies changes in opsin expression within individual cones. A new group of UVS cells appears in the

32 Fig. 31. Area (site of acute vision) in the ventrotemporal retina. a Sagittal section at the level of the lens 5 days after fertilization.

The boxed area is characterized by an increased ONL (small arrows), a tiering of cones (long arrow) and an absence of rods in the temporal area. Bar = 60 m. b Higher magnification of the boxed area in a. A tier of short single cones (SSC) lies vitread to a tier of longer single cones (LSC), which extend into the RPE (arrowheads). The ONL in this region is made up of two to three layers of tightly packed nuclei (small arrows) in contrast to the more dorsally placed nuclei (open arrows). Bar = 20 m. CF = Choroid fissure; D/V = dorso/ventral; NR = nasal retina. (From Schmitt and Dowling [44], with kind permission of Wiley-Liss.) Fig. 32. a Triad of photoreceptor synapse at 74 hpf: two lateral horizontal processes (1, 2) and a central bipolar process (3) opposing the ribbon. Synaptic vesicles (SV) and electron-dense postsynaptic zones (arrows) are observed. Bar = 120 nm. b At 74 hpf a presumptive synaptic ribbon (SR) within a bipolar cell axon terminal in the IPL is opposed by two postsynaptic processes (1, 2). Arrowheads point to synaptic vesicles and arrow to electrondense material associated with the postsynaptic membranes. Bar = 90 nm. (From Schmitt and Dowling [44], with kind permission of Wiley-Liss.)

31

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ventral retina during metamorphosis when the fish changes to a benthic form of life [46]. In the retina of the winter flounder Pleuronectes americanus only opsin for green cones was expressed and a corresponding middle wavelength visual pigment was observed before metamorphosis. After metamorphosis, rod opsin and three cone opsins were expressed, but cones were always produced before rods. However, the different cone types were produced simultaneously [47].

Discussion of the Appearance of Rod versus Cone Precursors

In teleosts with indirect development, the rods arise only after metamorphosis when the fish descend to a benthic habitat [46, 47]. This would explain why it has been

repeatedly stated that teleosts first develop cones and only later rods. However, as shown in this review, in fish with direct development (such as guppy and salmonids), both rod and cone precursors are present in the embryonic retina and differentiate first in the fundic region. Rods develop even faster than cones. Rod nuclei are smaller and usually positioned vitreally to the cone nuclei. Therefore, rods develop at a more vitreal level than cones and only extend sclerally when they are differentiated and in a light-adapted position. In the zebrafish embryo, early rod formation has been discovered in the ventronasal region of the retina [44, 45]. It has been suggested that retinoic acid regulates the differentiation of rods because the application of exogenous retinoic acid to zebrafish during the initial stages of photoreceptor differentiation resulted in a precocious development of rods and an inhibition of cones [48].

References 1 Kunz YW: Developmental Biology of Teleost Fishes. Dordrecht, Springer, 2004, chap 12, pp 239–250, and chap 17, pp 303–375. 2 Kunz YW: Development of the Eye of Teleosts; in Kapoor BG, Hara TJ (eds): Sensory Biology of Jawed Fishes. Enfield, Science Publishers, 2001, chap 1, pp 1–18. 3 Kunz YW: Ontogeny of retinal pigment epithelium: photoreceptor complex and development of rhythmic metabolism under ambient light conditions. Prog Retin Res 1990; 9:135–196. 4 Bielmaier O, Neuhauss SC, Kohler K: Onset and time course of apoptosis in the developing zebrafish retina. Cell Tissue Res 2001; 306:199–207. 5 Kishi S: Functional aging and gradual senescence in zebrafish. Ann NY Acad Sci 2004; 1019:521. 6 Genade T, Benedetti M, Terzibasi E, Roncaglia P, Valenzano DR, Cattaneo A, Cellerino A: Annual fishes of the genus Nothobranchius as a model system for aging research. Aging Cell 2005;4:223–233. 7 Yamamoto Y, Jeffery WR: Central role for the lens in cave fish eye degeneration. Science 2000;289:631–633. 8 Tabata M: Persistence of pineal photosensory function in blind cave fish, Astyanax mexicanus. Comp Biochem Physiol A 1982; 73: 125–127. 9 O’Brien P, Klein DC: Pineal and Retinal Relationships. New York, Academic Press, 1986, p 434. 10 Peirson SN, Bowmaker JK: The photopigment content of the goldfish pineal organ. Invest Ophthalmol Sci 1999;404:S158.

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11 Forsell J, Ekstrom IN, Flamarique IN, Holmquist B: Expression of pineal ultraviolet- und green-like opsins in the pineal organ and retina of teleosts. J Exp Biol 2001; 204: 2517–2525. 12 Ennis S, Kunz YW: Ageing of the collagenous layer in Bruch’s membrane of the teleost Poecilia reticulata. Cell Biol Int Rep 1984; 8:902. 13 Kunz YW: Distribution of lactate dehydrogenase (and its E-isozymes) in the developing and adult retina of the guppy (Lebistes reticulatus). Rev Suisse Zool 1971; 78: 761– 776. 14 von Frisch K: Ueber den Farbensinn der Fische. Verh Deutsch Zool Ges 1911, pp 20– 21, pp 220–225. 15 Hess C: Untersuchungen über den Lichtsinn bei Fischen. Arch Augenheilkde, Wiesbaden 1909;64(Suppl.):1–38. 16 Hess C: Ueber den angeblichen Nachweis von Farbensinn bei Fischen. Arch Ges Physiol 1910;134:1–14. 17 Bowmaker JK, Kunz YW: Ultraviolet receptors, tetrachromatic colour vision and retinal mosaics in the brown trout (Salmo trutta): age-dependent changes. Vision Res 1987; 217:2102–2108. 18 Kunz YW, Wise C: Ultrastructure of the ‘oildroplet’ in the retinal twin-cone of Lebistes reticulatus (Peters). Preliminary results. Rev Suisse Zool 1973;80:694–698. 19 Tohya S, Mochizuki A, Iwasa I: Difference in the retinal cone mosaic pattern between zebrafish and medaka: cell-rearrangement model J Theor Biol 2003; 221:289–300.

20 McCormack CA, Hayden TH, Kunz YW: Ontogenesis of diurnal rhythms of cAMP concentration, outer segment disc shedding and retinomotor movements in the eye of the brown trout Salmo trutta. Brain Behav Evol 1989;34:65–72. 21 Fürst CM: Zur Kenntnis der Histogenese und des Wachstums der Retina. Acta Universita Lund, 1892, pp 1–45. 22 Kunz YW, Wildenburg G, Goodrich L, Callaghan E: The fate of ultraviolet receptors in the retina of the Atlantic salmon (Salmo salar). Vision Res 1994; 34: 1375– 1383. 23 Allison WT, Dann SG, Veldhoen KM, Hawryshyn CW: Degeneration and regeneration of ultraviolet cone receptors during development in rainbow trout. J Comp Neurol 2006;499:702–715. 24 Allison WT, Dann SG, Helvik JV, Bradley C, Moyer HD, Hawryshyn CW: Ontogeny of ultraviolet-sensitive cones in the retina of rainbow trout (Oncorhynchus mykiss). J Comp Neurol 2003;461:294–306. 25 Zygar CA, Colbert S, Yang D, Fernald RD: IGF-1 produced by cone photoreceptors regulates rod progenitor proliferation in the teleost retina. Brain Res Dev Brain Res 2005; 154:91–100. 26 Dann SG, Allison WT, Levin DB, Hawryshyn CW: Identification of a unique transcript down-regulated in the retina of rainbow trout (Oncorhynchus mykiss) at smoltification. Comp Biochem Physiol B Biochem Mol Biol 2003; 136:849–860.

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27 Deutschlander ME, Greaves DK, Haimberger TJ, Hawryshyn CW: Functional mapping of ultraviolet photosensitivity during metamorphic transitions in a salmonid fish, Oncorhynchus mykiss. J Exp Biol 2001; 204: 2401–2413. 28 Hawryshyn CW, Martens G, Allison WT, Anholt BR: Regeneration of ultraviolet-sensitive cones in the retinal cone mosaic of thyroxin-challenged post-juvenile rainbow trout (Oncorhynchus mykiss). J Exp Biol 2003;2006:2665–2673. 29 Flamarique IN: The ontogeny of ultraviolet sensitivity, cone disappearance and regeneration in the sockeye salmon Oncorhynchus nerka. J Exp Biol 2000;203:1161–1172. 30 Losey GS, Cronin TW, Goldsmith TH, Hydes D, Marshall NJ, McFarland WN: The UV visual world of fishes: a review. J Fish Biol 1999; 54:921–943. 31 Portmann A: Einführung in die vergleichende Morphologie der Wirbeltiere, ed 6. Basel, Schwabe, 1983. 32 Kunz YW: Tracts of putative ultraviolet receptors in the retina of the two-year-old brown trout (Salmo trutta) and the Atlantic salmon (Salmo salar). Experientia 1987; 43: 1202–1204. 33 Goette A: Beiträge zur Entwicklungsgeschichte der Wirbelthiere. I. Der Keim des Forelleneies. Arch Mikr Anat 1873; 9: 679– 708.

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34 Goette A: Beiträge zur Entwicklungsgeschichte der Wirbelthiere. III. Ueber die Entwicklung des Zentral-Nervensystems der Teleostier. Arch Mikr Anat 1878;5:139–157. 35 Papan C, Campos-Ortega A: On the formation of the neural keel and neural tube in the zebrafish Danio (Brachydanio) rerio. Rouxs Arch Dev Biol 1994; 203:178–186. 36 Kimmel CB, Ballard WW, Kimmel SR, Ullmann B, Schilling TF: Stages of embryonic development of the zebrafish. Dev Dyn 1995; 203:253–310. 37 Henneguy LF: Recherches sur le développement des Poissons osseux. Embryogénie de la Truite. J Anat Physiol Paris 1888;24: 413– 617. 38 Schmitt E, Kunz YW: Retinal morphogenesis in the rainbow trout, Salmo gairdneri. Brain Behav Evol 1989;34:48–64. 39 Kunz YW, Ennis S, Wise C: Ontogeny of the photoreceptors in the embryonic retina of the viviparous guppy, Poecilia reticulata P. (Teleostei). An electron-microscopical study. Cell Tissue Res 1983;230:469–486. 40 Ennis S, Kunz YW: Myeloid bodies in the pigment epithelium of a teleost embryo, the viviparous Poecilia reticulata. Cell Biol Int Rep 1984;8:1009–1011. 41 Kunz YW, Ennis S: Ultrastructural diurnal changes of the retinal photoreceptors in the embryo of the viviparous teleost (Poecilia reticulata P.). Cell Differ 1983;13:115–123.

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42 Masai I, Stemple DL, Okamoto H, Wilson WS: Midline signals regulate retinal neurogenesis in zebrafish. Neuron 2000; 24: 251– 263. 43 Soules KA, Link BA: Morphogenesis of the anterior segment in the zebrafish eye. BMC Dev Biol 2005;5:12. 44 Schmitt E, Dowling JE: Early retinal development in the zebrafish Danio rerio : a light and electron microscopical analysis. J Comp Neurol 1999;404:515–536. 45 Schmitt E, Dowling JE: Comparison of topographical patterns of ganglion and photoreceptor cell differentiation in the retina of the zebrafish, Danio rerio. J Comp Neurol 1996; 371:222–234. 46 Helvik JV, Drivenes O, Torstein H, Seo HC: Topography of different photoreceptor cell types in the larval retina of Atlantic halibut (Hippoglossus hippoglossus). J Exp Biol 2001; 204:2553–2559. 47 Mader MM, Cameron DA: Photoreceptor differentiation during retinal development, growth and regeneration in a metamorphic vertebrate. J Neurosci 2004; 24: 11463– 11472. 48 Hyatt GA, Schmitt EA, Fadool JJ, Dowling JE: Retinoic acid alters photoreceptor development in vivo. Proc Natl Acad Sci USA 1996;93:13298–13303.

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Neuroembryol Aging 2006–07;4:61–75 DOI: 10.1159/000103877

Published online: June 13, 2007

The Pathology of Mueller Cells under Different Disease Conditions Shimpei Nishikawa Department of Ophthalmology, National Defense Medical College, Tokorozawa, Japan

Key Words Fovea development
Mueller cells
Preconditioning

Abstract The morphology and enzymes of Mueller cells were investigated in the developing retinas of Royal College of Surgeons (RCS) rats. Immunohistochemistry showed glutamine synthetase (GS) expression first on P10. It increased later in both normal and dystrophic retinas. Glial fibrillary acidic protein was not expressed in normal retinas, but Mueller cells of dystrophic retinas were stained on P35 and P100. The percentages of the area of Mueller cells in the outer nuclear layer in the macula, posterior pole, equator and periphery were 12.3 8 2.7, 22.1 8 4.5, 23.6 8 1 and 26.7 8 4.5%, respectively. In the foveal region, lower levels of GS and glutamate/aspartate transporter (GLAST) immunoreactivity were confirmed. Enzyme-linked immunosorbent assay and Western blot analyses revealed that the amounts of GS and GLAST in the foveal region were smaller than those in any other region. The role of Mueller cells in the preconditioned retinal ischemic rat was also investigated. Three and 7 days after ischemia, GS immunoreactivity recovered only in the preconditioned group. After 1 h of ischemia, the retinas showed increased GFAP immunoreactivity in the non-preconditioned group at 3 and 7 days. In the preconditioned group, only slight GFAP immunoreactivity was observed. These results suggest that the mechanism of preconditioned retinal ischemia may be related to Mueller cells in the retina. Copyright © 2007 S. Karger AG, Basel

© 2007 S. Karger AG, Basel 1661–3406/06/0042–0061$23.50/0 Fax +41 61 306 12 34 E-Mail [email protected] www.karger.com

Accessible online at: www.karger.com/nba

Mueller cells are giant cells that occupy the full thickness of the retina from the internal to the external limiting membrane and beyond as villous processes. They are recognized by their cytoplasmic density which is greater than that of the adjacent neurons. These cells are important because they furnish glucose to the nerve cells and are able to synthesize and store glycogen [1]. Their extensions enclose most of the neurons and insulate the nerve fibers. Mueller cells can also be considered space-occupying cells since their extensions fill all the retinal space not occupied by neurons, except in the inner retina, where astrocytes also play this role.

Cell Generation

Ganglion cells, horizontal cells and cones are always the first born cohort. They first develop in the fovea on fetal days 33–38 (8 weeks) and later in the periphery on fetal days 90–100 (24 weeks). This means that no new cones are generated after midgestation. Amacrines form a middle cohort and are generated slightly later. The last cohort includes bipolar cells, rods and probably Mueller glial cells. Mueller cells arise, together with rod photoreceptor cells, bipolar cells and a subset of amacrine cells, from common precursor cells during a late proliferative phase. One Mueller cell and a species-specific number of such neurons seem to form a columnar unit within the retinal tissue. In contrast, extracolumnar neurons (ganglion

Shimpei Nishikawa Department of Ophthalmology National Defense Medical College, 3-2 Namiki Tokorozawa, Saitama 359-8513 (Japan) Tel. +81 4 2995 1683, Fax +81 4 2993 5332, E-Mail [email protected]

cells, cone photoreceptor cells, horizontal cells and another subset of amacrine cells) develop and begin to differentiate before most Mueller cells are generated. It may be essential for such neurons to develop metabolic capacities sufficient to support their own survival, whereas late-appearing (columnar) neurons seem to depend on a nursing function of their Mueller cells. Thus, of all the cell types within a retinal column, it is exclusively Mueller cells that possess the enzyme for glycogen metabolism (fig. 1) [2].

Mueller Cells in Developing Rats with Inherited Retinal Dystrophy [3]

Riepe and Norenburg [4] first localized glutamine synthetase (GS), a key enzyme of the glutamate-glutamine cycle, in Mueller cells in rat retina by immunohistochemical techniques. Since then, many studies have examined the morphology and function of Mueller cells by focusing on GS. Mueller cells are thought to play an important role both structurally and functionally in the retina, and they have a close relationship with the neuronal environment. Furthermore, it appears that they are involved in the pathology and repair of various diseases. Mueller cells express glial fibrillary acidic protein (GFAP) when the retina is damaged by certain factors or is in a pathological condition, including inherited retinal dystrophy [5–8] and other cases of retinal degeneration, injury, or light damage [5, 9, 10]. GFAP appears to be a

Fig. 1. Semi-schematic drawing of a co-

lumnar unit and some cell types (G, H, C, A) not belonging to the units. A, A = Amacrine cell; B = bipolar cell; M = Mueller cell; R = rod photoreceptor cell; G = ganglion cell; H = horizontal cell; C = cone photoreceptor cell. The amacrine cell on the left is underlined by hatching to indicate that only a special population of amacrine cells is extracolumnar, whereas a different population of amacrine cells is columnar. Figure adapted from [2]. Normal

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ing): it is remarkable that from P21 onward, the outer nuclear layer becomes disarranged (degenerative) in dystrophic retina and its thickness decreases.

62

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glutamine and arginine is elevated in Mueller cells prior to the onset of photoreceptor death, due to anomalies in glutamate degradation [14]. After being taken up by Mueller cells, glutamate is intracellularly converted into glutamine by the enzyme GS, which is only expressed in Mueller cells [15] and which operates in concert with glutamate/aspartate transporter (GLAST) [16]. Likewise, after the uptake of
-aminobutyric acid (GABA) into Mueller cells, it is converted to glutamate [17], which then is fed into the GS pathway. Glutamine is transported back to neurons as a precursor for their synthesis of glutamate and GABA (glutamate-glutamine cycle of the retina) [18]. Due to the efficiency of the GS, demonstrable levels of glutamate are only present in Mueller cells when the GS is experimentally inhibited [19] or downregulated under pathological conditions [20]. When the GS is experimentally blocked in Mueller cells, retinal neurons lose their glutamate content, and the animals rapidly (within 2 min) become functionally blind [19, 21]. From postnatal day 21 (P21) onward, the outer nuclear layer (ONL) of the dystrophic retina becomes disarranged (degenerative) and its thickness decreases with development (fig. 2). Immunohistochemical Staining GS was first expressed in Mueller cells of both normal and dystrophic rats on P10, and staining increased afterwards (fig. 3a). In normal rats, GFAP was not expressed throughout the examined developmental stages. In dystrophic retinas, however, Mueller cells were stained with anti-GFAP antibody on P35 and P100. GFAP staining was hardly observed in Mueller cells on P1, P4, P10 and P21 (fig. 3b).

normal and dystrophic retinas. No significant differences were observed between both animals. b No GFAP expression was seen in normal retinas. Mueller cells of dystrophic retinas were immunostained on P35 and P100. Scale bar = 50 m.

Immunoblots of Extracts In both dystrophic and normal retinas, GS immunoblots were not recognized on P1 and P10, but a single band with similar densities of approximately 44 kDa was found on P21, P35 and P100 in these animals (fig. 4a). GFAP expression by Western blots showed a single band of about 50 kDa on P35 and P100 dystrophic retinas, and the intensity on P100 was stronger than that on P35. There was hardly any GFAP expression in rats examined at other stages (fig. 4b).

pathological marker of Mueller cells. Royal College of Surgeons (RCS) rats have autosomal recessive retinal dystrophy that results in progressive degeneration of (rod) photoreceptor cells [11–13]. In RCS rats, the content of

Developmental Profiles of GS and GFAP Determined by ELISA GS concentrations in both dystrophic and normal rat retinas increased according to development, but no remarkable difference was found between dystrophic and

Mueller Cell Pathology and Disease

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Fig. 5. Developmental profiles of GS and GFAP in rat retinas determined by ELISA. ) = Normal; $ = RCS. a GS concentrations

per protein increased with development in both normal and dystrophic retinas, although the difference between them was not significant. b In dystrophic retinas, GFAP concentrations per protein increased remarkably on P35 and P100. They were constantly at a low level in normal retinas throughout the developmental stages. Fig. 4. Immunoblots of extracts of rat retinas with GS and GFAP. a GS: a single band close to 44 kDa on P21, P35 and P100 can be

observed in both normal and dystrophic extracts of retinas. Both groups showed similar densities. b GFAP: the extracts of dystrophic retinas showed a single band close to 50 kDa only on P35 and P100. The density was greater on P100. + = Normal rat; – = dystrophic rat.

control retinas at any developmental stage (fig. 5a). GFAP concentrations in both dystrophic and normal rat retinas were similar to those on P21. In normal retinas, the concentrations were constant afterwards, whereas in dystrophic retinas they remarkably increased on P35 and P100 (p ! 0.05; fig. 5b). Data are shown as means 8 SD; p ! 0.05 was considered to be statistically significant (Student’s t test). The development of Mueller cells has been investigated using various animal models, including the rat [22], mouse [23] and rabbit [24, 25]. In normal rat retinas, GS, 64

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a specific enzyme of Mueller cells [4, 26, 27], was first detected on P5 by immunohistochemical techniques [22]. In our study, its expression was observed on P10, and no obvious staining was detected on P4. It was speculated that GS activities emerge within 1 week after birth with the development of Mueller cells. There was no significant difference between GS-specific activity per milligram protein of normal rats and that of dystrophic ones. We believe that GS converts the neurotransmitter glutamate into glutamine, which is then taken up by photoreceptor cells, synthesized into glutamate and released. Consequently, photoreceptor cells are the major source of glutamate in the retina. Even with progressive degeneration of photoreceptor cells, the function of Mueller cells does not seem to change. However, there is another possibility. It is well known that the thickness and total protein of the dystrophic rat retina Nishikawa

decrease with age [23]. Although there was no significant difference between GS-specific activity per milligram protein of two species, taking into account the total amount of GS, our observation might indicate a significant difference, and dystrophic retinas might display a rapid loss of GS. GS activity, the Mueller cells themselves, or the glioneuronal interactions were downregulated [13]. Columnar arrangements of cells in the developing retina can be observed by scanning electron microscopy. In the adult retina, each Mueller cell ensheaths a columnar group of neuronal cells. These cells consist of 1 Mueller cell, 11 rod photoreceptor cells, about 2 bipolar cells and 1–2 amacrine cells. It is thought that this unit arises from the migration of groups of preneurons along a common Mueller (precursor) cell, and that such columns might constitute metabolic and functional units. It is known that rod photoreceptor cell degeneration in RCS rats induces degenerative changes in secondary neurons of the inner retina [11, 13, 28]. Also, in RCS rats, Mueller cells are shorter than in normal rats, and their outer processes are hypertrophic [29]. GFAP is thought to be expressed due to gliosis and hypertrophy of macroglial cells [13, 30]. Moreover, usually GFAP is thought to be expressed in astrocytes in the retina, and in Mueller cells under pathological or stressful conditions [13]. The present results showed that immunoreactivity of GFAP and its content increased in dystrophic retinas at late developmental stages. Taken together, the following possibility is suggested. As the degeneration of rod photoreceptor progresses, other neurons might become degenerative, and the neuronal relationship between Mueller cells and neuronal cells might be damaged within the columnar unit. Also, Mueller cells might reactively become gliotic and hypertrophic to compensate for the structural defect.

