|Molecular Vision 1999;
Received 24 May 1999 | Accepted 2 November 1999 | Published 3 November 1999
Senescence of the retinal pigment epithelium
Hjelmeland,1 Vincent J. Cristofalo,2 Walter Funk,3
Elizabeth Rakoczy,4 Martin L. Katz5
1Department of Ophthalmology, School of Medicine, University of California, Davis, CA; 2Center for Gerontological Research, Allegheny University of the Health Sciences, Medical College of Pennsylvania, Hahnemann School of Medicine, Philadelphia, PA; 3Geron Corporation, Menlo Park, CA; 4Centre of Ophthalmology and Visual Science, University of Western Australia; 5University of Missouri School of Medicine, Mason Eye Institute, Columbia, MO 65212
Correspondence to: Leonard M. Hjelmeland, Ph.D., Vitreoretinal Research Laboratory, School of Medicine, University of California, One Shields Avenue,Davis, CA, 95616-8794; Phone: (530) 752-2250; FAX: (530) 752-2270; email: email@example.com
Senescence of human cells has largely been studied as an in vitro phenomenon resulting from replicative exhaustion. The literature contains many studies of retinal pigment epithelium (RPE) cells which document replicative senescence. Several studies by Burke and others illustrate the relationship between donor age and replicative lifespan, the relationship between geographical location of RPE in the posterior pole and replicative lifespan, and the phenomena of altered cellular morphology and decreased culture saturation density for senescent RPE cells. Other studies have focused on the alterations of the expression of specific genes or the alteration of enzymatic activities during the senescence of RPE cells in vitro. Recently, a technique utilizing a histochemical staining procedure for beta galactosidase has been developed which identifies senescent cells. Normal beta galactosidase histochemistry which identifies the lysosomal form of the enzyme is performed at pH 4.0, while senescence-associated beta galactosidase activity is observed at pH 6.0 and is observed in the cytoplasm. We have studied the replicative senescence of human RPE cells in vitro using this procedure and have also measured the length of chromosomal telomeres to identify the aging of cultures in vitro. Our results show that RPE cultures accumulate beta galactosidase positive cells as a function of the number of population doublings and that these data correlate with the shortening of chromosomal telomeres to a functional limit observed for many human cell types at senescence. We have also recently extended this work to the development of a senescence-associated beta galactosidase procedure for observing senescent RPE cells in vivo. Basically, the same histochemical procedure is used with a post-staining bleaching step to clearly visualize staining in the RPE. Our first studies were performed on globes from Rhesus monkeys at a variety of ages from 1 year to 29 years of age. The results show the accumulation of beta galactosidase positive cells in the older monkey eyes. We have also examined several human eyes in an attempt to observe whether any relationship exists between beta galactosidase staining and age, pathology (diabetes, basal laminar deposits), and geographical location (macula vrs. periphery). These studies represent a first effort to determine if senescent RPE are present in vivo. It will be important to extend these studies so that these data might be expressed on a quantitative bases.
Senescence of human cells has largely been studied as an in vitro phenomenon resulting from replicative exhaustion [1-10]. The literature contains several reports which document the senescence of retinal pigment epithelium (RPE) cells [11-18]. Studies by Burke and others illustrate the relationship between donor age and replicative lifespan, the relationship between topographical location of RPE in the posterior pole and replicative lifespan, and the phenomena of altered cellular morphology and decreased culture saturation density for senescent RPE cells. Other studies have focused on the alterations of the expression of specific genes or the alteration of enzymatic activities during the senescence of RPE cells in vitro [19-21].
What evidence suggests that senescent RPE cells exist in vivo?
Recently, a technique utilizing a histochemical staining procedure for beta galactosidase has been developed which identifies senescent cells . Normal beta galactosidase histochemistry which identifies the lysosomal form of the enzyme is performed at pH 4.0, while senescence-associated beta galactosidase activity is observed at pH 6.0 and is observed in the cytoplasm. A recent study in the literature has shown that human RPE cells at replicative senescence not only stain for senescence-associated beta galactosidase but also have critically shortened chromosomal telomeres . Withdrawal of senescent RPE cells from the cell cycle was documented by BrdU labeling. These results show that RPE cultures accumulate beta galactosidase positive cells as a function of the number of population doublings and that these data correlate with the shortening of chromosomal telomeres to a functional limit observed for many human cell types at senescence.
The senescence-associated beta galactosidase histochemistry procedure has also been recently modified for studies of the RPE cell in vivo . Basically, the same histochemical procedure was used with a post-staining bleaching step to clearly visualize staining in the RPE. The first studies were performed on globes from Rhesus monkeys at a variety of ages from 1 year to 29 years of age. The results show the accumulation of beta galactosidase positive cells in the older monkey eyes.
These studies represent a first effort to determine if senescent RPE are present in vivo. It will be important to extend these studies to the human eye with a quantitative treatment of data.
What is the function of the downregulation of EPC-1 at replicative senescence?