Aging of Mueller Cells of the Fovea [31]

Most frequently, the presence of macular edema is responsible for the decreased vision in patients with diabetic retinopathy [32, 33]. By compression of retinal neurons, nerve fibers and capillaries, edema contributes to photoreceptor degeneration and neuronal cell death, and exacerbates the ischemic conditions. The development of chronic edema depends on two parameters: the rate of fluid entry into the retinal parenchyma through leaky vessel walls and the rate of fluid reabsorption from the retinal tissue back into the blood. Fluid absorption is carried out by the pigment epithelium (which absorbs the Mueller Cell Pathology and Disease

fluid from the subretinal space) and by Mueller cells (which absorb the fluid from the inner retinal tissue by transcellular water transport) [34]. Macular diseases, including age-related macular degeneration (AMD), cystoid macular edema and macular holes, have recently received attention. The cardinal symptom is blurring of central vision. However, the primary pathogenic basis of AMD remains unclear. One of the reasons may be the poor understanding of the structural integrity of the fovea. The specialized structure in the macular region accounts for its tendency to be affected by certain disease processes. Our present knowledge on the morphology of the fovea centralis in the human retina is derived from the works of many investigators dating from the early 19th century. A historical survey and explanation of the foveal structure can be found in reviews by Kolmer [35], Polyak [36], Duke-Elder and Wybar [37] and Rohen [38]. We know that the foveola forms the central floor of the fovea and has a diameter of 0.35 mm. It is the thinnest part of the retina and is devoid of ganglion cells. Its entire thickness consists only of cones and their nuclei, and it subserves the most acute vision. The umbo is a tiny depression in the very center of the foveola that corresponds to the ophthalmoscopically visible foveolar reflex seen in most healthy eyes. The fovea is a zone 1.5 mm in diameter, and the parafoveal central retina, which surrounds the fovea, is about 0.5 mm in width. According to Yamada [39], a relatively large number of Mueller cell processes is found at the center of the fovea. These cells constitute the floor of the fundus and possess a watery cytoplasm among central cone fibers in an area about 50 m in diameter. The mitochondria of the foveal cone tend to be dispersed in the inner segment and outer cone fiber. At the center, no ellipsoid is discernible in the inner segment of the foveal cone. The basal lamina along the vitreal surface is extremely thin at the fundus of the fovea. The center of the fovea externa, about 80 m in diameter, shows a slight convex elevation. GS and GLAST in the foveal region were investigated by ELISA, Western blot analysis and immunocytochemistry techniques. Data are shown as means 8 SD; p ! 0.05 was considered to be statistically significant (Student’s t test). Conventional Electron Microscopy The ultrastructure of the human retina in the ONL (fig. 6) consists of (a) the macula, (b) the posterior pole, (c) the equator and (d) the periphery. The ONL contains the cell bodies of the rods and cones with their nuclei and Neuroembryol Aging 2006–07;4:61–75

65

Fig. 6. Ultrastructure of the human retina in the ONL. a Macula: many neurotubules (arrowhead) are aggregated around the rods and cone nuclei, which occupy much of the cell body. Posterior pole (b), equator (c) and periphery (d) are shown. The longer the distance from the macula, the wider the diameter of the Mueller cell fiber. The cytoplasm contains numerous well-developed fibrils measuring 100 Å in diameter, which are randomly oriented (arrow). The Mueller cells fill all the space between the processes of the rods and cones. M = Mueller cell. Bar = 5 m.

cytoplasm. Mueller cell processes are also seen. The greater the distance from the macula, the wider the diameter of the Mueller cell fiber. The cytoplasm is darker than that of the surrounding cells and contains numerous well-developed fibrils measuring 100 Å in diameter. These fibrils are randomly oriented. The Mueller cells fill all the space between the processes of the rods and cones. On the other hand, many neurotubules are aggregated around the rods and cone nuclei, which occupy much of the cell body (fig. 6). The percentages of the area of Mueller cells in the ONL in the macula, posterior pole, equator and periphery were 12.3 8 2.7, 22.1 8 4.5, 23.6 8 1 and 26.7 8 4.5%, respectively (fig. 7). There was a significant difference between the macula and each of the posterior 66

Neuroembryol Aging 2006–07;4:61–75

pole, the equator and the periphery (p ! 0.05). The mean density of Mueller cells was about 25,000, 15,000, 12,000 and 9,000 cells/mm2, respectively. Immunohistochemical Staining At the foveola, GS staining was faint compared with that of the parafoveal region (low magnification: fig. 8a). The long transversal course of the outer trunks of Mueller cells in the outer plexiform layer (OPL) followed the course of the fibers of Henle. The Mueller cell end-feet or astrocytes were stained except at the foveola (high magnification: fig. 8b). At the foveola, GLAST staining was faint compared with that of the parafoveal region (low magnification: Nishikawa

fig. 9a). The long transversal course of the outer trunks of Mueller cells in the OPL followed the course of the fibers of Henle. The stained outer processes of Mueller cells form the clear-cut outer limiting membrane, which also shows labeled microvilli extending in between the pho-

35

*

30

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25

*

20 15 1.6

10 5 0 Macula

Posterior pole

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Equator

Fig. 7. The areas of Mueller cells in the ten photographs for each region, i.e. 4 ! 4 ! 10 areas, were traced with the computer mouse [four human eyes, ten visual fields for each region (macula, posterior pole, equator and periphery)]. Cell areas were calculated directly based on the tracing. The percentages of the area of Mueller cells in ONL in the macula, posterior pole, equator and periphery were 12.3 8 2.7, 22.1 8 4.5, 23.6 8 1 and 26.7 8 4.5%, respectively. There was a significant difference between the macula and each of the posterior pole, the equator and the periphery (* p ! 0.05 vs. macula) by Tukey test.

toreceptor inner segments. Although the Mueller cell end feet were stained, there was only faint staining at the foveola (high magnification: fig. 9b). A photograph of the control experiment section, where the primary antibody was omitted, is shown in figure 9c. Immunoblots of Extracts of Human Retinas with GS Extracts from the macula, posterior pole, equator and periphery demonstrated a single band at the same position, which corresponded to 44 kDa for GS (fig. 10a). In the macula, the extracts yielded a slightly positive band for GS. The bands at the posterior pole, equator, and periphery were strong for GS. These results correlated with the results of immunohistochemical staining. Regional Profiles of GS and GLAST in Human Retinas The regional differences of GS concentration in the human retina were determined at the posterior pole, equator and periphery (fig. 11a). The optical absorbency at the macula, posterior pole, equator, and periphery were 0.20 8 0.03, 0.44 8 0.06, 0.45 8 0.07 and 0.49 8 0.05, respectively. Low GS concentrations were observed in the macula. The results obtained from ELISA agreed with those obtained from Western blot analysis. There was a significant difference between the macula and each of the posterior pole, the equator and the periphery (GS: p ! 0.05). Regional differences in the GLAST concentration in the human retina were determined at the posterior pole,

a

Fig. 8. GS immunoreactivity in the macula. a Low magnification: GS staining of the

foveola was faint compared with that of the parafoveal region. b High magnification: the long transversal course of the outer trunks of Mueller cells in the OPL followed the course of the fibers of Henle. The Mueller cell end-feet or astrocytes are stained except at the foveola. ILM = Inner limiting membrane; GCL = ganglion cell layer; IPL = inner plexiform layer; INL = inner nuclear layer; IS = inner segment. Bar = 100 m.

Mueller Cell Pathology and Disease

ILM GCL IPL INL OPL ONL IS

b

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67

Fig. 9. GLAST immunoreactivity in the macula. a Low magnification: GLAST

staining of the foveola was faint compared with that of the parafoveal region. b High magnification: the long transversal course of Mueller cells in the OPL followed the course of the fibers of Henle. The stained outer processes of Mueller cells form the outer limiting membrane. The Mueller cell end-feet were stained, although there is only faint staining at the foveola. c The control experiment section where the primary antibody was omitted is shown. Immunoreactivity was rarely observed. Bar = 100 m.

Macula

Posterior pole

Equator

Periphery

Fig. 10. Immunoblots of extracts of human retinas with GS. Characterization of mouse anti-GS antibody by Western blot and immunostaining techniques. Extracts from the macula, posterior pole, equator and periphery gave a single band at the same position, which corresponded to 44 kDa for GS. In the macula, the extracts yielded a slightly positive band for GS. The bands at the posterior pole, equator and periphery were strong for GS.

equator and periphery (fig. 11b). The optical absorbency at the macula, posterior pole, equator and periphery were 0.16 8 0.03, 0.24 8 0.05, 0.24 8 0.03 and 0.26 8 0.05, respectively. Low GLAST concentrations were observed 68

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in the macula. There was a significant difference between the macula and each of the posterior pole, the equator and the periphery (GLAST: p ! 0.05). Mueller cells play a central role in the homeostasis of glutamate in the retina because they possess two functionally linked properties [40], namely their capacity to take up glutamate from the extracellular space after its release by neurons [41] via the transporter GLAST [42] and their capacity to convert glutamate into glutamine using the enzyme glutamine synthetase [43, 44] for reexport to neurons, which then convert such glutamine back into glutamate [41]. At least four related but distinct eukaryotic glutamate transporters have been cloned to date: the rat brain glutamate transporter 1 (GLT-1) [45], the rat brain L-glutamate/L-aspartate transporter 1 (GLAST-1) [46], the excitatory amino acid carrier 1 (EAAC-1) [47] from rabbit intestine and the excitatory amino acid transporter 4 from the human brain (EAAT4) [48]. According to Rauen et al. [42], GLAST-1 immunoreactivity was seen in Mueller cells and astrocytes. EAAC-1 was found in horizontal cells, in amacrine and displaced amacrine cells, and in ganglion cells. GLT-1 was preferentially expressed by different types of bipolar cells. Derouiche and Rauen [49] demonstrated that the light- and electron-microscopic patterns of immunoreactivity of GS and GLAST were very similar, particularly in Nishikawa

the OPL, and they suggested that GS and GLAST may operate in concert to terminate the neurotransmitter action of glutamate. The modification in the general pattern of the neural elements in the retinal organization at the fovea has been known since early light-microscopic studies. For example, in 1866, Schultze [50] described the fovea, including the precise morphology of the foveal cone. In 1968, Wolff [51] demonstrated that there were no Mueller cells in the human fovea and reported that it consisted only of cone cells. On the other hand, in 1969, Yamada [39] showed that a relatively large number of Mueller cell processes resided at the center of the human fovea (Mueller cell cone). However, we observed that Mueller cell density is relatively

low. Mueller cell cones contain watery cytoplasm in contrast to the darker cytoplasm of ordinary Mueller cells. The function of the Mueller cell cones is unclear. Gass [52] hypothesized that it is a reservoir for retinal xanthophyll, the primary structural support for the fovea or a primary role player in age-related macular hole formation, and that congenital abnormalities affecting the Mueller cell cone are responsible for the pathognomonic biomicroscopic picture of foveomacular schisis. Some regional differences have also been reported recently. Distler and Dreher [29] showed that the thickness of the inner trunks of monkey Mueller cells increased significantly with increasing retinal eccentricity. According to the overall thickness of the retina, Mueller cells in the central retina were found to be longer than those in peripheral regions. In the parafoveal region, the outer trunks of Mueller cells in the OPL were immensely elongated. These Mueller fibers can reach lengths of several hundred micrometers as they travel through the OPL from the foveal center toward the foveal border where they enter the inner nuclear layer. In 1999, Burris et al. [53] reported a distribution of one Mueller cell per foveal cone in the macaque monkey. In this study, the percentages of the area of Mueller cells relative to the whole area in the ONL were 12.3 8 2.7, 22.1 8 4.5, 23.6 8 1 and 26.7 8 4.5% for the macula, posterior pole, equator and periphery, respectively. At least in the ONL, Mueller cells appeared to be burdened in the foveal region. Less GS and GLAST immunoreactivity was confirmed in the foveal region. ELISA and Western blot analysis revealed that the amounts of GS and GLAST were smaller in the macula than in any other region. It is thought that these histological and biochemical results were correlated and that the low density of Mueller cells in the fovea may make the area architecturally weak. Studies on other animals have shown that the Mueller cell volume fraction is higher in animals with avascular retinas than in animals with vascularized retinas, which may be an indication of the role of Mueller cells in retinal nutrition [54, 55]. This also supports the finding of a low volume fraction of Mueller cells in the central fovea where photoreceptors can be supplied entirely by diffusion from the choriocapillaris. AMD is one of the leading causes of blindness in Western countries. The primary pathogenic basis of AMD remains unclear, and although many studies have focused on the changes that occur in the retinal pigment epithelium and choroid [56], less consideration has been given to changes occurring in the neural retina. On the other hand, anti-astrocytes [57] and other anti-retinal autoantibodies [58] have

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Optical absorbency

0.6 0.5

*

*

*

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*

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0.4 0.3 0.2 0.1 0

a

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0.35

Optical absorbency

0.30

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0.25 0.20 0.15 0.10 0.05 0

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Fig. 11. a The regional profile of GS in human retinas was deter-

mined by ELISA. A regional difference in GS concentration was observed in the human retina. A low GS concentration was seen in the macula. Means 8 SD of three samples are shown. There was a significant difference between the macula and each of the posterior pole, equator and periphery (* p ! 0.05). b The regional profile of GLAST in human retinas was determined by ELISA (fig. 6b). A regional difference in GLAST concentration was observed in the human retina. A low GLAST concentration was seen in the macula. Means 8 SD of four samples are shown. There was a significant difference macula and each of the posterior pole, equator, and periphery (* p ! 0.05 vs. macula).

69

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Fig. 12. Representative ERG 3 days after 1 h of ischemia with and

without preconditioning. Preconditioning performed 24 h before ischemia significantly improved the recovery of a- and b-waves 3 days after 1 h of ischemia.

400 300 200 100 0 Pre

been detected in the sera of patients with AMD, suggesting that changes in the neural retina may be an early feature of the pathogenesis of AMD. These data may be relevant to the pathogenesis of AMD. In any case, further studies on the neural retina and Mueller cells are warranted. These results suggest that macular diseases including cystoid macular edema, AMD and macular hole may be related to the low density of glia in the foveal region.

b

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Fig. 13. ERG summary (a, a-wave, and b, b-wave). Preconditioning performed 24 h before ischemia significantly improved recovery of the a- and b-waves 1, 3 and 7 days after 1 h of ischemia. There was a nonsignificant trend toward greater recovery in the first 120 min after ischemia, especially in the b-wave. $ = Preconditioned; ) = non-preconditioned; Pre = 24 h after preconditioning. All values are means 8 SD (preconditioned: n = 10 per group, non-preconditioned: n = 4 per group).

Mueller Cells in the Preconditioned Retinal Ischemic Injury Rat [59]

Within 1 h of ischemia-reperfusion, Mueller cells become reactive, as reflected by an enhanced phosphorylation of extracellular signal-regulated kinases [60]. Inhibition of activation of these kinases results in an elevated degree of ganglion cell death [60], likely due to inhibition of the release of neurotrophic factors by Mueller cells. The neuronal cell loss accompanying ischemia results mainly from an accumulation of excessive amounts of glutamate, as well as from overstimulation of ionotropic glutamate receptors [61].

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The basis for the phenomenon of ischemic preconditioning is that short periods of ischemic stress do not lead to irreversible cell injury (in contrast to prolonged ischemic stress), but paradoxically to an adaptive mechanism that results in resistance to a subsequent ischemic stress. Cerebral ischemia is a potent modulator of gene expression. Gene regulation, including immediate early genes, is required for programmed neuronal death after trophic factor deprivation and is predicted to be involved in apoptosis triggered by cerebral ischemia [62]. Hypoxia induces upregulation of gene products [63]. It is reported that the tolerance to ischemia is induced by the incomNishikawa

Preconditioned

Non-preconditioned

a

Pre

60 min

1 day

3 days

Pre

60 min

1 day

3 days

Preconditioned

Fig. 14. Immunohistochemistry (a : GS, and b : GFAP). GS immunoreactivity

showed no significant differences between non-preconditioned and preconditioned groups prior to (Pre) and 60 min after ischemia. One day after ischemia, GS immunoreactivity decreased in both groups. Three days after ischemia, GS immunoreactivity had recovered only in the preconditioned group (a). Three days after 1 h of ischemia GFAP immunoreactivity increased in the retinas in the non-preconditioned group. In the preconditioned group, only slight GFAP immunoreactivity was found. Nuclei were stained by methyl green (b).

Non-preconditioned

b

pletely defined protective factors in hippocampal neurons [64]. In the myocardium [65, 66] and in the brain [67, 68], the phenomenon of ischemic tolerance has been shown. However, once ischemia has begun, preconditioning by a previous, brief non-damaging period of ischemia cannot protect the retina. This is significant in the search for effective methods to prevent or treat ischemic injury in the central nervous system because preconditioning utilizes the endogenous protective potential of tissues. In 1998, it was demonstrated that preconditioning provided complete protection against retinal ischemic injury in rats [69], although the mechanism was unclear. It was thought that Mueller cells play a central role in the homeostatic regulation of the retina [40]. In the present

Electroretinogram Figure 12 shows the representative electroretinogram (ERG) 3 days after 1 h of ischemia with and without preconditioning. Preconditioning performed 24 h before ischemia significantly improved the recovery of the awaves and b-waves 1 day after 60 min of ischemia (p ! 0.01). Thirty, 60, 90 and 120 min after ischemia, only recovery of the a-wave was observed (p ! 0.05). There was

Mueller Cell Pathology and Disease

Neuroembryol Aging 2006–07;4:61–75

study, GS, which is the marker enzyme of Mueller cells, and GFAP, which is expressed in unhealthy Mueller cells, was investigated in the preconditioned retina. Data are shown as means 8 SD; p ! 0.05 was considered to be statistically significant (Student’s t test).

71

30 min +

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Fig. 15. Immunoblots of extracts of retinas with GS (a) and GFAP (b). Extracts from retinas 30 and 90 min after ischemia yielded a

single band at the same position, which corresponded to 44 kDa for GS. In the non-preconditioned group, 1, 3 and 7 days after ischemia extracts yielded a slightly positive band. Three and 7 days after ischemia, the GS band was recovered only in the preconditioned group (a). + = Preconditioned; – = non-preconditioned. One, 3 and 7 days after 1 h of ischemia, the GFAP band had increased in the retinas, which corresponded to 50 kDa for GFAP in the non-preconditioned group. In the preconditioned group, the band was faint (b).

a nonsignificant trend toward greater recovery in the first 120 min after 60 min of ischemia, especially in the case of the b-wave (fig. 13). Immunohistochemical Staining There was no significant difference in GS immunoreactivity between non-preconditioned and preconditioned groups prior to and 60 min after ischemia. One day after ischemia, GS immunoreactivity decreased in both groups. Three days after ischemia, GS immunoreactivity recovered only in the preconditioned group (fig. 14a). Three days after 1 h of ischemia, the retinas showed increased GFAP immunoreactivity in the non-preconditioned group. In the preconditioned group, only slight GFAP immunoreactivity was observed (fig. 14b). 72

Neuroembryol Aging 2006–07;4:61–75

0.1 0

b

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30 min

60 min

90 min

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Fig. 16. Assay of relative GS (a) and GFAP content changes (b) measured by ELISA. A high GS concentration was observed in the extracts from retinas 30, 60, 90 and 120 min after ischemia. In the non-preconditioned group, 1, 3 and 7 days after ischemia, the extracts yielded low concentrations of GS. Three and 7 days after ischemia, GS had recovered in the preconditioned group only (p ! 0.05; a). One, 3 and 7 days after 60 min of ischemia, the retinas showed increased GFAP in the non-preconditioned group. In the preconditioned group, however, a low concentration of GFAP was found (p ! 0.05; b). $ = Preconditioned; ) = non-preconditioned; Pre = 24 h after preconditioning. All values are means 8 SD; preischemic: n = 4, others: n = 3 per group.

Immunoblots of Extracts of Retinas with GS and GFAP In the extracts from the retinas 30 and 90 min after ischemia, there was a single band at the same position, which corresponded to 44 kDa for GS. In the non-preconditioned group, 1, 3 and 7 days after ischemia extracts yielded a slightly positive band. Three and 7 days after ischemia, the GS band had recovered only in the preconditioned group (fig. 15a).

Nishikawa

One, 3 and 7 days h after 60 min of ischemia, the retinas demonstrated an increased GFAP band, which corresponded to 50 kDa for GFAP in the non-preconditioned group. In the preconditioned group, only a slight band was observed (fig. 15b). Assay of Relative GS and GFAP Contents Measured by ELISA The changes in GS and GFAP concentrations in the retina are shown in figure 16. A high GS concentration was observed in the extracts from the retinas 30, 60, 90 and 120 min after ischemia. In the non-preconditioned group, the extracts obtained 1, 3 and 7 days after ischemia yielded low GS concentrations. Three and 7 days after ischemia, GS had recovered only in the preconditioned group (p ! 0.05; fig. 16a). The retinas 1, 3 and 7 days after 60 min of ischemia showed increased GFAP in the non-preconditioned group. However, the preconditioned group had a low concentration of GFAP (p ! 0.05; fig. 16b). The results obtained from ELISA agreed with those obtained from Western blot analysis. Mechanisms possibly involved in the protection of the heart that results from classical ischemic preconditioning, i.e. the ability of short periods of ischemia to protect the heart against a subsequent, more prolonged period of ischemic stress initiated several minutes later, have been discussed [70]. In retinal ganglion and Mueller cells in culture, sublethal hypoxia induced tolerance to glutamate and anoxia and increased heat shock protein (HSP70) expression. The protective effect of sublethal hypoxia in these cultured cells was abolished by the HSP-70 synthesis inhibitor quercetin [71]. Another potentially protective protein whose expression was increased after ischemic preconditioning in the brain was the free-radical scavenger superoxide dismutase [72]. In 1999, Li and Roth [73] demonstrated that adenosine, acting through the A1 and A2a receptors, is a critical component in the induction of ischemic tolerance after preconditioning in the retina. On the other hand, they considered that Mueller cells play a central role in the homeostatic regulation of the retina. In the present study, we investigated the role of Mueller cells in the process of preconditioning rescue. We chose 5 min of ischemia to serve as a brief ischemic stimulus for preconditioning because the ERG exhibited a nearly complete recovery 1 day after ischemia. Studies were designed to test the ability of preconditioning to preserve retinal function after 60 min ischemia. This duration of ischemia was chosen because it induced an apMueller Cell Pathology and Disease

proximately 10% recovery of the baseline ERG a- and bwave amplitudes 7 days after ischemia, and because it caused significant histologic alterations. We regarded GS as the enzyme that serves functional glioneuronal interactions and GFAP as the pathological marker. Petito et al. [74] showed that 3 and 24 h after ischemia, GS immunoreactivity increased in the rat hippocampus. Tanaka et al. [75] demonstrated that GS immunoreactivity increased transiently on day 2 after ischemia and decreased to the initial level on day 5 in the gerbil hippocampus. On day 2 after ischemia, and more prominently on day 3, reactive astrocytes were intensely stained for GFAP in the hippocampal formation. In the present study, we observed that GS immunoreactivity showed no significant difference between the non-preconditioned and preconditioned groups 30, 60, 90 and 120 min after ischemia. One day after ischemia, GS immunoreactivity decreased in both groups. Three and 7 days after ischemia, GS immunoreactivity had recovered only in the preconditioned group. These results showed that Mueller cells were disrupted by an ischemic insult after 1 day but that the functions had recovered after 3 and 7 days only in the preconditioned group. One, 3 and 7 days after 1 h of ischemia, the retinas increased GFAP immunoreactivity in the non-preconditioned group, while in the preconditioned group, they exhibited only slight GFAP immunoreactivity. These results showed that Mueller cells were damaged after 1, 3, and 7 days in the non-preconditioned group but that they were rescued in the preconditioned group. In summary, our results suggest that the mechanism of preconditioned retinal ischemia might relate to Mueller cells in the retina.

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48 Fairman WA, Vandenberg RJ, Arriza JL: An excitatory amino-acid transporter with properties of a ligand-gated chloride channel. Nature 1995;375:599–603. 49 Derouiche A, Rauren T: Coincidence of Lglutamate/L-aspartate transporter (GLAST) and glutamine synthetase (GS) immunoreactions in retinal glia: evidence for coupling of GLAST and GS in transmitter clearance. J Neurosci 1995;42:131–143. 50 Schultze M: Zur Anatomie und Physiologie der Retina. Arch Mikrosk Anat 1866;2:175– 286. 51 Wolff E: Anatomy of the Eye and Orbit, ed 6. Philadelphia, Saunders, 1968. 52 Gass JDM: Mueller cell cone, an overlooked part of the anatomy of the fovea centralis: hypotheses concerning its role in the pathogenesis of macular hole and foveomacular retinoschisis. Arch Ophthalmol 1999; 117: 821–823. 53 Burris CJ, Sterling P, Schein S: One Mueller cell per foveal cone in macaque monkey. Invest Ophthalmol Vis Sci 1999;40:S238. 54 Rasmussen KE: The Mueller cell: a comparative study of rod and cone retinas with and without retinal vessels. Exp Eye Res 1974;19: 243–257. 55 Rasmussen KE: A morphometric study of the Mueller cell in rod and cone retinas with and without retinal vessels. Exp Eye Res 1975;20:151–166. 56 Sarks SH: Aging and degeneration in the macular region: a clinico-pathological study. Br J Ophthalmol 1976;60:324–341. 57 Penfold PL, Provis JM, Furby JH, Gatenby PA, Billson FA: Autoantibodies to retinal astrocytes associated with age-related macular degeneration. Graefes Arch Clin Exp Ophthalmol 1990;228:270–274.

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58 Gurne DH, Tso MO, Edward DP, Ripps H: Antiretinal antibodies in serum of patients with age-related macular degeneration. Ophthalmology 1991;98:602–607. 59 Nishiyama T, Nishikawa S, Tomita H, Tamai M: Mueller cells in the preconditioned retinal ischemic injury rat. Tohoku J Exp Med 2000;191:221–232. 60 Akiyama H, Nakazawa T, Shimura M, Tomita H, Tamai M: Presence of mitogen-activated protein kinase in retinal Mueller cells and its neuroprotective effect ischemiareperfusion injury. Neuroreport 2002; 13: 2103–2107. 61 Osborne NN, Casson RJ, Wood JP, Chidlow G, Graham M, Melena J: Retinal ischemia: mechanisms of damage and potential therapeutic strategies. Prog Retin Eye Res 2004; 23:91–147. 62 Atkins PT, Liu PK, Hsu CY: Immediate early gene expression in response to cerebral ischemia. Friend or foe? Stroke 1996; 27: 1682– 1687. 63 Takagi H, King GL, Ferrara N, Aiello LP: Hypoxia regulates vascular endothelial growth factor receptor KDR/Flk gene expression through adenosine A2 receptors in retinal capillary endothelial cells. Invest Ophthalmol Vis Sci 1996;37:1311–1321. 64 Kirino T, Tsujita Y, Tamura A: Induced tolerance to ischemia in gerbil hippocampal neurons. J Cereb Blood Flow Metab 1991;11:299– 307. 65 Parratt JR, Szekeres L: Delayed protection of the heart against ischemia. Trends Pharmcol Sci 1995;16:351–355. 66 Martin HB, Walter CL: Preconditioning: an endogenous defense against the insult of myocardial ischemia. Anesth Analg 1996; 83:639–645.

67 Kogure K, Kato H: Altered gene expression in cerebral ischemia. Stroke 1993; 24: 2121– 2127. 68 Simon RP, Niiro M, Gwinn R: Prior ischemic stress protects against experimental stroke. Neurosci Lett 1993;163:135–137. 69 Roth S, Li B, Rosenbaum PS, Gupta H, Goldstein IM, Maxwell KM, Gidday JM: Preconditioning provides complete protection against retinal ischemic injury in rats. Invest Ophthalmol Vis Sci 1998;39:777–785. 70 Parratt JR: Protection of the heart by ischemic preconditioning: mechanisms and possibilities for pharmacological exploitation. Trends Pharmacol Sci 1994;15:19–24. 71 Caprioli J, Kitano S, Morgan JE: Hyperthermia and hypoxia increase tolerance of retinal ganglion cells to anoxia and excitotoxicity. Invest Ophthalmol Vis Sci 1996; 37: 2376– 2381. 72 Kato H, Kogure K, Araki T, Liu XH, Kato K, Itoyama Y: Immunocytochemical localization of superoxide dismutase in the hippocampus following ischemia in a gerbil model of ischemic tolerance. J Cereb Blood Flow Metab 1995;15:60–70. 73 Li B, Roth S: Retinal ischemic preconditioning in the rat: requirement for adenosine and repetitive induction. Invest Ophthalmol Vis Sci 1999;40:1200–1216. 74 Petito CK, Chung MC, Verkhovsky LM, Cooper AJL: Brain glutamine synthetase increases following cerebral ischemia in the rat. Brain Res 1992;569:275–280. 75 Tanaka H, Araki M, Masuzawa T: Reaction of astrocytes in the gerbil hippocampus following transient ischemia: immunocytochemical observations with antibodies against glial fibrillary acidic protein, glutamine synthetase, and S-100 protein. Exp Neurol 1992;116:264–274.

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Neuroembryol Aging 2006–07;4:76–84 DOI: 10.1159/000103583

Published online: June 5, 2007

Disruption of Sonic Hedgehog Signaling Affects Axon Routing in the Mouse Optic Chiasm Yanli Hao a, b Jun Wang a, c Chung-Kit Chan a Sun-On Chan a Departments of Anatomy, a Chinese University of Hong Kong, Hong Kong, SAR, b Guangzhou Medical College, Guangdong, and c Department of Anatomy and Embryology, School of Basic Medical Sciences, Peking University, Beijing, China

Key Words Axon guidance
Mouse
Visual pathway, retina

Abstract The morphogen Sonic hedgehog (Shh) has recently been shown as an axon guidance molecule in the development of central neural pathways. We investigated here whether Shh serves its axon guidance role in patterning axon course in the mouse optic chiasm. Using immunocytochemistry, Shhlike staining was observed in the caudal regions of the ventral diencephalon in embryonic day (E) 13 embryos. At E14 and E15, the staining became obvious on the optic axons in lateral regions of the chiasm and in the optic tract, but not in the midline region. Blocking Shh signaling with cyclopamine in brain slice preparations of the optic pathway produced several abnormalities in routing of optic axons at the chiasm. Cyclopamine treatment in E13 brain slices reduced the midline crossing and generated a misrouting of axons in the post-midline region of the chiasm. This misrouting was further proven in E14 and E15 brain slices, in which a substantial increase in the retinoretinal axons was observed after cyclopamine treatment. Midline crossing at these developmental stages was not affected. These findings indicated that Shh protein was localized in a regionally and temporally specific pattern in the mouse optic chiasm, which might regulate midline crossing and targeting of axons into the optic tract at different stages of pathway development. Copyright © 2007 S. Karger AG, Basel

© 2007 S. Karger AG, Basel 1661–3406/06/0042–0076$23.50/0 Fax +41 61 306 12 34 E-Mail [email protected] www.karger.com

Accessible online at: www.karger.com/nba

Introduction

In the mouse retinofugal pathway, axons from both eyes meet at the ventral midline of the diencephalon to form the optic chiasm. Within this structure, most optic axons follow a stereotyped pathway crossing the midline and growing into the optic tract on the opposite side of the brain. A small population of axons deviates from the crossing axons before reaching the midline and projects to the ipsilateral optic tract, forming a bilateral projection. Another population of axons leaves the major fiber projection after crossing the midline, and projects to the contralateral optic stalk, generating an interretinal projection. This retinoretinal projection was viewed as an erroneous development of the pathway and is corrected during later stages of development. Although mechanisms of axon divergence in the mouse chiasm have been extensively studied [1], that of targeting of axons into the optic tract in the post-midline region of the chiasm remains to be elucidated. Sonic hedgehog (Shh), a member of the hedgehog family, is originally found in the notochord and the floor plate of the embryo and plays important roles in signaling spinal cord development [2]. Precursor of Shh is cleaved proteolytically to generate an N-terminal fragment (NShh) that is involved in all signaling functions, and a Cterminal fragment that catalyzes the process of cleavage and introduces cholesterol modification to N-Shh [3]. Shh binds to its receptor Patched (Ptc) and releases inhi-

Prof. S.O. Chan Department of Anatomy, Chinese University of Hong Kong Shatin, N.T. Hong Kong, SAR (China) Tel./Fax +852 2609 6898, E-Mail [email protected]

bition of Ptc to another surface molecule Smoothened (Smo), which triggers the signal transduction pathway [4, 5]. In mouse embryos, Shh transcripts have been demonstrated in the retinal ganglion cells by in situ hybridization, and have been shown to promote differentiation of neuroepithelial cells in the optic disk and the optic stalk [6]. Similar functions of Shh are reported in the chick retina, where ectopic expression of Shh produces multiple defects in retinal development, including expansion of pigment epithelium, enlargement of optic stalk and reduction of neural retina [7]. Shh has also been shown to inhibit extension of retinal neurites in vitro and affect routing of axons in the chick visual pathway [8]. In Pax-2 mutant mice, the presence of exuberant Shh at the ventral midline of the hypothalamus generates a failure of axon crossing at the midline [9], demonstrating a crucial role of Shh in early formation of the optic chiasm. However, it remains unknown whether Shh is involved in regulating axon growth and guidance in the optic chiasm of mouse embryos at later stages of pathway development. We have addressed this issue using brain slice cultures of the mouse optic pathway and showed an important function of Shh in guiding axons into the optic tract after crossing the midline of the chiasm.

Materials and Methods Animal and Tissue Preparations The experimental procedures were approved by the Animal Ethic Committee of the Chinese University of Hong Kong. Timemated C57 pigmented mice were obtained from the University Animal House. The date when the vaginal plug was found was designated as embryonic day 0 (E0). Embryos at the age of E13– E15 were used. Pregnant mothers were killed by cervical dislocation and embryos were taken out and decapitated. The heads were immersed in freshly prepared 4% paraformaldehyde in 0.1 M phosphate buffer at pH 7.4. Tissues were embedded in a gelatinalbumen mixture and sectioned using a vibratome to obtain frontal and horizontal sections of the retinofugal pathway. The brain sections were collected in 0.1 M phosphate-buffered saline (pH 7.4).

tions were processed with the same procedure but without addition of primary antibody. No obvious staining was detected in the control preparations. Preparation of Brain Slices Brain slices containing the primary visual pathway were prepared using the protocol described in our previous reports [13, 14]. Embryos aged from E13 to E15 were removed by cesarean section, decapitated and maintained in Dulbecco’s modified Eagle’s medium/F12 medium. Brain slices containing the optic pathway from the eyes to the optic tract were prepared and cultured for 5 h in Dulbecco’s modified Eagle’s medium/F12 supplemented with 10% fetal bovine serum (Life Technologies, Carlsbad, Calif., USA) at 37 ° C. During this incubation period, oxygen was supplied three times over the culture. Some brains slices were treated with cyclopamine (1 mg dissolved in 200
l 95% ethanol; catalogue No. C988400, from Toronto Research Chemicals, North York, Canada) at various concentrations at the start of the culture. This steroidal alkaloid has been shown to inhibit specifically Shh signaling in embryos [15] by direct binding to Smo [16]. Control preparations for this experiment included brain slices cultured without cyclopamine. After the culture, brain slices were fixed in 4% paraformaldehyde. Retinal axons from one eye were labeled with DiI (Invitrogen, Carlsbad, Calif., USA). The brain slices were then stored for 1–3 weeks, depending on the distance of dye diffusion, at room temperature. In another experiment, brain slices were immunostained with antibody against the stage-specific embryonic antigen-1 (SSEA-1) to visualize the arrangement of the chiasmatic neurons in the E14 diencephalon. Brain slices treated with cyclopamine (50
g/ml) and those in the control group (cultured without cyclopamine) were fixed with paraformaldehyde and sectioned. Horizontal sections containing the ventral diencephalon were processed for immunostaining using the protocol described above, with primary antibody against SSEA-1, which is expressed specifically on the chiasmatic neurons [17, 18]. Specificity of the antibody staining was tested by processing some sections in the absence of primary antibody. No significant staining was found in the sections examined. Confocal Microscopy Immunostaining in brain slices and DiI-labeled axons in whole-mount preparations of the optic pathway were examined using a confocal imaging system [BioRad MRC600 connected to a Zeiss Axiophot photomicroscope (Zeiss, Oberkochen, Germany) or Olympus FV300 (Olympus, Tokyo, Japan)]. Digital images were processed using Adobe PhotoShop (version 8.0; Adobe, Seattle, Wash., USA) and PowerPoint (Microsoft, Redmond, Wash., USA).

Immunohistochemistry The immunocytochemical procedure followed that described in our earlier studies [10, 11]. Brain sections were incubated in 10% normal donkey serum in 0.1 M phosphate-buffered saline for 1 h, followed by primary antibody 5E1 (IgG, 1:10; from Developmental Studies Hybridoma Bank, Iowa City, Iowa, USA) that recognizes N-Shh [12]. After incubation in Cy3-conjugated secondary antibody (1:100, Jackson Laboratories, Bar Harbor, Me., USA), sections were mounted and examined using confocal microscopy (BioRad MRC600; BioRad, Hemel Hempstead, UK). Control sec-

Analyses of Axon Trajectory at the Optic Chiasm The methods for analyzing axon routing in the chiasm followed those described in our earlier report [19]. Two parameters were measured in this study. First, the degree of axon crossing was measured in two defined areas (each 100 ! 200
m2) flanking both sides of the midline of the chiasm in projected images of the E13–E14 pathways using the MetaMorph software (Universal Imaging, Downingtown, Pa., USA). Midline crossing was represented as ratio of pixel intensity of axons and growth cones in the postmidline versus pre-midline region of the chiasm. Second, develop-

Shh Signaling in the Mouse Optic Chiasm

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ment of the retinoretinal projection was measured in the E14–E15 pathways. It was represented as average pixel intensity in a defined area straddling the first 200-
m segment into the contralateral optic stalk. The data in the experimental groups were compared with the corresponding controls using one-way ANOVA in the InStat software (GraphPad, San Diego, Calif., USA).

Results

Shh Expression at the Mouse Optic Pathway During the early period of axon growth (E13–E15) in the mouse retinofugal pathway, Shh-like immunoreactivity was barely detected in the retina. Only a weak staining was observed in the optic stalk in the embryos examined (n = 7 at each developmental stage; fig. 1a, b). In the ventral diencephalon, where the optic axons form the chiasm, Shh-like staining was found in specific domains in the optic pathway. For example, at E13 (n = 5), Shh-like staining was localized largely in an inverted V-shaped region in the caudal regions of the diencephalon (fig. 1c), a pattern similar to that of a population of early generated chiasmatic neurons [17, 20]. This pattern was altered at later developmental stages. In E14 and E15 embryos, Shh-like staining was most obvious in the optic tract and less in the optic stalk and the regions of the chiasmatic neurons, but not in the midline region of the chiasm (fig. 1d, e). This pattern of staining was supported by labels in frontal sections of the E14 diencephalon, in which strong Shh-like signals were observed in the optic stalk, lateral regions of the chiasm and the optic tract (fig. 1f, g). These signals were substantially reduced at the midline. Similar changes in Shh-like staining were observed in the E15 pathway (fig. 1h, i), suggesting a regionally regulated localization of Shh protein in the developing mouse optic pathway. Cyclopamine Disrupted the Routing of Early Axons in E13 Chiasm In mouse embryos, the first axons arrived at the chiasm and the optic tract at E13 [21, 22]. In control brain slices prepared from E13 embryos, which had been cultured in the absence of cyclopamine, a number of dyefilled optic axons were observed in the chiasm (fig. 2a). These axons traversed the midline in parallel bundles and some had already entered the contralateral optic tract (fig. 2b). Treatment with cyclopamine, a specific inhibitor of the Shh signaling mediator Smo, produced two obvious routing abnormalities in the chiasm. First there was an obvious reduction in axon crossing at the chiasmatic midline (fig. 2c), which was supported by quantitative analyses of axon crossing in brain slices treated with var78

Neuroembryol Aging 2006–07;4:76–84

ious concentrations of cyclopamine (fig. 2e). Significant reduction in axon crossing (p ! 0.05, compared with control) was only observed when the concentration of cyclopamine was 625
g/ml but not below (fig. 2e). Second, the optic axons, after crossing the midline, tended to grow to the optic tract in a defasciculated manner (fig. 2d). Some of these axons, instead of growing as a bundle towards the optic tract, were deviated rostrally to the contralateral optic stalk, which was not seen in any control preparations at similar age. Routing Abnormalities Caused by Cyclopamine in the E14 and E15 Optic Chiasm At E14, more retinal axons were added to the retinofugal pathway (fig. 3a). Treatment of the pathway with cyclopamine at this stage did not generate obvious reduction in the crossing of axons at the midline (compare fig. 3a, c), which was supported by quantitative measurements showing no significant difference in axon crossing in brain slices treated with cyclopamine at the concentrations tested (5–50
g/ml) compared with the control (p 1 0.05). However, it was noted that there was a dramatic increase in axons that project to the contralateral optic stalk (compare fig. 3b, d). This routing abnormality was dose dependent, as evidenced by the quantitative measurements of retinoretinal axons in the contralateral optic stalk that shows a significant increase after treatment with cyclopamine at concentrations of 25–50
g/ml (p ! 0.05; fig. 3f). Similar results were obtained in the E15 brain slices. Whilst there was no apparent difference in the axon crossing at the midline (fig. 4a, c), there was an obvious enlargement in the retinoretinal projection in brain slices treated with cyclopamine (fig. 4b, d). Measurement of these axons revealed a significant increase, compared with control, after treatment with cyclopamine at 25 or 50
g/ml (p ! 0.05), but not at 5
l/ml (p 1 0.05; fig. 4e). The findings confirmed the effect of cyclopamine on the formation of an enlarged interretinal projection, and suggested that Shh signaling via the Ptc-Smo pathway is crucial for the targeting of some retinal axons to the optic tract in the post-midline region of the chiasm. Cyclopamine Treatment Did Not Affect Arrangement of the Chiasmatic Neurons To investigate whether the changes in axon routing after cyclopamine treatment are secondary effects due to alteration of cellular patterning in the ventral diencephalon, we examined the arrangement of the chiasmatic neurons in control and cyclopamine-treated brain slices of Hao /Wang /Chan /Chan

Fig. 1. Confocal micrographs showing

Shh-like immunoreactivity in the retinofugal pathway of mouse embryos. Rostral is to the top in a–e, midline is indicated by the white arrows in c–i ; to the right in a, and to the left in b. a, b No obvious Shhlike staining was detected in the retina (R) and the lens (L), but a weak staining was consistently found in the optic stalk (OS) of E14 and E15 embryos. c In horizontal sections of the E13 diencephalon, Shh-like staining was localized largely in an inverted V-shaped domain (bound by the broken lines) whose rostral tip extends to the midline of the chiasm (indicated by the tip of the vertical arrow). d, e In horizontal sections of the E14 and E15 diencephalon, Shh-like staining was most intense in the optic tract (empty arrow). Moderate signal was detected in the optic stalk and the chiasmatic neurons (asterisk) but not in the midline region of the chiasm. f–i This regional change in Shh-like staining was also observed in frontal sections of the diencephalon, showing localization of Shh protein in the optic stalk (OS), optic tract (OT) and lateral regions of the chiasm. The staining was reduced dramatically at the midline. Scale bars = 200
m in a–e, and 100
m in f–i.

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80

Pixel intensity of axons (%, post- vs. pre-midline region)

Fig. 2. Blocking Shh signaling with cyclopamine in brain slice cultures of E13 optic pathways produced abnormalities in axon routing in the chiasm. Rostral is to the top. The midline is indicated by the arrows. a, b In control preparations, retinal axons from the dye-injected eye traversed the optic stalk (OS), optic chiasm and arrived at the optic tract (OT). c, d After cyclopamine treatment (50
g/ml), there was an obvious reduction in midline crossing at the chiasm. Some axons strayed away from the major bundle that heads toward the optic tract and innervated inappropriate regions in the brain, including the contralateral optic stalk (axons indicated by empty arrow). e A plot showing significant reduction in midline crossing after treatment of brain slices with 25 or 50
g/ml cyclopamine (* p ! 0.05 vs. control). Numbers in the columns indicate the number of animals examined. Scale bars = 200
m in a and c, 100
m in b and d.

70

8

60

7

50

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8

40 30 20 10 0 Control

e

E14 embryos using antibody against SSEA-1. In horizontal sections of control brain slices (n = 3), SSEA-1-positive chiasmatic neurons existed as an inverted V-shaped domain in the caudal regions of the ventral diencephalon (fig. 5a). Rostral to this domain was an extension of cells that protrudes into the midline of the chiasm (fig. 5b). In brain slices treated with cyclopamine (50
g/ml, the highest dose used in the axon routing study; n = 3), we found no obvious change in the arrangement of these chiasmatic neurons (fig. 5c). The characteristic configuration of these cells was observed in the caudal parts of the diencephalon and the midline (fig. 5d), showing cyclopamine did not produce a major reorganization in the ar80

*

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5 µg/ml

25 µg/ml

50 µg/ml

Cyclopamine

rangement of chiasmatic neurons, which are believed to be crucial in directing axon routing in the mouse optic chiasm.

Discussion

We have investigated in this study the localization of Shh protein in the retinofugal pathway of mouse embryos and the effects of blocking Shh signaling on axon routing in the optic chiasm. The major findings are: (1) Shh protein is not detectable in the retina at the developmental stages examined. (2) Shh protein is localized largely in Hao /Wang /Chan /Chan

8

7

8

7

Control

5 µg/ml

e

25 µg/ml

Pixel intensity of retinoretinal axons

Pixel intensity of axons (%, post- vs. pre-midline region)

100 90 80 70 60 50 40 30 20 10 0

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Cyclopamine

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Cyclopamine

Fig. 3. Perturbation of Shh signaling generated an enlarged retinoretinal projection in E14 optic pathway. Rostral is to the top. Midline is indicated by the vertical arrow. a, b Most axons in the control preparations were routed into the optic tract (OT) after crossing the midline. A few axons were deviated to the contralateral optic stalk (empty arrow). c, d In brain slices treated with cyclopamine (50
g/ml), there was a substantial increase in axons that project to the contralateral optic stalk (empty arrow). e Cyclopamine at the concentrations tested had no apparent effect on the midline crossing in the E14 chiasm. f However, treatment with 25 or 50
g/ml cyclopamine produced a significant enlargement of the retinoretinal projection (* p ! 0.05 vs. control). Numbers in the columns indicate the number of animals examined. Scale bars = 200
m in a and c, 100
m in b and d.

the lateral regions of the chiasm and at the optic tract, and to a lesser extent in the caudal regions of the diencephalon. (3) Disruption of Shh signaling with cyclopamine produces a reduction in midline crossing in the E13 chiasm and generates an increase in misrouting of axons into the contralateral optic stalk in E14 and E15 chiasms.

The current findings on the localization of Shh protein are apparently different from the expression pattern of Shh transcripts in the optic pathway of mouse embryos at similar developmental stages. Previous reports using in situ hybridization have shown Shh mRNA expression in the ganglion cell layer in the retina [6, 23], where Shh protein is not detectable. Moreover, Shh transcripts are

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Pixel intensity of retinoretinal axons

Fig. 4. Blocking Shh signaling with cyclopamine generated an enlargement in the retinoretinal projections in E15 optic pathways. Rostral is to the top. Midline is indicated by the solid arrows. a, b In control preparations, a few axons were found in the contralateral optic stalk (empty arrow). c, d Treatment with cyclopamine (25
g/ml) produced an obvious enlargement in the retinoretinal projections (empty arrow). e This change was supported by quantitative measurements shown in the plots (* p ! 0.05 vs. control). Numbers in the columns indicate the number of animals examined. Scale bars = 200
m in a and c, 100
m in b and d.

180

*

160 140

7

100

7

80 60

7 6

40 20 0 Control

e

found in diencephalic domains rostral and caudal to the path of retinal axons [8, 24], but not in the chiasm per se where Shh protein is localized. One possible explanation for these discrepancies is that Shh protein is transported away or diffuses from its sites of synthesis to its sites of actions in the optic pathway. Our finding of a weak but consistent staining of Shh in the optic stalk supports an anterograde transport of Shh protein in the optic axons, which is probably responsible for its reported functions in the specification of the glial lineage from neuroepithelial cells in the mouse optic stalk [6]. Similarly, it would not be unreasonable to argue that Shh in the chiasm is deposited from its site of synthesis in the diencephalic cells, which are also immunopositive to the Shh antibody. Given the diffusible nature of a morphogen, Shh may dif82

*

120

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5 µg/ml

25 µg/ml

50 µg/ml

Cyclopamine

fuse across cells and deposit in the lateral regions of the chiasm to exert its action on the growing retinal axons. However, a retinal contribution to the Shh protein in the chiasm cannot be ruled out. Another important finding in this study is that interruption of Shh signaling generates a number of routing abnormalities in brain slice preparations of the optic pathway. The most obvious routing error is the increase in misrouting of axons into the contralateral optic stalk, forming an enlarged retinoretinal projection. This misrouting takes place in the post-midline regions of the chiasm, indicating a crucial role of Shh in regulating axon targeting to the optic tract after crossing the midline. Furthermore, interruption of Shh signaling reduces substantially the crossing of the early axons in the E13 chiasm, but Hao /Wang /Chan /Chan

Fig. 5. Cyclopamine treatment did not

generate any changes to the configuration of the chiasmatic neurons in the mouse diencephalon. Rostral is to the top. Midline is indicated by the arrow. a, b In the horizontal section of the ventral diencephalon in control slices, the chiasmatic neurons marked by the SSEA-1 antibody were found in the caudal diencephalon and at the midline of the chiasm (empty arrow). b Magnified view of a. c, d This characteristic arrangement was obviously not disrupted by treatment with cyclopamine (50
g/ml). Note the section in c, d was taken at a level more dorsal to a, b, as indicated by the presence of the third ventricle (asterisk), therefore there is a slight variation in the arrangement of these neurons. Scale bars = 200
m in a, c, and 100
m in b, d.

has no apparent effect on the crossing of those generated later. The action of Shh on retinal axon growth at the midline and the mechanism of channeling axons into the optic tract are unknown. Previous experiments have shown in the chick that Shh is able to inhibit retinal neurite growth, and ectopic expression of Shh interferes with axon growth and prevents axon crossing in the chiasm [8]. This inhibitory nature of Shh to retinal axon growth gains support from an earlier finding in the mouse showing the crossing of the first generated retinal axons is highly impeded in the presence of high levels of Shh at the chiasmatic midline in Pax-2 mutants [9]. On the contrary, in the spinal cord of mouse and rat embryos, Shh has been shown to collaborate with netrin-1 as chemoattractants for guiding commissural axons to the ventral midline [25]. This variation in activity suggests that Shh may regulate axon growth depending on the neuronal type and species of animals studied. In fact, a recent study has demonstrated that it is the level of Shh that determines whether it is promotive or inhibitory to neurite extension in chick retinal explants [26]. It remains to be determined whether Shh acts to pattern the chiasmatic routing in the mouse through an inhibitory action, as demonstrated in the chick optic pathway and the mouse mutants. Preliminary findings in our laboratory have shown a localization of Shh receptors Ptc and Smo on retinal axons and growth cones, supporting an interaction of retinal axons with Shh

protein in the chiasm through the Ptc-Smo pathway. The current functional studies using the Smo inhibitor cyclopamine support this possible interaction, but the spatial and temporal localization of Ptc and Smo in the chiasm at different developmental stages have to be worked out in future investigations. It is noted that the localization of Shh in the mouse chiasm is remarkably similar to that of heparan sulfate proteoglycans [10]. This coincidence in spatial distribution suggests Shh may interact with these proteoglycans for its actions on axon routing. It has been shown in the mouse cerebellum that heparan sulfate is able to regulate the Shh response for granule cell proliferation [27]. Moreover, mice deficient in sulfation or polymerization of heparan sulfate proteoglycans also display phenotypes of enlarged retinoretinal projection [28, 29], providing further support to the interaction of Shh with other molecules in the extracellular environment to control axon routing at the mouse optic chiasm.

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Acknowledgment This study was partially supported by grants from the Research Grant Council of the Hong Kong Special Administrative Region (project Nos. CUHK 4417/03M and CUHK 4420/05M).

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References 1 Williams SE, Mason CA, Herrera E: The optic chiasm as a midline choice point. Curr Opin Neurobiol 2004;14:51–60. 2 Marti E, Bumcrot DA, Takada R, McMahon AP: Requirement of 19K form of Sonic hedgehog for induction of distinct ventral cell types in CNS explants. Nature 1995;375: 322–325. 3 Goodrich LV, Scott MP: Hedgehog and Patched in neural development and disease. Neuron 1998;21:1243–1257. 4 Alcedo J, Ayzenzon M, Von Ohlen TV, Noll M, Hooper JE: The Drosophila smoothened gene encodes a seven-pass membrane protein, a putative receptor for the hedgehog signal. Cell 1996;86:221–232. 5 Murone M, Rosenthal A, de Sauvage FJ: Sonic hedgehog signaling by the PatchedSmoothened receptor complex. Curr Biol 1999;9:76–84. 6 Dakubo GD, Wang YP, Mazerolle C, Campsall K, McMahon AP, Wallace VA: Retinal ganglion cell-derived sonic hedgehog signaling is required for optic disc and stalk neuroepithelial cell development. Development 2003;130:2967–2980. 7 Nasrallah I, Golden JA: Brain, eye and face defects as a result of ectopic localization of sonic hedgehog protein in the developing rostral neural tube. Teratology 2001;64:107– 113. 8 Trousse F, Marti E, Gruss P, Torres M, Bovolenta P: Control of retinal ganglion cell axon growth: a new role for Sonic hedgehog. Development 2001;128:3927–3936. 9 Torres M, Gomez-Pardo E, Gruss P: Pax2 contributes to inner ear patterning and optic nerve trajectory. Development 1996; 122: 3381–3391. 10 Chung KY, Leung KM, Lin L, Chan SO: Heparan sulfate proteoglycan expression in the optic chiasm of mouse embryos. J Comp Neurol 2001;436:236–247.

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11 Lin L, Taylor JSH, Chan SO: Changes in expression of fibroblast growth factor receptors during development of the mouse retinofugal pathway. J Comp Neurol 2002; 451: 22–32. 12 Ericson J, Morton S, Kawakami A, Roelink H, Jessell TM: Two critical periods of sonic hedgehog signaling required for the specification of motor neuron identity. Cell 1996; 87:661–673. 13 Chung KY, Taylor JSA, Shum DKY, Chan SO: Axon routing at the optic chiasm after enzymatic removal of chondroitin sulfate in mouse embryos. Development 2000; 127: 2673–2683. 14 Leung KM, Taylor JSH, Chan SO: Enzymatic removal of chondroitin sulphates abolishes the age-related axon order in the optic tract of mouse embryos. Eur J Neurosci 2003; 17: 1755–1767. 15 Incardona JP, Gaffield W, Kapur RP, Roelink H: The teratogenic Veratrum alkaloid cyclopamine inhibits Sonic hedgehog signal transduction. Development 1998; 125: 3553– 3562. 16 Chen JK, Taipale J, Cooper MK, Beachy PA: Inhibition of Hedgehog signaling by direct binding of cyclopamine to Smoothened. Genes Dev 2002;16:2743–2748. 17 Marcus RC, Mason CA: The first axon growth in the mouse optic chiasm: axon patterning and the cellular environment. J Neurosci 1995;15:6389–6402. 18 Lin L, Cheung AWS, Chan SO: Chiasmatic neurons in the ventral diencephalon of mouse embryos – changes in arrangement and heterogeneity in surface antigen expression. Dev Brain Res 2005;158:1–12. 19 Lin L, Chan SO: Perturbation of CD44 function affects chiasmatic routing of retinal axons in brain slice preparations of the mouse retinofugal pathway. Eur J Neurosci 2003;17: 2299–2312. 20 Leung KM, Margolis RU, Chan SO: Expression of phosphacan and neurocan during early development of mouse retinofugal pathway. Brain Res Dev Brain Res 2004;152: 1–10.

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21 Godement P, Vanselow J, Thanos S, Bonhoeffer F: A study in developing visual systems with a method of staining neurons and their processes in fixed tissue. Development 1987;101:697–713. 22 Colello RJ, Guillery RW: The early development of retinal ganglion cells with uncrossed axons in the mouse: retinal position and axonal course. Development 1990; 108: 515– 523. 23 Wallace VA, Raff MC: A role for Sonic hedgehog in axon-to-astrocyte signaling in the rodent optic nerve. Development 1999; 126: 2901–2909. 24 Marcus RC, Shimamura K, Sretavan D, Lai E, Rubenstein JLR, Mason CA: Domains of regulatory gene expression and the developing optic chiasm: correspondence with retinal axon paths and candidate signaling cells. J Comp Neurol 1999;403:346–358. 25 Charron F, Stein E, Jeong J, McMahon AP, Tessier-Lavigne M: The morphogen Sonic hedgehog is an axonal chemoattractant that collaborates with netrin-1 in midline axon guidance. Cell 2003;113:11–23. 26 Kolpak A, Zhang JH, Bao ZZ: Sonic hedgehog has a dual effect on the growth of retinal ganglion axons depending on its concentration. J Neurosci 2005;25:3432–3441. 27 Rubin JB, Choi YJ, Segal RA: Cerebellar proteoglycans regulate sonic hedgehog responses during development. Development 2002; 129:2223–2232. 28 Inatani M, Irie F, Plump AS: Mammalian brain morphogenesis and midline axon guidance require heparan sulfate. Science 2003;302:1044–1046. 29 Pratt T, Conway CD, Tian NMML, Price DJ, Mason JO: Heparan sulphation patterns generated by specific heparan sulfotransferase enzymes direct distinct aspects of retinal axon guidance at the optic chiasm. J Neurosci 2006;26:6911–6923.

Hao /Wang /Chan /Chan

Neuroembryol Aging 2006–07;4:85–101 DOI: 10.1159/000103735

Published online: June 8, 2007

Aging of the Mammalian Circadian Timing System: Changes in the Central Pacemaker and Its Regulation by Photic and Nonphotic Signals Marilyn J. Duncan Department of Anatomy and Neurobiology, University of Kentucky Medical Center, Lexington, Ky., USA

Key Words Circadian rhythm
Dorsal raphe nucleus
Entrainment
Phase shift
Serotonin
Suprachiasmatic nucleus
Vasoactive intestinal peptide

Abstract Aging alters many aspects of endogenously regulated, 24hour (circadian) rhythms, such as their amplitude, relationship to the ambient lighting cycle, and sensitivity to phase resetting signals. In order to elucidate the mechanisms responsible for these age-related changes, many studies have investigated age-related changes in the neural components of the circadian timing system, which include the hypothalamic suprachiasmatic nucleus (SCN), the site of the mammalian master circadian pacemaker, its afferent projections from the retina, thalamus, and midbrain raphe, and its efferent projections to the hypothalamus, thalamus and limbic system. Studies have shown that the SCN exhibits age-related changes in electrical activity rhythms in the absence of neurodegeneration. Also, aging selectively decreases SCN expression of the circadian clock genes, Clock and Bmal1, as well as vasoactive intestinal peptide. The latter participates in mediating light-induced phase resetting of the circadian pacemaker and the amplitude of the preovulatory luteinizing hormone surge, which are both attenuated by aging. Furthermore, by decreasing serotonin 5-HT7 receptors in the dorsal raphe, aging reduces one type of nonphotic input to the SCN. This review describes which aspects of circadian

© 2007 S. Karger AG, Basel 1661–3406/07/0042–0085$23.50/0 Fax +41 61 306 12 34 E-Mail [email protected] www.karger.com

Accessible online at: www.karger.com/nba

rhythm regulation exhibit changes during aging, as well as which mechanisms appear to be spared or remain to be explored. Copyright © 2007 S. Karger AG, Basel

Introduction to the Circadian Timing System

A broad range of biological processes and behaviors, including but not limited to body temperature, hormone secretion, rest-activity, and performance and alertness, are governed by an endogenous circadian (approximately 24-hour) timing system. The circadian timing system helps to ensure that physiological and behavioral processes occur at appropriate times, in relation to changes in the external environment. This system enables organisms to anticipate and prepare for predictable daily changes in environmental conditions, such as illumination and temperature, and optimize the time of their activities. For example, in nocturnal, fossorial rodents, circadian regulation of the onset of daily activity allows them to emerge from their burrows at dusk, when the risk of predation is lower. The circadian timing system also coordinates multiple biological processes. For example, the rhythm of circulating cortisol levels is characterized by an increase in the early morning that mobilizes energy stores, which facilitates waking and the onset of activity [1]. Also, lower nocturnal temperatures characterize the human body temperature rhythm, promoting energy savings during

Dr. Marilyn J. Duncan Department of Anatomy and Neurobiology, University of Kentucky Medical Center 800 Rose Street Lexington, KY 40536-0298 (USA) Tel. +1 859 323 4718, Fax +1 859 323 5946, E-Mail [email protected]

the time of rest. Furthermore, the onset of sleep is closely associated with the body temperature rhythm; the latency to fall asleep is shortest during the nocturnal decline in body temperature [2, 3]. As anyone with experience with shift work or transmeridian jet travel knows, disruption of the circadian rhythms leads to fatigue and malaise. Research has shown that chronic desynchronization of circadian rhythms also has more severe consequences, such as increased risk of cardiovascular disease, impaired responsiveness to chemotherapy, impaired cognition and memory, and decreased longevity [4–16]. Therefore, the circadian timing system is essential for maintaining optimal health and well-being in humans as well as animals and the age-related changes identified in this system may be deleterious to health. The circadian timing system has several components, including an internal master circadian pacemaker that is not only intrinsically rhythmic but is also responsive to environmental time-giving cues, such as daily changes in ambient light. In mammals, the circadian pacemaker is located in a cluster of hypothalamic neurons called the suprachiasmatic nucleus (SCN) [17–21]. Retinal ganglion cells densely innervate the SCN, as well as a secondary circadian structure, the thalamic intergeniculate leaflet (IGL), providing the circadian timing system with exquisite sensitivity to light [22–26]. Therefore, the circadian timing system has also been called ‘the circadian visual system’ [27, 28]. Light can reset the phase of the endogenous rhythms generated by the circadian pacemaker in the SCN. By using a variety of efferent neuronal pathways, the SCN communicates timing information to many neural and hormonal substrates that regulate various physiological processes [29–31]. Although many tissues and organs exhibit intrinsic oscillations, based on in vitro studies, the phase of the majority of these oscillations appears to be coordinated by the SCN [32].

Overview of the Effects of Aging on the Circadian Timing System

Circadian rhythmicity begins before birth and persists throughout the life span, but undergoes functional changes both during development and later with aging. The cells that give rise to the SCN are born as early as gestational days 13–17 in rats (gestational length
21–22 days) and 9.5–13 in hamsters (gestational length
15–16 days) [33–35]. In rats, the circadian pacemaker begins to function by gestational days 19–22, when the SCN exhibits day-night differences in glucose metabolism in vivo or 86

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electrical activity in vitro [36, 37]. During the prenatal period, before retinal innervation of the SCN has developed, and during the early postnatal period, the phase of the SCN pacemaker is determined by maternal signals, including circulating levels of melatonin [38–40]. Within several days to a week after birth, the rodent SCN acquires retinal innervation, and environmental lighting can override maternal signals to entrain the circadian pacemaker [41–43]. Light continues to be a powerful entraining signal throughout the life span, although other signals influence the phase of circadian rhythms as well. As individuals progress through middle age and old age, many alterations in the circadian timing system occur; these alterations will be the focus of this review. They include a decrease in circadian rhythm amplitude, increased fragmentation, a decrease in the synchrony or coupling among rhythms, altered phase relationship of rhythms to the external temporal environment, and diminished capacity to respond to timing signals [44–50]. These alterations have been observed during aging in humans as well as laboratory animals. The present review will focus on animal studies and the underlying aging mechanisms that they have revealed. Although some findings concerning aging changes in human circadian rhythms will be described, the reader is referred to two recent reviews for more extensive information on this topic [51, 52]. A decrease in circadian rhythm amplitude is a common effect of aging that has been observed in many studies of the circadian rhythms of body temperature, sleep, and activity in a variety of rodent species [53–55]. Also, decreases in the amplitude of endocrine rhythms, such as circulating melatonin, prolactin, and luteinizing hormone rhythms, have been observed in humans and rodents [56, 57]. A decreased amplitude is sometimes considered an indicator of decreased rhythm robustness. Relevant to this issue, increased fragmentation of locomotor rhythms, or breakdown into multiple components, during aging has been observed in some studies [58–61]. However, in contrast to fragmentation of circadian rhythms under entrained conditions (exposure to a light: dark cycle), old hamsters appear to be resistant to treatments (e.g. long-term exposure to constant light or exposure to dark pulses during constant light) that in young hamsters typically induce ‘splitting’ of circadian wheel running rhythms into two discrete components beginning at approximately 12-hour intervals [62, 63]. Aging affects the timing, or phase, of circadian rhythms. It is well known that in humans, the onset of Duncan

nocturnal sleep and morning waking occur earlier in the elderly than in young adults. Laboratory studies in nocturnal rodents have also indicated that aging alters the circadian phase angle, i.e. the relationship of the timing of a circadian rhythm to the lighting cycle, although the magnitude and direction of this effect is variable. In mice, for example, the onset of nocturnal running wheel activity was approximately 40 min later in older individuals (19–22 months of age) than in younger adults (6–9 months of age) [64]. In hamsters, in contrast, aging has been reported to advance the onset of running activity such that middle-aged hamsters (12–13 months old) began nocturnal running activity approximately 15 min before lights off, while young hamsters (3 months old) began running approximately 30 min after lights off [50]. Another study of hamsters showed that the influence of aging on the time of activity onset depended upon the environmental lighting cycle (photoperiod) [58]. In that study, aging was associated with significant advances (
1–2.5 h) in activity onset in middle-aged and old hamsters as compared to young hamsters when the animals were exposed to a photoperiod with 6 h of light per day. In contrast, when these animals were exposed to a 14-hour light photoperiod, the time of activity onset was not significantly different among the ages. Although it has been suggested that the age-related change in the time of activity onset might be related to a decrease in the endogenous circadian period length with aging, the latter effect has not been observed consistently [60, 62, 63, 65]. Most studies of the effects of aging on circadian rhythms have used a cross-sectional approach comparing groups of individuals of different ages. Furthermore, in many cases the experimental animals had been exposed to one or more lighting cycles before determination of endogenous circadian period length under constant conditions, and, often, the animals’ histories are not described. Since light history influences period length, differences in light history might contribute to the variability in the effects of aging on period length among studies. Interestingly, two longitudinal studies of hamsters in which the animals were exposed to unchanged lighting conditions from early adulthood have reported different effects of aging on endogenous circadian period length. In one study, in which the hamsters were bilaterally enucleated (i.e. exposed to constant darkness) at 35 days of age, the endogenous circadian period gradually and significantly decreased from
24.2 hours at 15 weeks of age to
24.0 hours at 70 weeks of age [62]. In contrast, another study, in which hamsters were exposed to constant dim light (!1 lx) from 8 weeks of age until death,

found no significant change in the average endogenous circadian period length throughout the rest of the life span [60]. Thus, a change in endogenous circadian period length seems to be a variable rather than a consistent effect of aging and it may be influenced by lighting conditions and/or other conditions occurring during the life span. Aging changes in phase angle of circadian rhythms may also result from changes in sensitivity to entraining signals or time givers (‘zeitgeber’), such as light. Indeed, age-related changes in sensitivity to light have been observed in both mice and hamsters [66, 67]. In mice, aging decreases phase shifts induced by a range of irradiances, including those that induce maximal responses in young adults [67]. In hamsters, aging increases the threshold level of light needed to induce a detectable phase shift and decreases the magnitude of phase shifts induced by intermediate levels of light [66]. However, when exposed to high-intensity light, old hamsters respond as well as young hamsters, suggesting that increased illumination can overcome aging attenuation of light sensitivity [66]. This may also be the case in humans, since a recent study using light intensities which people typically encounter has shown that healthy elderly adults exhibit equivalent phase shifts to those of young adults [68]. Circadian rhythms in humans and rodents are phase shifted not only by light, but also by many so-called ‘nonphotic’ stimuli, including exposure to several hours of darkness, exercise, sleep deprivation, melatonin, and some anxiolytic or serotonergic drugs [for review, see ref. 69, 70]. Aging decreases the phase shifts induced by some of these nonphotic stimuli, e.g. the benzodiazepine, triazolam, or the serotonergic drug, 8-OH-DPAT [48, 49, 71, 72]. However, the effectiveness of other phase-shifting stimuli, such as melatonin injections or dark exposure, is either not altered by aging or not consistently affected in all studies, respectively [48, 63, 73]. Therefore, aging does not decrease the responsiveness of the circadian pacemaker to all time-giving signals. Finally, aging appears to modulate interactions between photic and nonphotic time-giving signals. For example, administration of serotonergic drugs that act on either 5-HT1A or 5-HT1B receptors attenuates photic phase shifts [74, 75]; this effect of 5-HT1A agonists (e.g. 8-OH-DPAT) is reduced by aging [76]. When considered together, these data demonstrating various alterations in the expression and regulation of circadian rhythms suggest that aging induces multiple, discrete changes in the mechanisms underlying the circadian timing system, rather than a simple, overall loss

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of function. Indeed, aging may differentially affect various circadian rhythms within an individual, although few studies have investigated this issue. When body temperature, drinking, and activity rhythms were monitored simultaneously in old rats, aging differentially affected these rhythms [47]. While circadian rhythms were generally weaker or absent in these old rats, some individuals maintained a robust circadian rhythm in one parameter, but not in the other two [47]. Thus, loss of rhythmicity was not an all-or-none phenomenon. Theoretically, the age-related changes observed in overt circadian rhythms might involve aging changes in central circadian rhythm generation, reception or processing of environmental timing information, and/or changes in transmission of circadian information to the anatomical substrates regulating various overt rhythms. Before reviewing the aging changes that have been identified in each of these properties, the basic structure and function of the circadian timing system will be described.

Basic Anatomy and Physiology of the Circadian Timing System

The circadian timing system consists of a central pacemaker, input pathways conveying temporal information to the pacemaker, and output pathways communicating temporal information from the pacemaker to multiple brain regions and other organs [for review, see ref. 28, 77]. The anatomical location of the central pacemaker, the SCN, was first identified by lesion studies demonstrating that destruction of this nucleus eliminated circadian rhythms in many processes, including locomotor activity, body temperature, drinking, and circulating corticosterone levels [17, 18]. Later studies showed that the SCN in vitro exhibited
24-hour rhythms in electrical activity and release of the neuropeptide, vasopressin [19, 20]. Furthermore, transplantation of fetal SCN tissue to SCNlesioned, arrhythmic hosts reinstated circadian rhythms with a period length identical to that of the donor [21]. Thus, the SCN fulfills the basic requirements of a circadian pacemaker, i.e. endogenous rhythmicity and ability to drive overt circadian rhythms. The molecular mechanisms underlying the circadian pacemaker are thought to consist of interacting transcriptional and translational feedback loops, based on investigations throughout the last decade in several organisms, including fungi, yeast, and mice. In the mouse SCN, the transcription factors CLOCK and BMAL1 form het88

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erodimers that bind to E-boxes in the promoter regions of the Per1, Per2, Per3, Cry1 and Cry2 genes, thereby activating their transcription [78–81]. The expression of the Per and Cry genes in the SCN exhibits circadian rhythms with peaks during the subjective day and troughs during the subjective night [82, 83]. The proteins PER1, PER2, CRY1 and CRY2 also exhibit circadian rhythms that follow their respective mRNA rhythms by
4–6 hours [83]. Cytoplasmic PER and CRY proteins form complexes that enter the nucleus and inhibit Clock:Bmal1 activation of transcription. Thus, once the nuclear levels of PER and CRY increase to a critical point, these proteins inhibit their own transcription, thus forming a negative feedback loop. Over time, the levels of PER and CRY decrease as a result of degradation, thereby allowing their transcription to begin again. The entire cycle lasts about 24 hours. Other genes encoding proteins that interact with and modulate these basic feedback loops have been identified as well. In spite of the fact that this transcription-translation model has been generally accepted as the molecular underpinnings of circadian rhythms, as yet this model may be incomplete [84, 85]. As reported recently, genetic deletion of Clock, a gene encoding one member of the CLOCK: BMAL1 heterodimer forming the positive feedback loop, does not prevent expression of circadian rhythms [85]. Also, single deletion of genes involved in the negative feedback loop, such as Per1, Per2, Cry1 and Cry 2, does not disrupt circadian rhythms; the effect of such deletions appears to depend on environmental illumination and/or the presence or absence of other genes [for review, see ref. 84]. The molecular mechanisms driving circadian rhythms are still under active investigation that will no doubt lead to further elucidation. Although circadian rhythmicity in electrical activity or gene expression is expressed autonomously by individual cells [86, 87], the SCN is not a collection of homogeneous, rhythmic cells, but a heterogeneous structure [77, 88–90]. The SCN contains both neurons, most of which are GABA immunoreactive (IR) [91], and a large proportion of astrocytes [92]. Furthermore, the SCN exhibits anatomical and functional subdivisions, which were first described for the rat SCN using the terms ventrolateral and dorsomedial, and subsequently described as the SCN ‘core’ and ‘shell’ [93]. Extensive studies of the SCN of several mammalian species now indicate that this terminology is simplistic and that the organization of the SCN is more complex and consists of overlapping regions and gradients (for detailed descriptions, the reader is directed to recent reviews [28, 77] and the articles described Duncan

therein). Although two important functions of the circadian pacemaker, generation of circadian rhythms and photoresponsiveness, are generally associated with certain topographical areas of the SCN and specific peptidergic cell types, they are not strictly localized to discrete, sharply bounded regions [94]. Generation of circadian rhythmicity, or pacemaking, has been associated with the dorsomedial region of the SCN, because this region exhibits circadian rhythms in Per1 and Per2 mRNA expression during exposure to constant darkness, suggesting endogenous rhythmicity [95– 98]. The dorsomedial region exhibits many cells that synthesize vasopressin, and vasopressin mRNA expression in the dorsomedial SCN also exhibits a circadian rhythm in hamsters and rats exposed to constant darkness [95, 96, 98]. In contrast, expression of mRNA for vasoactive intestinal peptide (VIP), a neuropeptide localized mainly in the ventral SCN, does not exhibit circadian rhythms during constant conditions [99]. In addition to the dorsomedial SCN, a small subregion of the caudal aspect of the ventral region of the hamster SCN, called the calbindin-D28-containing subnucleus, has also been implicated in circadian rhythm generation [21, 100]. Lesion studies have shown that this region is necessary for the expression of overt circadian rhythms in several functions, including locomotor activity, drinking, body temperature, circulating melatonin levels and circulating corticosterone levels [100]. Surprisingly, while the calbindin subnucleus is required for the expression of circadian rhythms, it does not exhibit intrinsic circadian rhythms in either electrical activity or circadian clock gene expression although it does express a rhythm in subcellular calbindin localization [97, 101, 102]. The calbindin-containing cells in the SCN co-express several neuropeptides, including gastrin-releasing peptide (GRP), VIP, and substance P, but not vasopressin [103]. Furthermore, calbindin-IR SCN cells make direct, reciprocal connections with most other known types of SCN cells, with the exception, once again, of the vasopressin-IR cells [103]. VIP has been implicated in the generation of circadian rhythms. When mutant mice lacking VIP or one of the VIP receptor subtypes (VPAC2) were exposed to constant darkness, they exhibited abnormal circadian rhythms or arrhythmicity [104–106]. Furthermore, GRP administration can induce circadian rhythms in electrical activity in arrhythmic SCN neurons from mice lacking the VPAC2 receptor [106, 107]. Thus, VIP and GRP, which are both synthesized in the ventral SCN and are concentrated in the calbindin region in the ham-

ster SCN [94, 108], appear to be involved in circadian rhythm generation. Although tract-tracing studies show that retinal fibers innervate broad aspects of the SCN [23, 25, 94], most functional studies suggest that the ventrolateral aspect of the SCN responds the most quickly to photic stimulation [95–98]. Photoresponsiveness of the SCN, which is necessary for light-induced phase resetting, has been measured by light induction of Per1 and Per2 mRNA expression or Fos expression. Light exposure at times that reset the circadian pacemaker induces Per1 and Per2 mRNA more rapidly in the ventrolateral SCN, although this induction gradually spreads throughout most of the SCN [95–98]. Furthermore, light induces acute expression of Per1 in VIP-IR cells, located in the ventral SCN, but not vasopressin-IR cells [109], which are more abundant in the dorsal SCN [94, 108]. As well as Per induction, photic induction of Fos in the SCN occurs most robustly in the ventral aspect [110, 111]. Furthermore, in rats, photic induction of Fos was identified in SCN neurons immunoreactive for GRP but not for VIP [110]. In contrast, in mice, photic induction of Fos occurred in several cell types in the SCN, including cells immunoreactive for VIP or vasopressin, with a greater percentage of the former cell type showing Fos expression [112]. The calbindin subnucleus of the hamster SCN is not only necessary for the expression of overt circadian rhythms, as described above, but also participates in photic phase shifts. Light pulses stimulate Per1 and Per2 expression in the calbindin subnucleus and calbindin-expressing SCN cells receive direct retinal input [97, 103]. Because administration of calbindin antisense oligonucleotides into the SCN blocks photic phase shifts and photic induction of Per2 mRNA expression, calbindin in the SCN appears to be necessary for light-induced resetting of the circadian pacemaker [102]. Thus far, the calbindin-containing subnucleus has been observed and extensively studied only in the hamster SCN. It is possible that other species exhibit a functionally analogous SCN subregion with somewhat different neurochemical characteristics. For example, in the mouse SCN, a region with a similar neuroanatomical location has been demonstrated to expresses calretinin but little or no calbindin [94]. However, the potential function of calretinin in the mouse SCN has not been reported. As mentioned above, many cells of the hamster calbindin subnucleus co-express GRP [103]. The SCN of the rat and mouse also contain GRP cells that exhibit some of the functional characteristics of the calbindin-con-

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IGL

G

Fig. 1. Diagram representing the major af-

ferent projections to the SCN. DRN = Dorsal raphe nucleus; MRN = median raphe nucleus; GHT = geniculohypothalamic tract; RHT = retinohypothalamic tract [modified from fig. 6 shown in ref. 219].

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DRN

SCN MRN Eye

taining cells in the hamster SCN. For example, GRP neurons in the mouse and rat SCN are retinorecipient and express c-fos after light exposure at night, when photic phase shifts occur [108, 110, 113–115]. In addition, GRPcontaining cells of the rat or mouse SCN do not exhibit detectable rhythms of Per expression, similar to cells of the hamster calbindin subnucleus [96, 115]. GRP participates in photic phase shifts, as shown by its ability to reset the circadian pacemaker in a manner similar to light when it is microinjected into the hamster SCN in vivo; this effect is potentiated by co-administration with VIP or peptide histidine isoleucine [116, 117]. Also, mice lacking GRP receptors exhibit attenuated photic phase shifts [118]. Thus, GRP-IR neurons in the SCN, which in the hamster are localized mainly to the calbindin subnucleus, appear to be physiologically important in circadian timekeeping in several species. The SCN pacemaker can be reset by timing information of two basic types, photic and nonphotic, as indicated above. The effect of either type of timing signal is determined by the circadian phase at which the signal occurs [for review, see ref. 69]. Light in the late subjective day or early subjective night induces phase delays, while light in the mid- to late subjective night induces phase advances and light in the early or midsubjective day has little or no effect [119]. In contrast to light, nonphotic signals presented in the midsubjective day typically induce phase advances, while those at night induce small phase delays or no change in phase [69]. As suggested by the differential temporal sensitivity of the circadian pacemaker to photic and nonphotic signals, different neural mechanisms control the phase resetting effects of these two types of signals. 90

HT

RHT

Tract-tracing and lesion experiments have demonstrated that different pathways convey photic and nonphotic information to the SCN (see fig. 1 for a diagram of these pathways). Light information is transmitted to the SCN by the retinohypothalamic tract, which originates in a discrete subset of retinal ganglion cells [22, 23, 26] and innervates SCN GRP neurons in rats and hamsters and also VIP neurons in the rat SCN [120–122]. The majority of retinal ganglion cells innervating the SCN are intrinsically photosensitive and express the photopigment, melanopsin, which modulates the responsiveness of the SCN to light [26, 123–125]. The major neurotransmitter released by the retinohypothalamic tract is glutamate, although pituitary adenylate cyclase-activating peptide appears to act as a co-transmitter and modulator [126–129]. Light presented during the subjective night, when it induces phase shifts, increases expression of Per1 and Per2 first in the ventral SCN and later in the dorsal SCN [98]. Induction of Per1 in the SCN is necessary for photic phase shifts [98]. Nonphotic timing signals, such as triazolam injections, 8OH-DPAT injections, and dark ‘pulses’ (periods of dark exposure preceded and followed by constant light exposure), decrease the expression of Per1 and Per2 in the SCN when administered in the midsubjective day [130–132]. Attenuation of SCN Per1 expression, by administration of antisense oligonucleotides, in the midsubjective day induces phase shifts, mimicking nonphotic signals [133]. The major neuroanatomical pathway conveying nonphotic timing signals to the SCN is the geniculohypothalamic tract originating in IGL [134–138]. The geniculohypothalamic tract consists of fibers that are immunoreactive for neuropeptide Y (NPY), GABA, and; in some species, enkephalin or neurotensin [134, 139]. Nerve fiDuncan

Fig. 2. Diagram representing the major ef-

ferent projections from the SCN (constructed from information presented by Watts [220]). BNST = Bed nucleus of the stria terminalis; DMN = dorsal medial nucleus; LS = lateral septum; MPN = medial preoptic nucleus; PMA = premammillary area; PVN = paraventricular nucleus; PVT = paraventricular thalamic nucleus; RCA = retrochiasmatic area; VMN = ventromedial nucleus.

LS BNST MPN

PVT PVN

IGL DMN

SCN

PMA

VMN RCA

bers containing NPY contact VIP- and calbindin-containing cells in the ventral SCN region [103, 140]. These fibers are important because microinjections of NPY near the SCN mimic nonphotic phase shifts by inducing circadian phase advances when administered during the midsubjective day [116, 141]. Furthermore, lesions of the IGL block many nonphotic phase shifts [136, 138]. Serotonin-containing cells of the median raphe nucleus of the midbrain, which project to the SCN as well as to other brain regions, are also necessary for nonphotic phase shifts induced by triazolam injections [142–144]. Circadian information generated by the SCN pacemaker is transmitted to multiple targets in the hypothalamus, thalamus and limbic system, by efferent projections arising from both the dorsal and ventral SCN regions (see fig. 2 for a diagram summarizing these pathways) [29, 30]. The physiological role of all of these efferent projections is not yet understood, although different neuroanatomical pathways appear to mediate circadian regulation of different functions [for review, see ref. 31]. For example, VIP projections from the SCN to the gonadotropin-releasing hormone (GnRH) neurons have been implicated in regulating circadian release of the preovulatory surge of luteinizing hormone (LH) [145–151], while vasopressin projections from the SCN to the paraventricular nucleus regulate the circadian rhythm of corticosterone secretion in rats [152]. SCN GABAergic projections to the ventrolateral preoptic area of the rat affect sleep maintenance [153]. As well as communicating by efferent neuronal pathways, the SCN communicates by releasing diffusible signals, such as transforming growth factor-
and prokineticin 2, that appear to regulate the circadian rhythm of locomotor activity [154–157].

Although aging renders some brain regions vulnerable to neurodegeneration, there is relatively little evidence suggesting that this phenomenon occurs to any major extent in the SCN. The dorsolateral but not the ventromedial SCN region exhibits decreased cell density in young adults and aged rats as compared to weanlings [158]. Also, the dorsolateral SCN region of aged rats exhibits a higher percentage of cells with ultrastructural changes associated with degeneration, such as organelle and cytoplasmic vacuolation, lysosomal aggregation, and lipofuscin granules [158]. In spite of the appearance of these markers of neurodegeneration, there is no significant difference in cell numbers in the dorsolateral, ventromedial or total SCN between young adulthood and old age [158]. Similarly, a study using unbiased stereological methods found no significant effects of aging on the SCN volume, cell size, total number of cells, number of neurons, or number of astrocytes in rats [159]. In contrast to the absence of striking morphological changes during normal aging, the SCN does exhibit functional changes. One of the first functional changes reported was an alteration in the SCN circadian rhythm of glucose utilization in vivo in middle-aged female rats [160]. This finding suggested that aging might alter SCN neuronal activity rhythms, and subsequent in vitro studies showed that rats, hamsters, and mice exhibit age-related reductions in the amplitude and/or precision of circadian rhythms of SCN electrical activity [47, 161–163]. Studies of mice have shown that aging increases the proportion of electrically silent cells in the SCN during the

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91

*

SCN VIP mRNA (grain areal/cell)

50

25

0 Young

Middle aged

Old

Fig. 3. The effect of aging on VIP mRNA expression in the hamster SCN. Each bar represents the mean 8 SEM of the hybridization signal for each age group. Aging significantly decreased VIP mRNA expression [data redrawn from ref. 172] * p ! 0.05.

daytime, when the electrical activity is highest, and also decreases the spontaneous inhibitory postsynaptic currents [163]. Aging changes in SCN function appear to be functionally important because transplantation of aged hamsters with fetal SCN tissue can ameliorate aging changes in circadian rhythms in older animals [164–166]. For example, transplants of fetal SCN but not other brain tissue increase the endogenous free-running period of circadian activity rhythms in old but not young hamsters, and greatly potentiate phase shifts to triazolam [164, 165]. Furthermore, transplants of fetal SCN tissue to aged rats improve or restore circadian rhythms in body temperature, drinking and locomotor activity [166]. The effect of aging on SCN molecular rhythms appear to be less pronounced and more variable than age-related dampening of SCN electrical activity rhythms. Studies using in situ hybridization or RT-PCR found that aging decreased Cry 1 mRNA expression in the rat SCN but did not affect Per1 or Per2 mRNA expression in either the rat or hamster SCN [167, 168]. In the mouse SCN, aging attenuated Per2 but not Per1 mRNA expression at one of the two time points examined [169]. When the circadian rhythm of Per1 expression was monitored continuously in vitro from SCN of Per1-luciferase transgenic rats, the only observed effect of aging was a decrease in the endogenous period length [170]. However, age-associated decreases in the expression of Clock mRNA and changes in the circadian rhythm of Bmal1 mRNA were found in the hamster 92

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SCN [168]. The recent finding that Clock is not essential for circadian rhythms may help to explain the finding that normal circadian rhythms of Per1 and Per2 mRNA persist in the SCN of old hamsters in spite of significant reduction of Clock mRNA expression [85]. Taken together, the results of these studies suggest that although there are some alterations, the molecular mechanisms underlying circadian rhythm generation are not strongly disrupted by aging. It is interesting to note that in the studies of aging described above, expression of circadian clock genes was assessed in the SCN as a whole, rather than in SCN regions or in specific types of neurons, such as those that synthesize the known neuropeptides or calbindin. Other studies have investigated the effects of aging on gene expression of specific neuropeptides located most abundantly in either the dorsal SCN (vasopressin) or ventral SCN (VIP). In male and female rats and male hamsters (fig. 3), aging decreases SCN expression of VIP mRNA but not vasopressin mRNA [171–173]. The selectivity of the effect of aging on VIP mRNA expression is also emphasized by findings that SCN mRNA expression of somatostatin, typically located in the dorsal SCN, or GAD67, an enzyme required for the synthesis of the neurotransmitter GABA that is broadly distributed throughout the SCN, are also apparently unaffected by aging [172, 174]. The age-related decrease in SCN VIP mRNA expression may help to explain the age-related decrease in the number of VIP-IR cells in the rat SCN that has been observed in some but not all studies [175–177]. Decreased VIP mRNA expression might suggest that there is decreased synthesis of VIP, and hence a decrease in the number of neurons expressing VIP at detectable levels. However, it is also possible that aging includes a loss of VIP synthesizing neurons. Because these neurons constitute only
3–5% of the total SCN neurons, a decrease in this population would not be incompatible with the finding that the total number of neurons in the rat SCN is not significantly altered by aging [159]. Aging has also been found to be associated with a decrease in the number of SCN VIP-IR cells in humans; this effect was observed in middle-aged and old men, but not women [178]. In contrast to the lack of effect of aging on vasopressin mRNA expression in the rat and hamster SCN, described above, an age-related decrease in vasopressin-IR neurons has been observed in the SCN of rats, voles, and humans [176, 179–181]. In humans, because the decrease in SCN vasopressin-IR cells was only observed in normal subjects at advanced ages, 81–100 years, but was also seen in younger demented patients, this phenomenon was thought to represent neurodegeneration, and to be unDuncan

likely to contribute to the normal aging changes in circadian rhythms that usually appear much earlier in the life span [179]. However, unlike the loss of SCN vasopressinIR cells late in life, a loss of the circadian rhythm of expression of these cells was observed in a younger human population (50–91 years old) [179]. In rats, the age-related decrease in SCN vasopressin-IR cells was found in 24- or 32-month-old animals, and apparently was not investigated at earlier ages [176, 177, 181]. Age-related decreases in neuropeptide expression in the rat SCN can be reversed or prevented by several treatments. For example, increased light intensity during the daytime prevented the age-related loss of vasopressin-IR cells in the rat SCN [177]. Also, intracerebroventricular administration of nerve growth factor to old rats restored SCN expression of both vasopressin-IR and VIP-IR cells to the levels observed in young, control-treated rats [176]. Thus, the aging circadian pacemaker exhibits some plasticity. Also, aging may induce compensatory changes, such as increased expression of mRNAs for the V1a and V1b vasopressin receptors in the rat SCN [173].

The findings that aging decreased the responsiveness of the circadian timing system to a variety of phase resetting signals suggested that aging might not only induce changes in the SCN pacemaker itself, but might alter input pathways conveying either photic or nonphotic information. As mentioned above, photic information that resets the mammalian circadian pacemaker is conveyed to the SCN by the retinohypothalamic tract, which originates from a subset of retinal ganglion cells. Light contacts the retina after passing through the lens. In humans, aging is known to be associated with increased lens opacity and occurrence of cataracts. Therefore, the effect of aging on lens transmittance was investigated in laboratory rodents. Although transmittance of light at a range of wavelengths (200–700 nM) was reduced by 8–50% in excised lens obtained from old (18–22 months) versus young (4–5 months) hamsters, this reduction was not large enough to account for the order of magnitude decrease in phase shifting to low intensity light [66, 182]. This study also investigated the effect of aging on the retinohypothalamic tract, by conducting monocular injections of the tract-tracer, cholera toxin/horseradish peroxidase. Aging did not significantly affect the histological appearance of the retinohypothalamic tract projec-

tions or the volume of the SCN that they innervated, although the density of the projections was not assessed quantitatively [182]. The foregoing study, in combination with others, suggested that rather than decreasing photic phase shifts by inducing major changes in lens opacity or retinohypothalamic tract projections, aging instead may have stronger effects on either the functional properties of the retinohypothalamic tract or the SCN processing of photic information. Although the effects of aging on glutamate release from the retinohypothalamic tract or glutamate receptors in the SCN have not been reported, effects of aging on photic intracellular signaling pathways have been identified. At the circadian times at which light induces phase shifts, light also induces activation of several immediate early genes, including c-fos and jun-B [183, 184]. Induction of these genes is required for photic phase shifts, because administration of antisense to c-fos or junB into the SCN reduces the magnitude of these phase shifts, similar to the effect of aging [185]. Aging significantly reduces photic induction of SCN expression of cfos mRNA or FOS protein in rats, hamsters and mice, as well as SCN expression of jun-B mRNA in mice [66, 67, 186, 187]. The age-related reduction in photic induction of SCN c-fos mRNA expression is associated with decreased photic induction of SCN cells expressing immunoreactivity for phosphorylated cyclic-AMP response element-binding protein (CREB), in the absence of any change in the number of CREB-IR SCN cells, suggesting that aging decreases light-induced activation of CREB [66]. Aging has also been found to decrease photic induction of Per1 expression in the hamster SCN [168]. The age-related reduction in light induction of Per1 and c-fos in the SCN may contribute to the age-related attenuation of photic phase shifts, which require these phenomena for maximal expression [185]. It is interesting to note that although photic stimulation of FOS in the ventral SCN occurs preferentially in GRP cells, it also occurs in some VIP cells [110, 113, 114]. VIP as well as GRP has been implicated in the phase resetting effect of light [104, 188]. Co-administration of VIP potentiates the phase shifts induced by SCN microinjections of GRP [117, 188]. Furthermore, photic phase shifts are impaired in mice deficient in VIP/peptide histidine isoleucine [104]. The age-related reduction in SCN VIP mRNA expression may lead to decreased VIP, and thus contribute to the attenuation of photic phase shifts observed in older animals [66, 67, 116, 117, 171, 172, 188]. Furthermore, aging is associated with decreased SCN sensitivity to VIP, as shown by decreased VIP stimulation

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of cAMP in the SCN in middle-aged rats [189]. Whether aging also attenuates photic phase shifts by downregulating GRP expression or signaling in the SCN has not been reported. As well as attenuating phase shifts induced by light, aging also reduces phase shifts stimulated by nonphotic signals, such as dark pulses and systemic injections of triazolam or 8-OH-DPAT [48, 71]. It is likely then that aging modulates either SCN processing of this information or the afferent pathways conveying nonphotic information to the SCN. Investigation of the former possibility has been hampered by the lack of a known cellular marker for SCN responsiveness to nonphotic stimuli. In contrast to light, nonphotic stimuli do not induce detectable Fos expression in the SCN [190–192]. Although most nonphotic zeitgebers utilize the NPYergic geniculohypothalamic tract, the effect of aging on this pathway has received relatively little attention. One study reported that the SCN content of NPY is decreased in middle-aged male rats by
35% compared to young adult rats [193]. Since the NPY in the SCN is derived virtually exclusively from the geniculohypothalamic tract, this finding suggested that NPY release in the SCN decreases during aging. SCN sensitivity to the phase-shifting effect of midday microinjections of NPY in vivo does not appear to change significantly with aging [Duncan, unpubl.]. Similarly, cAMP administration to the rat SCN in vitro during the midsubjective day induces robust phase shifts in the SCN electrical activity rhythm in old as well as young rats [194]. Thus, the aged SCN appears to be able to respond robustly to at least some types of signals that convey nonphotic information. In addition to the geniculohypothalamic tract, serotonergic fibers from the midbrain median raphe nucleus project to the SCN and this projection is necessary for the occurrence of nonphotic phase shifts that are induced by triazolam [142–144]. Although aging has been reported to lead to degenerative changes in serotonin terminals in many rat brain regions, this effect has not been shown to occur in the SCN [195, 196]. However, aging is associated with functional changes in the SCN which mimic partial deafferentation by decreasing extracellular serotonin levels. For example, the hamster SCN exhibits age-related increases in 5-HT1B receptor binding sites [197], which are typically located presynaptically and whose activation decreases serotonin release. Age-related changes in the hamster SCN also include increased serotonin reuptake sites [197], which would be expected to increase the clearance of serotonin from the synaptic cleft. These changes in the serotonergic system in the SCN that occur 94

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during aging may contribute to the reduced responsiveness of older hamsters to phase resetting by triazolam, since this phenomenon depends on serotonergic innervation of the SCN [48, 49]. In contrast to the mechanisms described above, other mechanisms may mediate the age-related decrease in responsiveness to phase resetting by 8-OH-DPAT [71], a serotonergic drug with high affinity for the 5-HT1A receptor subtype and moderate affinity for the 5-HT7 receptor subtype [198]. Because some studies have demonstrated that the SCN is a sufficient target site for 8-OHDPAT phase resetting [199, 200] (but also see [201]), the SCN appeared to be a likely site of age-related loss of serotonin receptors. However, autoradiographic studies did not detect any aging changes in the SCN in specific [3H]8OH-DPAT binding to either the 5-HT1A or 5-HT7 receptor subtype [202]. In contrast, the dorsal raphe of old hamsters exhibited a significant decrease of 50% in specific [3H]8-OH-DPAT binding to 5-HT7 but not the 5-HT1A receptor subtype, implicating this region as a site where aging affects the function of the circadian timing system (fig. 4a) [202]. Indeed, aging inhibits phase advances induced by microinjections of 8-OH-DPAT into the dorsal raphe [72] (fig. 4b). These phase shifts are mediated by the 5-HT7 receptor subtype, because they can be blocked by pretreatment with SB-26770A, a selective 5-HT7 receptor antagonist, or Rp-cAMPs, an antagonist for cAMP, the second messenger stimulated by 5-HT7 receptor activation [72, 203]. The absence of aging changes in 5-HT1A receptors in the SCN or dorsal raphe may help to explain the finding that aging does not affect the potentiation of photic phase shifts by the 5-HT1A agonist/ antagonist BMY7378 [204].

Effect of Aging on Output Pathways from the SCN

Another means by which aging might alter overt circadian rhythms is through changes in the output pathways that convey circadian signals from the master pacemaker to neural substrates regulating specific physiological functions or behaviors. Although these changes have not been extensively studied, there is evidence suggesting their importance in the effect of aging on the circadian regulation of the preovulatory LH surge in rats. In rats and other laboratory rodents, the LH surge occurs once every 4–5 days, in the late afternoon of proestrous, in response to an antecedent increase in plasma levels of estradiol secreted from the maturing ovarian follicles [205, 206]. This rise in estradiol exerts a positive feedback acDuncan

a

2

*

*

*

8-OH-DPAT Vehicle

20

Phase shift (h)

Specific [3H] 8-OH-DPAT binding (fmol/mg TE)

30

1

10

0

Young

Old

b

0

Young

Old

Fig. 4. The effect of aging on 5-HT7 receptors in the dorsal raphe nucleus of young (3–5 months) and old hamsters (17–19 months). a Specific binding of [3H]8-OH-DPAT (1.8 nM) to 5-HT7 receptors, using ritanserin (500 nM) to define nonspecific binding. TE = Tissue equivalent. * p ! 0.001. b Phase shifts induced by micro-

injections of 8-OH-DPAT into the dorsal raphe in the midsubjective day (zeitgeber time 6, i.e. 6 h before lights off). * p ! 0.05. [Data were redrawn from ref. 202 (a) and ref. 72 (b).]

tion on the GnRH neurons that increases the secretion of this hormone, causing enhanced pituitary LH release [207]. During the LH surge, GnRH neurons, located mainly in the septum/diagonal band of Broca/rostral preoptic area, are activated, as indicated by an increase in their expression of Fos [208–211]. In addition to the positive feedback action of estradiol, the rodent LH surge is also regulated by a circadian signal transmitted from the SCN [212–215]. Both VIP- and vasopressin-synthesizing neurons in the rat SCN project to the region of the GnRH neurons, but only the former have been demonstrated to make synaptic contacts with them [145]. GnRH neurons not only express the VIP/pituitary adenylate cyclase-activating peptide receptor subtype 2, but the percentage of VIP-innervated GnRH neurons expressing Fos increases at the time of the LH surge [145, 146, 149, 150]. Furthermore, intracerebroventricular administration of VIP antiserum or intra-SCN administration of VIP antisense oligonucleotides delays and attenuates activation of GnRH neurons and the LH surge [147, 148, 151]. These findings show that VIP projections from the SCN are important for regulating the timing and amplitude of the circadian preovulatory LH surge. In female rats, aging decreases VIP mRNA expression in the SCN [171]. In particular, the peak of VIP mRNA

rhythm that precedes the onset of LH surge in young animals does not occur in old animals [171]. Furthermore, Fos induction in GnRH neurons receiving VIP innervation is reduced during aging [150]; this effect can be mimicked in young rats by reducing SCN VIP with antisense oligonucleotides [151]. Although some evidence suggests that vasopressin projections from the SCN may also participate in regulating the LH surge [216–218], aging does not appear to alter vasopressin mRNA expression in the SCN [171, 172]. Thus, selective age-related changes in the SCN VIP output pathway to the GnRH neurons appear to mediate age-related changes in the circadian regulation of the LH surge.

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Summary and Conclusions

The circadian timing system emerges early in development and functions throughout the life span. During middle and old age, circadian rhythms undergo changes that include decreases in amplitude, alterations in their timing relative to the environment, and increased risk of fragmentation or cessation [45–50]. These changes vary among rhythms within an individual and among species. With normal aging, as opposed to neuropathological conditions, there is very little evidence of neurodegen95

eration of the components of the circadian timing system [159]. Instead, aging brings about many functional changes. In the master circadian pacemaker in the SCN, aging alters electrical activity rhythms [47, 161–163], expression of circadian clock genes (e.g. Clock and Bmal1) [168], and neuropeptide gene expression (e.g. VIP) [171–173]. With the exception of the latter, these studies have considered the SCN as a homogeneous unit. It is now evident that the SCN is a neuroanatomically and functionally heterogeneous structure, and further investigations of the effects of aging in discrete SCN regions, especially the calbindin subnucleus, are needed. Aging causes changes in pacemaker generation of electrical and molecular circadian rhythms and also reduces the sensitivity of the circadian timing system to both photic and nonphotic phase resetting signals [48, 49, 66, 67, 71, 72]. This reduced sensitivity is mediated in part by attenuation of the molecular mechanisms by which the SCN pacemaker responds to light [66, 67, 186, 187]. In addition, by decreasing 5-HT7 receptors in the dorsal raphe nucleus, aging attenuates

the sensitivity of one of the nonphotic input pathways to the SCN [72, 202]. Expression of overt circadian rhythms, such as rhythms in rest-activity-rest, body temperature, or circulating hormones, depends on SCN output to multiple brain regions and peripheral organs. Aging-induced deficits in the VIPergic SCN output pathway to the GnRH neurons regulating the LH surge have been demonstrated [150, 171]. Whether aging-induced alterations in VIP pathways are also part of the mechanism leading to aging changes in other circadian rhythms, and whether aging also affects other SCN output pathways, remains to be elucidated.

Acknowledgments Financial support from the National Institute on Aging (AG13418), as well as helpful comments from Dr. Sandra J. Legan, Dr. Lothar Jennes, and two anonymous reviewers on an earlier version of the manuscript, are gratefully acknowledged.

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172 Duncan MJ, Hill SA, Herron JM: Aging selectively suppresses vasoactive intestinal peptide messenger RNA expression in the suprachiasmatic nucleus. Brain Res Mol Brain Res 2001;87:196–203. 173 Kalamatianos T, Kallo I, Coen CW: Ageing and the diurnal expression of the mRNAs for vasopressin and for the V1a and V1b vasopressin receptors in the suprachiasmatic nucleus of male rats. J Neuroendocrinol 2004;16:493–501. 174 Duncan MJ, Wheeler DL: Aging and photoperiod regulate glutamic acid decarboxylase(67) messenger RNA expression. Brain Res Mol Brain Res 1999;71:325–331. 175 Chee CA, Roozendaal B, Swaab DF, Goudsmit E, Mirmiran M: Vasoactive intestinal polypeptide neuron changes in the senile rat suprachiasmatic nucleus. Neurobiol Aging 1988;9:307–312. 176 Pereira PA, Cardoso A, Paula-Barbosa MM: Nerve growth factor restores the expression of vasopressin and vasoactive intestinal polypeptide in the suprachiasmatic nucleus of aged rats. Brain Res 2005;1048:123–130. 177 Lucassen PJ, Hofman MA, Swaab DF: Increased light intensity prevents the age related loss of vasopressin-expressing neurons in the rat suprachiasmatic nucleus. Brain Res 1995;693:261–266. 178 Zhou JN, Hofman MA, Swaab DF: VIP neurons in the human SCN in relation to sex, age, and Alzheimer’s disease. Neurobiol Aging 1995;16:571–576. 179 Swaab DF, Fliers E, Partiman TS: The suprachiasmatic nucleus of the human brain in relation to sex, age and senile dementia. Brain Res 1985;342:37–44. 180 Van der Zee EA, Jansen K, Gerkema MP: Severe loss of vasopressin-immunoreactive cells in the suprachiasmatic nucleus of aging voles coincides with reduced circadian organization of running wheel activity. Brain Res 1999;816:572–579. 181 Roozendaal B, van Gool WA, Swaab DF, Hoogendijk JE, Mirmiran M: Changes in vasopressin cells of the rat suprachiasmatic nucleus with aging. Brain Res 1987; 409: 259–264. 182 Zhang Y, Brainard GC, Zee PC, Pinto LH, Takahashi JS, Turek FW: Effects of aging on lens transmittance and retinal input to the suprachiasmatic nucleus in golden hamsters. Neurosci Lett 1998;258:167–170. 183 Kornhauser JM, Nelson DE, Mayo KE, Takahashi JS: Photic and circadian regulation of c-fos gene expression in the hamster suprachiasmatic nucleus. Neuron 1990; 5: 127–134. 184 Abe H, Rusak B, Robertson HA: NMDA and non-NMDA receptor antagonists inhibit photic induction of Fos protein in the hamster suprachiasmatic nucleus. Brain Res Bull 1992;28:831–835.

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185 Wollnik F, Brysch W, Uhlmann E, Gillardon F, Bravo R, Zimmermann M, Schlingensiepen KH, Herdegen T: Block of c-Fos and JunB expression by antisense oligonucleotides inhibits light-induced phase shifts of the mammalian circadian clock. Eur J Neurosci 1995;7:388–393. 186 Cai A, Lehman MN, Lloyd JM, Wise PM: Transplantation of fetal suprachiasmatic nuclei into middle-aged rats restores diurnal Fos expression in host. Am J Physiol 1997;272:R422–R428. 187 Sutin EL, Dement WC, Heller HC, Kilduff TS: Light-induced gene expression in the suprachiasmatic nucleus of young and aging rats. Neurobiol Aging 1993; 14: 441– 446. 188 Albers HE, Liou SH, Stopa EG, Zoeller RT: Interaction of colocalized neuropeptides: functional significance in the circadian timing system. J Neurosci 1991; 11: 846– 851. 189 Krajnak K, Lillis TO: Aging alters light- and PACAP-induced cAMP accumulation in the suprachiasmatic nucleus of female rats. Brain Res 2002;950:297–303. 190 Mead S, Ebling FJP, Maywood ES, Humby T, Herbert J, Hastings MH: Nonphotic stimulus causes instantaneous phase advances of the light-entrainable circadian oscillator of the Syrian hamster but does not induce the expression of c-fos in the suprachiasmatic nuclei. J Neurosci 1992; 12: 2516–2522. 191 Cutrera RA, Kalsbeek A, Pevet P: No triazolam-induced expression of Fos protein in raphe nuclei of the male Syrian hamster. Brain Res 1993;602:14–20. 192 Mikkelsen JD, Vrang N, Mrosovsky N: Expression of Fos in the circadian system following nonphotic stimulation. Brain Res Bull 1998;47:367–376. 193 Sahu A, Kalra PS, Crowley WR, Kalra SP: Evidence that hypothalamic neuropeptide Y secretion decreases in aged male rats: implications for reproductive aging. Endocrinology 1988;122:2199–2203. 194 Ruby NF, Edgar DM, Heller HC, Miller JD: The aged suprachiasmatic nucleus is phaseshifted by cAMP in vitro. Brain Res 1998; 779:338–341. 195 van Luijtelaar MG, Tonnaer JA, Steinbusch HW: Aging of the serotonergic system in the rat forebrain: an immunocytochemical and neurochemical study. Neurobiol Aging 1992;13:201–215. 196 van Luijtelaar MGPA, Steinbusch HWM, Tonnaer JA: Similarities between aberrant serotonergic fibers in the aged and 5,7DHT denervated young adult rat brain. Exp Brain Res 1989;78:81–89.

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197 Duncan MJ, Crafton CJ, Wheeler DL: Aging regulates 5-HT1B receptors and serotonin reuptake sites in the suprachiasmatic nucleus and other brain regions. Brain Res 2000;856:213–219. 198 Lovenberg TW, Baron BM, de Lecea L, Miller JD, Prosser RA, Rea MA, Foye PE, Racke M, Slone AL, Siegel BW, Danielson PE, Sutcliffe JG, Erlander MG: A novel adenylyl cyclase-activating serotonin receptor (5-HT7) implicated in the regulation of mammalian circadian rhythms. Neuron 1993;11:449–458. 199 Challet E, Scarbrough K, Penev P, Turek FW: Roles of suprachiasmatic nuclei and intergeniculate leaflets in mediating the phase-shifting effects of a serotonergic agonist and their photic modulation during the subjective day. J Biol Rhythms 1998; 13: 410–421. 200 Ehlen JC, Grossman GH, Glass JD: In vivo resetting of the hamster circadian clock by 5-HT7 receptors in the suprachiasmatic nucleus. J Neurosci 2001;21:5351–5357. 201 Mintz EM, Gillespie CF, Marvel CL, Huhman KL, Albers HE: Serotonergic regulation of circadian rhythms in Syrian hamsters. Neuroscience 1997;79:563–569. 202 Duncan MJ, Short J, Wheeler DL: Comparison of the effects of aging on 5-HT7 and 5-HT1A receptors in discrete regions of the circadian timing system in hamsters. Brain Res 1999;829:39–45. 203 Duncan MJ, Davis VA: Cyclic AMP mediates circadian phase shifts induced by microinjection of serotonergic drugs in the hamster dorsal raphe nucleus. Brain Res 2005;1058:10–16. 204 Byku M, Gannon RL: Effects of the 5-HT1A agonist/antagonist BMY 7378 on light-induced phase advances in hamster circadian activity rhythms during aging. J Biol Rhythms 2000;15:300–305. 205 Ferin M, Tempone A, Zimmering PE, van de Wiele RL: Effect of antibodies to 17-estradiol and progesterone on the estrous cycle of the rat. Endocrinology 1969;85:1070– 1078. 206 Labhsetwar AP: Role of estrogen in spontaneous ovulation: evidence for positive feedback in hamsters. Endocrinology 1972; 90: 941–946. 207 Levine JE, Ramirez VD: Luteinizing hormone-releasing hormone release during the rat estrous cycle and after ovariectomy as estimated with push-pull cannulae. Endocrinology 1982;89:331–337. 208 Jennes L, Stumpf WE: LHRH-systems in the brain of the golden hamster. Cell Tissue Res 1980;209:239–256.

209 Hofman GE, Gibbs FP: LHRH pathways in rat brain: ‘deafferentation’ spares a subchiasmatic LHRH projection to the median eminence. Neuroscience 1982; 7: 1979– 1993. 210 Silverman AJ, Jhamandas J, Renaud LP: Localization of luteinizing hormone-releasing hormone (LHRH) neurons that project to the median eminence. J Neurosci 1987;7: 2312–2319. 211 Lee WS, Smith MS, Hoffman GE: Luteinizing hormone-releasing hormone neurons express Fos protein during the proestrous surge of luteinizing hormone. Proc Natl Acad Sci 1990;87:5163–5167. 212 Everett JW, Sawyer CH: A 24-hour periodicity in the ‘LH-release apparatus’ of female rats, disclosed by barbiturate sedation. Endocrinology 1950;47:198–218. 213 Norman RL, Blake CA, Sawyer CH: Delay of the proestrous ovulatory surge of LH in the hamster by pentobarbital or ether. Endocrinology 1972;91:1025–1029. 214 Stetson MH, Watson-Whitmyre M: The neural clock regulating estrous cyclicity in hamsters: gonadotropin release following barbiturate blockade. Biol Reprod 1977; 16: 536–542. 215 McCormack CE, Sridaran R: Timing of ovulation in rats during exposure to continuous light: evidence for a circadian rhythm of luteinizing hormone secretion. J Endocrinol 1978;76:135–144. 216 Hoorneman EMD, Buijs RM: Vasopressin fiber pathways in the rat brain following suprachiasmatic nucleus lesioning. Brain Res 1982;243:235–241. 217 Funabashi T, Aiba S, Sano A, Shinohara K, Kimura F: Intracerebroventricular injection of arginine-vasopressin V1 receptor antagonist attenuates the surge of luteinizing hormone and prolactin secretion in proestrous rats. Neurosci Lett 1999; 260: 37–40. 218 Palm IF, van der Beek EM, Wiegant VM, Buijs RM, Kalsbeek A: Vasopressin induces a luteinizing hormone surge in ovariectomized, estradiol-treated rats with lesions of the suprachiasmatic nucleus. Neuroscience 1999;93:659–666. 219 Gannon RL, Millan MJ: Serotonin1A autoreceptor activation by S 15535 enhances circadian activity rhythms in hamsters: evaluation of potential interactions with serotonin2A and serotonin2C receptors. Neuroscience 2006;137:287–299. 220 Watts AG: The efferent projections of the suprachiasmatic nucleus: anatomical insights into the control of circadian rhythms; in Klein DC, Moore RY, Reppert SM (eds): Suprachiasmatic Nucleus: The Mind’s Clock. Oxford, Oxford University Press, 1991, pp 77–106.

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Published online: June 6, 2007

Neuroembryol Aging 2006–07;4:102–111 DOI: 10.1159/000103584

Extrapineal Melatonin: General Conception, Localization and Biological Role in the Visual System Igor M. Kvetnoy a Irina O. Smirnova c Victoria O. Polyakova b a

Laboratory of Pathology, Ott Institute of Obstetrics and Gynecology, and b Department of Cell Biology and Pathology, St. Petersburg Institute of Bioregulation and Gerontology, Russian Academy of Medical Sciences, and c Department of Dermatology and Venereology, Faculty of Medicine, St. Petersburg State University, St. Petersburg, Russian Federation

Key Words Diffuse neuroendocrine system
Circadian clock
Extrapineal melatonin
Pineal gland
Retina

though the physiological role of melatonin in the visual system is not exactly determined yet, it apparently includes regulation of gene expression, visual sensitivity and the protection of ocular structures from oxidative damage. Copyright © 2007 S. Karger AG, Basel

Abstract During the last decade, much attention has centered on melatonin, which was considered to be only a hormone of the pineal gland for many years. As soon as highly sensitive antibodies to indolealkylamines became available, melatonin was identified not only in the pineal gland, but also in extrapineal tissues and cells, e.g. the retina, Harderian gland, gut mucosa, cerebellum, airway epithelium, liver, kidney, adrenals, pancreas, thyroid gland and thymus. Also, melatonin has been found in some non-endocrine cells, e.g. mast cells, natural killer cells, eosinophilic leukocytes, platelets and endothelial and other cells. Functionally, melatonin-producing cells are part and parcel of the diffuse neuroendocrine system as a universal system of response, control and organism protection. The influence of aging upon melatonin-synthesizing cells in the gut is summarized. Melatonin synthesis in the visual system and its hormone level, similarly to the pineal gland, reflect the interaction between the circadian clock and the photic environment. Since retinal melatonin does not contribute to circulating levels and melatonin receptors are present in the retina, its effects on the visual system are primarily mediated by a paracrine mechanism. Al-

© 2007 S. Karger AG, Basel 1661–3406/06/0042–0102$23.50/0 Fax +41 61 306 12 34 E-Mail [email protected] www.karger.com

Accessible online at: www.karger.com/nba

Introduction

Melatonin (5-methoxy-N-acetyltryptamine) is the main hormone produced by the pineal gland. In 1958, the American dermatologist Aaron Lerner and his group at Yale University were the first to discover melatonin in the bovine pineal gland [1]. Since their discovery, it was clearly demonstrated that melatonin plays the key role in the regulation of daily and seasonal rhythms, reproductive function, immune response, pigment metabolism and other vitally important physiological processes in all photoperiodic species [2]. Melatonin synthesis depends on the duration of the day and light intensity, reflecting annual photoperiodic changes. Being an endocrine messenger, melatonin quantitatively transfers the photic signal to others tissues, expressing its own receptors, and thus delivers timing information to the organism. The metabolic pathway of biogenic amines in the pineal gland is well investigated. Tryptophan is the precursor of melatonin, which is metabolized consistently into 5-hy-

Prof. Igor M. Kvetnoy Department of Pathology, Ott Institute of Obstetrics and Gynecology RAMS 3 Mendeleevskaia Linia, Vasiljevsky Island St. Petersburg 199034 (Russian Federation) Tel. +7 812 328 98 30, Fax +7 812 328 23 61, E-Mail [email protected]

droxy-tryptophan (by tryptophan-hydroxylase), 5-HT (by aromatic amino acid decarboxylase), N-acetylserotonin (by arylalkylamine-N-acetyltransferase, AA-NAT) and then into melatonin (by hydroxyindole-O-methyltransferase, HIOMT). It has been established that AA-NAT and HIOMT are the key enzymes of this pathway [3]. Melatonin synthesis is mainly observed at night and correlates with the peak of AA-NAT activity. Recent reports indicate that the main factor regulating rhythmic and light-induced changes in AA-NAT activity is the steady-state level of AA-NAT protein, which in turn reflects the balance of its synthesis and degradation [4, 5]. Both of these processes can be regulated by distinct mechanisms and the relative importance of each of them is species dependent [6]. In humans, nocturnal production of melatonin in the pineal gland is mainly regulated by the central circadian clock, situated in the hypothalamic suprachiasmatic nucleus [7]. The circadian clock stimulates norepinephrine release from dense pineal sympathetic fibers. Norepinephrine elevates the intracellular cAMP concentration via
-adrenergic receptors and activates the cAMP-dependent protein kinase A – the crucial pathway for the regulation of AA-NAT synthesis and activity. In some mammals, cAMP/protein kinase A protects the enzyme from degradation [8]. Thus, in primates pinealocytes constantly synthesize AA-NAT from continually available AA-NAT mRNA. During the day, in the absence of noradrenergic stimulation, this protein immediately undergoes proteasomal proteolysis, while nocturnal elevation of the cAMP level causes phosphorylation of AANAT by protein kinase A and protects the enzyme from degradation. Consequent increments in the intracellular concentration of AA-NAT are paralleled by increases in enzyme activity. In rodents, the cAMP/protein kinase pathway induces transcriptional activation of the AANAT gene, the primary mechanism initiating melatonin biosynthesis. In contrast, the importance of a transcriptional regulation of the AA-NAT gene for melatonin production in humans is questionable [9]. Thus, analysis of pineal tissue, taken from regular autopsies, showed significant rhythm for the correlation between time of death and both AA-NAT activity and melatonin content, while AA-NAT and HIOMT mRNA, as well as HIOMT activity showed no diurnal rhythm [9]. Although dominant mechanisms for the generation of rhythmic melatonin synthesis in the human pineal gland remain to be elucidated, it is generally accepted that AA-NAT is switching melatonin synthesis with photoperiodic variation in duration, while the level of HIOMT activity may tune the seasonal magnitude of hormone production [10, 11].

The rhythmical production of melatonin in the pineal gland reliably reflects environmental light conditions – light exposure can acutely suppress nocturnal melatonin secretion [12]. It has been reported that this nonvisual effects of light are mediated via the retinohypothalamic tract, a distinct neural pathway mediating the regulation of melatonin production by light [13, 14], which is initiated in the photoreceptor system. Assumably, light and dark cycles are perceived through the mammalian eyes, entrain the neural activity of the suprachiasmatic nucleus and alter the rhythmic secretion of melatonin from the pineal gland [15]. However, it has been shown recently that rod and cone photoreceptors of the retina are not participating in this pathway [16]. Thus, light at a wavelength of 555 nm, providing a peak effect on the visual system [17], suppresses melatonin synthesis in healthy humans approximately four times less than monochromatic light at 505 nm [15], although the visual ocular photoreceptors mediating these effects remain unknown [15]. Moreover, light-induced melatonin suppression and circadian entrainment have been demonstrated in humans with specific color vision deficiencies [18] or complete visual blindness [19]. The results of these studies suggest that melatonin biosynthesis is controlled, at least in part, by photoreceptors that differ from the known visual photoreceptors [15]. Recent studies on various vertebrate species have identified several new molecules that may serve as circadian photopigments. These pigments include opsin-based molecules (e.g. vertebrate ancient opsin [20], melanopsin [21] and peropsin [22]) as well as non-opsin molecules (bilirubin [23] and cryptochrome [15, 24]). Among these new photopigments only melanopsin has been discovered in the human neural retina [25]. Consequently, the diurnal rhythmical production of melatonin is controlled by strikingly diverse mechanisms apparently characterized by high adaptive plasticityregulatory mechanisms, most of which need to be elucidated.

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Extrapineal Melatonin: General Conception

However, pinealectomy does not abolish the animal’s circadian rhythm in rest-activity though it facilitates resynchronization of the animal to a new photoperiod [26]. In experimental models it has been shown that after pinealectomy the melatonin level at night attenuated, but daytime levels of melatonin in blood remained unaffected, indicating the existence of extrapineal sources of melatonin in the organism [27]. Now it has been securely established that the pineal gland is not the exclusive organ of 103

melatonin synthesis. Extrapineal melatonin is widespread in the organism of humans and animals: melatonin-producing cells are found in the gastrointestinal tract, airway epithelium, pancreas, suprarenal glands, thyroid gland, thymus, urogenital tract, placenta and in other organs. Moreover, active melatonin synthesis in non-endocrine cells has been demonstrated, such as mast cells, natural killer cells, eosinophilic leukocytes, platelets, endotheliocytes and others [28–31]. Today the pineal gland is undoubtedly not the exclusive site of melatonin production. During the last two decades, melatonin synthesis was found in many different organs, tissues and cells: in the gastrointestinal tract [29], in the liver, kidneys, adrenals [30], in lymphocytes, in mast cells, natural killer cells, eosinophilic leukocytes, thymocytes, some endothelial cells, and cells of the placenta and endometrium [31, 32]. The melatonin content in the organism and its blood concentration are accounted for not only by pineal gland secretion, but also by extrapineal sources of synthesis, changes in the volume of extracellular fluid, hormone binding with blood proteins, metabolism and excretion rates depending on different outer and inner regulatory factors. Functionally, cells producing extrapineal melatonin are certain to be part and parcel of the diffuse neuroendocrine system (DNES). In 1968–1969, Pearse [33] was the first to describe the new cell system – the DNES. He undertook an extensive series of experiments on distinguishing endocrine cells in different organs, identifying endocrine cell-generated products and making a thorough cytochemical and ultrastructural analysis of these cells. He suggested that a specialized, highly organized cell system should exist in organisms, whose main feature was the ability of component cells to produce peptide hormones and biogenic amines. He has shown that different types of cells widely dispersed throughout the organism have a common ability of absorbing monoamine precursors (5-hydroxytryptophan and L-dihydroxyphenylalanine) and decarboxylating them, thus producing biogenic amines. That ability accounts for the term APUD, an abbreviation of ‘amine precursor uptake and decarboxylation’ used by Pearse to designate the cell series [34]. The APUD series includes over 60 types of cells located in the gut, pancreas, urogenital tract, airway epithelium, pineal gland, thyroid gland, adrenals, adenohypophysis, hypothalamus, carotid body, skin, sympathetic ganglia, thymus, placenta and other organs [32, 35, 36]. Meanwhile the appearance of radioimmunological methods and rapid development of immunohistochemistry re104

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sult in establishing a completely unexpected phenomenon, i.e. the same biogenic amines and peptide hormones were identified in neurons and endocrine cells. Among APUD cells, cells which produce serotonin, melatonin, catecholamines, histamine, endorphins, endothelin, matrilysin, natriuretic peptide, vasoactive intestinal peptide, neuropeptide Y, vasopressin, oxytocin, somatostatin, endothelin, insulin, substance P and others have been found [37, 38]. Within the whole DNES, two compartments may be distinguished in the melatonin-producing cells, viz. central and peripheral. The central compartment includes the melatonin-producing cells, which are associated with the pineal gland and the visual system (retina, Harderian gland and possibly others) whose secretion rhythm complies with the light-dark rhythm. The peripheral compartment seems to account for all cells located outside the above areas, and its function probably does not depend on the degree of illumination. It includes melatonin-producing cells of the DNES, mainly gut enterochromaffin (EC) cells. Some time later, using reverse transcription-polymerase chain reaction, the two key enzymes of melatonin synthesis have been established in a wide variety of tissues: gut, testis, spinal cord, raphe nuclei, stomach fundus and striatum [39]. The functional morphology of EC cells has been studied sufficiently well [40]. EC cells as well as other melatonin-producing cells can serve as a classic example of APUD cells in which the biogenic amines serotonin and melatonin and peptide hormones (substance P, motilin and enkephalins) coexist [41]. Very important and interestingly, colocalization of melatonin and calcitonin has been obtained in thyroid C-cells; of melatonin and histamine in mast cells; of melatonin, somatostatin and
-endorphins in natural killer cells, and of melatonin and prostaglandin F2 in thymic reticuloepithelial cells [42, 43]. This testifies that peptide hormones and biologically active amines, produced by ‘non-endocrine’ cells in different tissues, are part of a universal system of response, control and organism protection. The hormonal substances of APUD cells act as paracrine signal molecules for the local coordination of intracellular, intertissue and interorgan relationships [44]. The accumulated data did not fit with the traditional concepts of hierarchical dependence within two main regulatory systems, viz. nervous and endocrine ones. It became more and more evident that the mechanism of biological regulation should be founded on the coordinated functional interaction between the endocrine sysKvetnoy /Smirnova /Polyakova

tem and the central and peripheral nervous system based on the common type of information perception and transmission at subcellular, cellular and tissue levels. Recent data on the identification of the same and similar physiologically active substances, acting as neurotransmitters and neurohormones within the nervous system, and, locally or distantly as hormones within the endocrine system, enable the incorporation of both systems into the universal DNES. Actually, it should be possible to unite the structurally isolated nervous and endocrine systems in the organisms by means of functional relationships between biogenic amines and regulatory peptides and, to a certain extent, to provide a basis for the concept of integrated functions [45, 46]. Obviously peripheral ‘non-endocrine’ cells of the DNES take part in immune responses, inflammatory reactions, cell growth and proliferation, and play an important role in the control of normal and pathological processes in the organism. The ascertainment of the morphologic fundamentals of the hormonal function of non-endocrine cells would enable to better understand the intercellular mechanisms of adaptation and compensation of the functional disorders appearing permanently in the organism during the vital activity. Further research into the nature of hormone synthesis and deposition of hormones in non-endocrine cells seems to be very promising. Thus, currently the extrapineal localization of melatonin shows a wide spectrum of cells in the different organs. The list of the cells producing and storing melatonin indicates that melatonin has a unique position among the hormones of the DNES, being found in practically all organ systems. In spite of data showing an active participation of melatonin in adaptive responses, as well as in pathophysiology, the functional significance of the peripheral department of extrapineal melatonin-producing cells remains practically unknown. Showing unique properties as a free radical scavenger, regulator of biological rhythms and cell proliferation, melatonin attracts indefatigable interest of pharmacologists as a potential medicinal substance. The current literature contains a lot of data about successful opportunities for the clinical application of melatonin [47]. Being a most powerful antioxidant, it has been stated that melatonin shows considerable promise in the treatment of Alzheimer’s and Parkinson’s diseases [48, 49]. Administration of low doses of exogenous melatonin can have a positive effect upon the establishment of a normal sleepwake cycle in children with neurological syndromes, mental retardation, blindness and epilepsy [50]. Melatonin/light treatment of patients suffering seasonal affec-

tive disorders and depressive disorders in Northern Europe is widely discussed [51, 52]. The mechanism of action of melatonin on the immune system has been reported [53]. Besides, antitumoral activity of melatonin, especially in combination with immunotherapy, has been demonstrated in clinical and animal model studies [54, 55]. However, in spite of the growing interest in the role of APUD cells in the coordination and regulation of intercellular relationships under normal and pathological processes, studies on the functional role of extrapineal melatonin are limited to oncological and gastroenterological regions. During the last 10 years, our team has studied the functional morphology and behavior of extrapineal melatonin-producing cells as well as other main APUD cells under different pathological and environmental conditions (e.g. ionizing and non-ionizing radiation, tumor growth and cytostatic therapy, autoimmune and gastrointestinal diseases, and pharmacological and toxic effects). The data obtained testify active participation of extrapineal melatonin and other hormones in the pathogenesis of various diseases [46, 56, 57].

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Gastrointestinal Melatonin and Aging

Melatonin is located in all portions of human and animal gut mucosa (from the esophagus to the rectum), where it is synthesized in EC cells [29, 31], which are also producing serotonin, the main precursor of melatonin. EC cells were shown to contain melatonin, as well as the enzyme HIOMT that catalyzes transformation of N-acetylserotonin into melatonin. Similarly to various hormones whose synthesis and presence were revealed in the central nervous system and gut, the effects of melatonin are mediated by humoral, neurocrine, paracrine and autocrine mechanisms. In many studies, involution of the pineal gland as well as a decrease in melatonin blood concentration has been shown during aging [54, 57]. However, the number of melatonin-immunopositive EC cells in human gut mucosa is stable until 60 years, and only after 70 years melatonin expression in the gut is decreasing. It must be noted that at the same time the optical density of melatonin-immunopositive EC cells does not change during aging, confirming that the decrease in melatonin synthesis in the gut after 70 years does not reflect a decreased synthesis in EC cells but a decrease in the total population of EC cells in the gut of elderly people [30, 31]. 105

It is interesting that during aging the behavior of extrapineal gut melatonin-producing EC cells differs between animals and humans. In our study on the functional morphology of stomach melatonin-immunopositive EC cells during aging in BALB/c-nu mice (4, 21 and 34 months old), the number of these cells has a tendency to increase concomitant with age (325 8 52 at 4 months, 345 8 49 at 21 months, and 389 8 46 cells/mm2 at 34 months). No statistically significant differences with age have been found in both the total number of epithelial cell nuclei (6,486 8 92 at 4 months, 6,374 8 95 at 21 months and 6,121 8 55 cells/mm2 at 34 months) and proliferative activity of mucous epitheliocytes (the PCNA index was 15.7 8 0.7 at 4 months, 15.7 8 0.8 at 21 months and 17.6 8 1.1 at 34 months). On the contrary, the apoptotic index rises with advancing age (0.48 8 0.06 at 4 months, 0.47 8 0.06 at 21 months and 0.85 8 0.09 at 34 months) [45]. Many data have proved that the relationships between proliferative cell activity and cell apoptosis play a key role in the progression of aging. Because it is well known at present that melatonin has a strong antiproliferative effect and can trigger apoptosis, we attempted to find the correlations between the functional activity of melatonin-producing stomach EC cells and these biological processes. Our investigations showed an unusual phenomenon: the proliferative activity of epithelial cells does not depend on the activity of EC cells and remained constant with aging, but the index of apoptosis increased simultaneously with advancing age. These data suggest that the level of cell proliferation is sufficiently high to surpass apoptotic elimination, and further investigations are required to clarify the interrelations and the role of melatonin in the endogenous mechanisms of aging. In special experiments we used pinealectomy in Wistar rats as the model of premature aging in order to study the neuroimmunoendocrine interactions and role of extrapineal melatonin under these conditions. For 30 days the increase in the total number of melatonin-immunopositive cells has been assessed in the gut. In the pancreas, the number of glucagon-producing A-cells did not differ compared with the control group, but hyperplasia of somatostatin-immunopositive D-cells and calcitoninimmunoreactive C-cells was noted in the thyroid. After 45 days the number of serotonin- and melatonin-immunopositive cells increased in the gut, but the number and immunoreactivity of islet cells in the pancreas and thyroid C-cells in pinealectomized animals returned to control levels. Thus, these results indicate that pinealectomy causes the changes in neuroimmunoendocrine interac106

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tions in the organism. Extrapineal gut melatonin may function as the key paracrine signal molecule for the local coordination of intercellular relationships in the absence of the pineal gland, a model of premature aging. The data presented here demonstrate that gut melatonin-immunopositive EC cells could play an important role in the endogenous mechanisms of aging.

Melatonin: Localization in the Visual System

The availability of highly sensitive techniques of analysis and identification spurred the detection of melatonin and its precursors as well as catalytic enzymes in extrapineal tissues, primarily those anatomically connected with the visual system – the retina and Harderian gland. Melatonin was localized in the retina immunohistochemically [58, 59]. The fact that pinealectomy did not result in any alterations in retinal melatonin levels may indicate that this hormone synthesis in the retina is independent of the epiphysis [60, 61]. Furthermore, the presence of key enzymes of melatonin biosynthesis (AA-NAT and HIOMT) was shown in retinal tissue, and melatonin synthesis from labeled precursors (tryptophan and serotonin) was demonstrated [62–65]. The comparative study of the capacity of HIOMT of the pineal gland and the retina of the mole to synthesize 5-methoxyindoles demonstrated the similarity of indoleamine metabolism in both tissues [66]. Moreover, both eyes of these mammals, having an atrophied visual system, synthesized 2–30 times more 5-methoxyindoles than the pineal gland [66]. Evidence indicates that melatonin synthesis in the layer of photoreceptor cells is more likely in the cytoplasm of these cells [59, 67, 68]. Several studies have demonstrated that the retinal photoreceptors synthesize melatonin rhythmically and the hormone level reflects the interaction between the circadian clock and the photic environment [13]. It is firmly established now that neurons of the retina contain autonomous circadian clocks [69, 70], regulating different retinal functions and particularly the activity of AA-NAT and melatonin synthesis in retinal photoreceptor cells. In non-mammalian vertebrates, the retinal clock controlling melatonin synthesis is made of photoreceptor cells. Thus, photoreceptor cell cultures derived from chick embryo neural retina contain a complete circadian clockwork system that is entrained by the light-dark cycle, and has a core timekeeping mechanism and circadian output in the form of melatonin synthesis [71]. ExpresKvetnoy /Smirnova /Polyakova

sion of clock genes, Bmal1 and Npas2 (also known as Mop4), in cultured chicken neural retina cells peaked late in the day in light-dark cycle and after transfer to constant darkness. Clock mRNA was high at night in the light-dark cycle, but arrhythmic in constant darkness. Rhythmic expression of clock genes was not observed in cultures not exposed to a light-dark cycle. While the retinal clock controlling melatonin synthesis in mammals has not been definitively localized [72], the persistence of the circadian rhythm in the mRNA levels of AA-NAT in the rat retina after lesioning the suprachiasmatic nucleus is well documented and indicates its relative independence from the master clock in the brain [72, 73]. Investigation of the developmental aspects of the clock system in the mammalian retina has shown that rhythmic changes in AA-NAT mRNA levels occur on postnatal day 2 in rats kept under a daily light-dark cycle, and on postnatal day 20 in animals transferred to constant darkness. Multiple experiments found that biosynthesis of melatonin is relatively independent in cell cultures derived from mammals, and light suppresses AANAT activity and hormone production not only in vivo, but also in vitro [69, 70, 72]. These facts indicate that the cells responsible for these events reside within the eye, i.e. within the rod and cone photoreceptors [69, 70]. The changes in the melatonin levels in the retina [74, 75] and in the pineal gland [5] are a direct reflection of the alterations in the rate of transcription and activity of AA-NAT. First, AA-NAT transcription is directly regulated by the circadian clock via the E-box present in the promoter region of the gene [72]. Second, the photic environment and circadian clock influence AA-NAT transcription via cAMP-responsive elements [13, 72, 75]. The outcome of experimental studies with cultured rat retinas, performed by Fukuhara et al. [13, 75, 76] in 2001, 2002 and 2004, have led to the hypothesis that the circadian clock gates melatonin synthesis by regulating the expression of the type 1 adenylylcyclase (AC1) and the synthesis of cAMP in photoreceptor cells [13]. Darkness depolarizes the photoreceptor plasma membrane, leading to stimulation of cAMP synthesis late in the day and at night, when AC1 protein expression is high, but not during the early morning, when AC1 protein expression is low. An increased cAMP level stimulates AA-NAT transcription and AA-NAT protein synthesis; in addition, it phosphorylates and stabilizes AA-NAT, protecting it from degradation by proteasomes. Thus, AC1 expression provides a gate controlling the time of day when cAMP levels are maximally stimulated by darkness, and melatonin synthesis is stimulated by darkness late in the Extrapineal Melatonin and the Visual System

day and at night, but not during the morning. Light exposure at night induces hyperpolarization of the photoreceptor cell, leading to decreased cAMP level, dephosphorylation and degradation of AA-NAT, and a consequent melatonin decline. Last, several studies have also shown that dopamine plays an important role in the regulation of AA-NAT activity by acting via D2/D4-like receptors that are present on the photoreceptors [72]. The Harderian gland (a special type of intraorbital lacrimal gland) is one of the sources of extrapineal melatonin synthesis [28, 77]. Evidently, melatonin synthesis in the Harderian gland is comparable to that in the epiphysis; at least, one of two key enzymes of melatonin biosynthesis, HIOMT, was found in the Harderian gland [78]. Melatonin synthesis in the Harderian gland of birds and mammals has been shown to comply with circadian rhythm, typical for the epiphysis, but does not depend on it [79, 80]. Moreover, a compensatory increase in the melatonin content has been observed in the Harderian gland of rats some weeks after pinealectomy [60]. The physiological role of melatonin in the Harderian gland remains to be elucidated.

Extrapineal Melatonin and Regulation of Visual System Functions

Melatonin appears to participate in the regulation of retinal physiology and it has been implicated in many retinal functions from gene expression to visual sensitivity [69, 81, 82]. Probably, the effects of melatonin synthesized in the retina are primarily mediated by a paracrine mechanism, while pineal gland melatonin produces changes via humoral and neurocrine pathways. In the retina, hormone function is mediated by melatonin receptors, MT1 and MT2 (formerly Mel1a and Mel1b [83], respectively), presenting in different ocular cell types of humans and other vertebrates. Thus, MT1 and MT2 receptors in horizontal, amacrine, ganglion cell and inner segments of rod photoreceptor cells were found in the human retina immunohistochemically [84]. Although the current literature provides some data of cellular targets of melatonin in the human retina, the functions of the hormone in human vision are still unclear. Thus, in the light of evidences of inhibition of dopamine release by melatonin in the retinas of vertebrates [84–86], the presence of MT1 receptors on dopaminergic amacrine cells in the human retina suggests that it may similarly affect dopaminergic function in humans [84]. The expression of MT1 receptors on horizontal cells in Neuroembryol Aging 2006–07;4:102–111

107

the human retina [84] may suggest the participation of melatonin in light adaptation. Immunohistochemical detection of MT1 on retinal photoreceptors [84] provided the evidence for the direct action of melatonin on these cells, suggesting effects of melatonin on phototransduction processing. In contrast to previous reports ascertaining the prevalence the MT2 receptors in the human retina [87], recent studies indicated similar MT1 and MT2 expression in the human retina, assuming an essential and possibly distinct role for both receptor subtypes in human vision [84]. As melatonin is a potent endogenous free radical scavenger, one of its possible functions in the human visual system is the protection of ocular structures from oxidative damage. In this connection it is necessary to emphasize the high susceptibility of the retina to oxidative stress because of its high consumption of oxygen, high proportion of polyunsaturated fatty acids and exposure to visible light [88]. Current evidence suggests that the pathogenesis of the majority of ocular diseases is connected with oxidative stress [89]. Oxidative damage to the eye leads to apoptotic degeneration of retinal neurons and fluid accumulation [89]. Retinal degeneration decreases visual sensitivity, and even a small change in the fluid content of the cornea and crystalline lens is sufficient to disrupt ocular transparency. Gradual waning of melatonin synthesis in the retina with increasing age increases the risk of free radical-based degenerative eye diseases in the elderly. Moreover, decreases in the melatonin level are accompanied by disturbances in the hormone capacity to control eye pigmentation and consequently regulate the amount of light reaching the photoreceptors [90] that amplify retina damage by light. From this standpoint, melatonin is postulated as a potentially useful agent in the prophylaxis and treatment of free radical-based ocular diseases, e.g. photokeratitis, cataract, glaucoma, retinopathy of prematurity, ischemia/reperfusion injury and retinal pigment epithelium dysfunction [89–91]. In a comparative study of the efficacy of different pineal indoleamines (melatonin, N-acetylserotonin and pinoline) with vitamin E against nitric oxide-induced lipid peroxidation in rat retinal homogenates, vitamin E and indoleamines significantly reduced malondialdehyde levels in a dose-dependent manner [92]. When vitamin E was combined with indoleamines, the protective effect was synergistically enhanced. In animal models, potent effects of melatonin were demonstrated to prevent retinal ischemia-reperfusion injury [93, 94] and experimental uveitis [95, 96] in guinea pigs, and inhibition of the for108

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mation of buthionine sulfoximine (glutathione synthesis inhibitor)-induced cataract in newborn rats [97]. Clinical studies demonstrated that at least 3 months of oral administration of melatonin for patients with agerelated macular degeneration, which is initiated by retinal pigment epithelium dysfunction and a leading cause of blindness in the developed world, evidently reduced pathological macular changes [90]. Continuation of the treatment for 6 months and longer not only helped to stabilize visual acuity but also reduced pathologic macular changes [90].

Conclusion

In conclusion, it is necessary to underline that by now melatonin is found practically in all organs and systems. The list of the cells producing and storing this hormone indicates that melatonin has a unique position among the hormones of the DNES, as a universal system of response, control and organism protection. Taking into account the wide spectrum of biological activities of melatonin and especially its main property as a universal regulator of biological rhythms, it is now possible to consider extrapineal melatonin as a key paracrine signal molecule for the local coordination of intercellular relationships. The diurnal rhythmical production of melatonin by the pineal gland is characterized by high adaptive plasticity-regulatory mechanisms and is controlled in part by photoreceptors in the retina, a structural part of the retinohypothalamic tract. In addition, local synthesis and storage of melatonin by photoreceptors seems to play a crucial role in retinal functions and in the regulation of several aspects of the visual system physiologically and pathologically. Nevertheless, the precise role of melatonin in visual system physiology, the exact mechanisms of the interaction between the circadian clock and the photic environment in hormone production as well as cellular targets and the whole spectrum of melatonin effects on the retina are still far from understanding. It seems that further investigations are required to improve our understanding in this field.

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90 Yi C, Pan X, Yan H, Guo M, Pierpaoli W: Effects of melatonin in age-related macular degeneration. Ann NY Acad Sci 2005; 1057: 384–392. 91 Head KA: Natural therapies for ocular disorders, part two: cataracts and glaucoma. Altern Med Rev 2001;6:141–166. 92 Siu AW, Reiter RJ, To CH: Pineal indoleamines and vitamin E reduce nitric oxide-induced lipid peroxidation in rat retinal homogenates. J Pineal Res 1999;27:122–128. 93 Yilmaz T, Celebi S, Kukner AS: The protective effects of melatonin, vitamin E and octreotide on retinal edema during ischemiareperfusion in the guinea pig retina. Eur J Ophthalmol 2002;12:443–449.

94 Celebi S, Dilsiz N, Yilmaz T, Kukner AS: Effects of melatonin, vitamin E and octreotide on lipid peroxidation during ischemia-reperfusion in the guinea pig retina. Eur J Ophthalmol 2002;12:77–83. 95 Kukner A, Colakoglu N, Serin D, Alagoz G, Celebi S, Kukner AS: Effects of intraperitoneal vitamin E, melatonin and aprotinin on leptin expression in the guinea pig eye during experimental uveitis. Acta Ophthalmol Scand 2006;84:54–61. 96 Kukner AS, Kukner A, Naziroglu M, Colakoglu N, Celebi S, Yilmaz T, Aydemir O: Protective effects of intraperitoneal vitamin C, aprotinin and melatonin administration on retinal edema during experimental uveitis in the guinea pig. Cell Biochem Funct 2004;22: 299–305. 97 Abe M, Reiter RJ, Orhii PB, Hara M, Poeggeler B: Inhibitory effect of melatonin on cataract formation in newborn rats: evidence for an antioxidative role for melatonin. J Pineal Res 1994;17:94–100.

Neuroembryol Aging 2006–07;4:102–111

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Neuroembryol Aging 2006–07;4:112–114 DOI: 10.1159/000103585

Published online: June 6, 2007

Melanopsin in Lower Vertebrates: The Race Is On to Show that It Is Really There V. Benno Meyer-Rochow Faculty of Engineering and Sciences, International University Bremen (now: Jacobs University), Bremen, Germany, and Department of Biological Sciences, Zoological Museum, University of Oulu, Oulu, Finland

Key Words Antarctic
Biorhythm
Bipolar cells
Ganglion cells
Photopigment
Retina
Trematomus bernacchii
Vision

Abstract The retina of the Antarctic teleost Trematomus bernacchii was examined immunocytochemically, using antisera against bovine rhodopsin (1:1,000) and visinin (1:250). There were no surprises with regard to rhodopsin, but positive visinin reactions occurred not only in the cones (visinin has a broad range of sensitivity to opsins), but also in some cells of the ganglion and bipolar cell layers. The reactions could be due to the presence of melanopsin or another opsin-like photopigment, but since experimental work on Antarctic fish is difficult, more accessible fish species should be examined under summer and winter conditions and different light/dark regimens. Furthermore, the search for melanopsin or similar proteins should also involve the pineal organs of lower vertebrates. Copyright © 2007 S. Karger AG, Basel

melanopsin [3], was postulated to explain how some blind mice ‘can still tell day from night’ [4]. Melanopsin was subsequently shown to be housed in certain ganglion cells of the mammalian as well as avian retina [6]. Its three-dimensional structure has recently been predicted from the Djungarian hamster Phodopus sungorus [6]. But what about lower vertebrates or even invertebrates? Do we not have to assume that melanopsin has an evolutionary history? It is hard to believe that in view of the ubiquity of circadian rhythmicity melanopsin should occur only in connection with mammals and birds, and indeed there have been suggestions that it could be present in the inner retinal photoreceptors of teleost fish [7]. There has even been a report that it may occur in the cephalochordate Amphioxus [8]. A couple of years before the identification of melanopsin, we examined immunocytochemically the retina of some Antarctic fishes [9], amongst them those of Trematomus bernacchii, a species caught under 2.5- to 3.0-meter-thick sea ice at a depth of approximately 100 m just above the seafloor. The fish examined weighed about 35 g and measured 14.5 cm in length.

Brief History of Melanopsin Antarctic Experience and Postantarctic Procedures

In 1991, the discovery that mice, which lack retinal rods as well as cones and are unable to perform visual tasks nevertheless responded to photic entrainment of their circadian rhythm [1], led to the prediction that a novel class of photoreceptive cells might exist in the eyes of mammals [2]. A new photosensitive pigment, termed

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The retinae of the lateral eyes and the pineal organs of freshly caught specimens were quickly excised under dim red light, immediately immersed in cold Zamboni’s fixative (2% paraformaldehyde in 0.1 M phosphate buffer containing 0.2% picric acid at pH 7.4) and allowed to fix

V. Benno Meyer-Rochow Faculty of Engineering and Science, Jacobs University Bremen Campus Ring 6, Research II D–28759 Bremen (Germany) Tel. +49 421 200 3242, Fax +49 421 200 3249, E-Mail [email protected]

for 2–3 weeks. After fixation, the tissues were rinsed with phosphate buffer. Some paraffin sections (10
m thick) were prepared from the Zamboni-fixed material and stained with haematoxylin-eosin, while frozen sections, which were prepared with a cryostat, were mounted on gelatin-coated slides and air-dried. The latter were then treated with antisera against bovine rhodopsin (diluted 1:1,000) and visinin (1:250). Not knowing what kind of opsin to expect, visinin was chosen rather than a more specific cone opsin antibody, because visinin has a broader range of sensitivity. The antibody was diluted in phosphate-buffered saline (PBS), containing 1% bovine serum albumin. Sections were incubated overnight at 4 ° C with the respective primary antisera, then rinsed three times with PBS for 10 min each, incubated overnight at 4 ° C with goat anti-rabbit IgG-conjugated FITC (diluted 1:250) in PBS, rinsed with PBS, and mounted using a mixture of glycerol and PBS (1:1). Photomicrographs were taken under a fluorescence microscope.

Rhodopsin and Other Opsins

There were no surprises with regard to the rhodopsincontaining retinal rods, whose outer segments and cell bodies were clearly distinguishable from the non-staining retinal cells (fig. 1a). However, what puzzled us with regard to the reactivity to visinin was that, in addition to the cones, a few scattered cells, chiefly in some regions of the ganglion cell layer and also (but to a lesser extent) amongst the bipolar cells, had reacted positively to the antibody (fig. 1b). In view of what is now known about non-visual photopigment-containing retinal cells like melanopsin-containing ganglion cells in the mammalian retina, for example, it seems possible that the anti-visinin-reactive cells contained small amounts of melanopsin or another opsin-like photopigment. It is noteworthy that the reactive cells were not randomly distributed: certain regions seemed to contain none of them, while there were cell clusters in others. Since melanopsin possesses an absorption maximum of 424 nm and dark/light entrainment appears to be most effective in the blue range of the visual spectrum, the rather low wavelength spectral sensitivity peak of 482–490 nm in Pagothenia borchgrevinki (another Antarctic under-ice sea fish), determined by behavioural methods [10] and electroretinograme recordings [11], could possibly have been the result of a masked contribution by melanopsin. Our anti-visinin immunocytochemistry also identified large numbers of immunoreactive cells in the pineal Melanopsin in Lower Vertebrates

Fig. 1. a The immunoreactive response to rhodopsin clearly shows the presence of retinal rods in the retina of the Antarctic fish T. bernacchii. b Anti-visinin immunoreactivity not only showed the presence of opsin-containing retinal cones, but also identified scattered opsin-containing cells in the ganglion and bipolar cell layers. a, b R = Rod outer segments; C = cone outer segments; ON = outer nuclear layer; OP = outer plexiform layer; IN = inner nuclear/bipolar cell layer; IP = inner plexiform layer; GC = ganglion cell layer. Scale bar = 50
m.

organ of the Antarctic fish, and since immunocytochemical data have been said ‘to indicate that pineal photoreceptive cells share many molecular features with retinal photoreceptors’ [12], the plea is herewith made to also search for melanopsin-containing cells in the visual cells of the pineal eyes of lower vertebrates. After all, ontogenetically the pineal eye is known to be functional before the lateral eyes are active [13]. Neuroembryol Aging 2006–07;4:112–114

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What Next?

Assuming that melanopsin (or another equivalent, but as yet unidentified photopigment) was present in some non-visual retinal cells of the eye of the Antarctic fish T. bernacchii, what could its function be? Would the day/ night fluctuations in ambient light intensities under the sea ice during the Antarctic summer not be absent or unreliable as a zeitgeber? Yes and no. At the latitude at which the T. bernacchii occurs light intensities can be higher at night (due to shifting clouds and snow) than during the day, and consequently it came as no surprise that circadian rhythmicity of the amounts of retinal in the eyes of the Antarctic under-ice amphipod Orchomene plebs was not apparent [14]. However, a 12-hour biorhythm was clearly present in the amount of retinal, but it was rather in synchrony with the day/night oscillations than with the 12-hour intertidal rhythm that prevailed at the time in McMurdo Sound. It was concluded that tidal currents, and in particular the reduced turbulence with less loose platelet ice under the sea ice cover during the turnaround phase, could have provided the amphipods with a more reliable clue than light intensity and position of the sun could have done. The light that does penetrate the layers of snow on top of the sea ice, the sea ice itself and the water column with its ice platelets is heavily blue-shifted when it reaches the eyes of T. bernacchii at the seafloor. This would favor a blue-sensitive pigment like melanopsin present in nonvisual cells like, for instance, some specialized cells of the

inner retina. Light to ‘see’ with and to use for detecting and catching prey may not be sufficient (as had been suggested earlier [10]), but it may still be intensive enough to interact with photopigment-containing ganglia and other non-visual cells in the T. bernacchii retina to entrain the fish and govern some of its biorhythmic activities. If Antarctica were not such an inaccessible place and the fish under debate were readily available, assuming that the antibody of visinin shows specificity to melanopsin, some additional observations could help to determine the contribution of the protein in non-visual cells to fish vision. Immunohistochemistry of T. bernacchii specimens captured at different times of the day and different seasons ought to be conducted to relate protein expression patterns to the day/night and summer/winter oscillations and compared with observations of teleost species that live in other natural habitats with light spectra and photoperiods different from those of the Antarctic. Unfortunately, to carry out research of this nature in Antarctica is still almost impossible (particularly during the winter months) and would require considerable financial and logistic backing.

Acknowledgments The author wishes to thank the Department of Scientific and Industrial Research, Antarctic Division, for the assistance and hospitality received at Scott Base (Antarctica), Dr. J. Macdonald for his help with the ‘fishing’ and Prof. Y. Morita for his collaboration on Antarctic fish eyes.

References 1 Foster RG, Provencio I, Hudson D, Fiske S, DeGrip W, Menaker M: Circadian photoreception in the retinally degenerate mouse (rd/rd). J Comp Physiol A 1991;169:39–50. 2 Brainard GC, Hanifin JM, Greeson B, Byrne B, Glickman G, Gerner E, Rollag MD: Action spectrum for melatonin regulation in humans: evidence for a novel circadian photoreceptor. J Neurosci 2001;21:6405–6412. 3 Provencio I, Jiang G, DeGrip WJ, Hayes WP, Rollag MD: Melanopsin: an opsin in melanophores, brain, and eye. Proc Natl Acad Sci USA 1998;95:340–345. 4 Provencio I, Rollag MD, Castrucci AM: Photoreceptive net in the mammalian retina. Nature 2002;415:493. 5 Hattar S, Liao HW, Takao M, Berson DM, Yau KW: Melanopsin-containing retinal ganglion cells: architecture, projections, and intrinsic photosensitivity. Science 2002;295: 1065–1070.

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6 Hermann R, Poppe L, Pilbak S, Boden C, Maurer J, Weber S, Lerchl A: Predicted 3Dstructure of melanopsin, the non-rod, noncone photopigment of the mammalian circadian clock, from Djungarian hamsters (Phodopus sungorus). Neurosci Lett 2005; 376:76–80. 7 Foster RG, Bellingham J: Inner retinal photoreceptors (IRPs) in mammals and teleost fish. Photochem Photobiol Sci 2004;6:17–27. 8 Koyanagi M, Kubokawa K, Tsukamoto H, Shichida Y, Terakita A: Cephalochordate melanopsin: evolutionary linkage between invertebrate visual cells and vertebrate photosensitive retinal ganglion cells. Curr Biol 2005;15:1065–1069. 9 Meyer-Rochow VB, Morita A, Tamotsu S: Immunocytochemical observations of the pineal organ and retina of the Antarctic teleosts Pagothenia borchgrevinki and Trematomus bernacchii. J Neurocytol 1999; 28: 125– 130.

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10 Pankhurst NW, Montgomery JC: Visual function in four Antarctic nototheniid fishes. J Exp Biol 1989; 142:311–324. 11 Morita Y, Meyer-Rochow VB, Uchida K: Absolute and spectral sensitivities in dark- and light-adapted Pagothenia borchgrevinki, an Antarctic nototheniid fish. Physiol Behav 1997;61:159–163. 12 Meissl H: Photic regulation of pineal function. Biol Cell 1997; 89:549–554. 13 Oksche A: Pineal complex – the third or first eye of vertebrates? A conceptual analysis. Biomed Res 1989;10:187–194. 14 Hariyama T, Terakita A, Meyer-Rochow VB: Rhythmicity of chromophore turnover of visual pigment in the Antarctic amphipod Orchomene plebs (Crustacea; Amphipoda). J Comp Physiol A 1993;173:615–619.

Meyer-Rochow

Author Index Vol. 4, No. 1–2, 2006–07

Chan, C.-K. 76 Chan, C.Y. 8, 13 Chan, S.-O. 76 Do, C.W. 8 Duncan, M.J. 85

Meyer-Rochow, V.B. 112 Nag, T.C. 19 Nishikawa, S. 61 Pang, C.C.P. 13 Polyakova, V.O. 102

Hao, Y. 76 Smirnova, I.O. 102 Kong, C.W. 8 Kunz, Y.W. 31 Kvetnoy, I.M. 102 Lam, C. 8 Lam, D.S.C. 13 Li, W.W.-Y. 13 Lorke, D.E. 5

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To, C.H. 8 Wadhwa, S. 19 Wang, J. 76 Yew, D.T. 13

115

Subject Index Vol. 4, No. 1–2, 2006–07

Aging 5 Antarctic 112 Anterior angle 13 Axon guidance 76 Biorhythm 112 Bipolar cells 112

Development 5, 19 Differentiation 19 Diffuse neuroendocrine system 102 Dorsal raphe nucleus 85

Retina 5, 19, 102, 112 Retinomotor movements 31

Ganglion cells 112 Glaucoma 8

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Neurochemical maturation 19 Optic nerve 5

Fetal eye 13 Fovea development 61

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Melatonin 5 Mouse 76 Mueller cells 61

cGMP 8 Chloride transport 8 Ciliary epithelium 8 Circadian clock 102 – rhythm(s) 31, 85 Collagen fibers 13 Color vision 31 Cone apoptosis 31

Elastic fibers 13 Embryonic fissure 31 Entrainment 85 Extrapineal melatonin 102

© 2007 S. Karger AG, Basel

Intraocular pressure 8

Peripheral growth zones 31 Phase shift 85 Photopigment 112 Photoreceptor disk shedding 31 Pineal gland 102 Preconditioning 61

Serotonin 85 Suprachiasmatic nucleus 85 Trematomus bernacchii 112 Ultraviolet vision 31 Vasoactive intestinal peptide 85 Vision 112 Visual pathway, retina 76