Replicative senescence is also characterized by changes in gene expression and dramatic phenotypic alterations [25,26]. The most profound of these is the loss of a proliferative response to mitogens, but alterations in expression of genes for matrix proteins, extracellular proteases, and inflammatory cytokines has also been documented.. Senescent growth arrest is distinct from the quiescent or G0 state induced in early passage cells by serum deprivation or contact inhibition. The phenotypic alterations which human fibroblasts exhibit at replicative senescence have been used to study alterations in the expression of specific genes. The expression of one of these genes, EPC-1, is detected as a 1.4 kb transcript in young (early passage), serum-deprived WI-38 cells . The EPC-1 gene product, a 50 kD secreted protein, is identical to the retinal pigmented epithelium derived factor (PEDF) produced by retinal pigmented cells and exhibits approximately 30% sequence identity to serpins, a family of secreted mammalian serine protease inhibitors.
Not only is EPC-1 G0-specific and potentially inhibitory for cellular proliferation, but EPC-1 expression is gradually lost during the fibroblast proliferative lifespan. Two questions of significance are: (1) What is the molecular mechanism by which EPC-1 slows/blocks cell cycle progression? and (2) What is the consequence of losing EPC-1 expression in senescent cells and can the ability to enter G0 be restored by overexpressing EPC-1 in these cells?
EPC-1 may also have growth inhibitory, and perhaps tumor suppressive, effects. Several findings support this view: (1) The number of WI-38 cells entering S phase can be increased by the addition of anti-EPC-1 antibodies to the culture media; (2) EPC-1 induces neuronal differentiation of Y79 retinoblastoma cells; (3) EPC-1 inhibits proliferation of endometrial carcinoma cells; and (4) The recombinant EPC-1 protein reduces the number of growing cells entering S phase by approximately 50%. The engineered overexpression of EPC-1 in mouse fibroblasts also results in a significant reduction in proliferation of the cells.
What is the mRNA phenotype of senescent cells?
Recently, studies of senescent gene expression patterns in multiple cell types and in response to various stimuli have been conducted by microarray analysis [27,28]. A custom DNA microarray was generated by Geron Corporation in collaboration with Synteni and was subsequently assessed for sensitivity and reproduciblity. mRNA was produced from early passage or senescent primary diploid cell lines, converted to fluorescently labeled probes, and hybridized to the array. Relative mRNA abundance was calculated after balancing signals from internal and exogenous controls. A comparison of senescence-associated expression patterns was generated for retinal pigment epithelial cells, dermal fibroblasts and vascular endothelial cells. Commonly dysregulated genes were identified, as well as those specific for a given cell type. A comparison of hyperoxia-induced senescence reveals partial overlap with replicative exhaustion. Senescence induces gene expression patterns that are shared partially amongst different cell types and may provide markers such as IGFBP-2 that can be assessed in aging or diseased tissues.
What is the possible role of RPE lysosomes in the formation of autoflourescent debris and the pathogenesis of AMD?
One of the most important roles of the RPE is the lysosomal digestion of the continuously growing photoreceptor outer segments. A decrease in lysosomal enzyme activity has been previously proposed to accompany the general aging process and it has been suggested that an impairment of RPE lysosomal enzymatic activity might be linked to the development of AMD [29-37]. Enzymatic studies could not reliably establish any relationship between aging and a decrease in enzymatic activity.
Of the lysosomal enzymes, an aspartic protease, cathepsin D (CatD), appears to be the most important enzyme for the proteolysis of opsin. The molecular and enzymatic characteristics of CatD are well understood, including the enzymatic or autolytic conversion of the proenzyme (52 kD) into the single chain active form of CatD (34 kD). CatD is present in a wide range of retinal cells with particularly high levels of expression present in the retinal pigment epithelium . Studies on the modulation of CatD activity in RPE cells have been conducted using retroviral transduction with both sense and antisense constructs . To monitor the lysosomal processing of ROS, the proteolysis of the major protein opsin was followed by Western blot analysis. In vitro transduction of RPE cells with the antisense recombinant virus (Ad.MLP.CatDAS) resulted in an inhibition of opsin proteolysis within the RPE cells. Subretinal injections of the recombinant adenovirus, carrying CatD in sense orientation did not induce any noticeable retinal changes during the course of the experiments (2 weeks). In contrast, subretinal injection of the recombinant adenovirus caused dramatic changes. The RPE cells were enlarged and they were full of debris which was autofluorescent. In addition, in large areas the degeneration of the photoreceptor layer was observed and by 2 weeks only a few layers of photoreceptors were left.
What is the mechanism of the formation of lipofuscin in RPE cells?
The progressive accumulation of autofluorescent lysosomal storage material (lipofuscin) is characteristic of senescent RPE cells in vivo [38-55]. At present, a role for RPE lipofuscin accumulation in AMD remains to be proven. An important step toward evaluating a possible association between RPE lipofuscin accumulation and AMD is the characterization of the molecular and cellular mechanisms by which RPE lipofuscin forms. A variety of studies in the literature have been published concerning the chemical composition and formation of lipfuscin in vitro and in vivo. A possible role for lipofuscin in the pathogenesis of AMD as a photooxidative sensitizer has also been published.
Cellular senescence, the alterations of lysosomal function, and the role of aging pigments as sensitizers for photooxidative stress all appear to be fruitful avenues for exploration. The following questions represent issues which may help to determine how related these studies are and in what ways each may be involved in the pathogenesis of AMD.
1. What evidence exists for the presence of senescent RPE cells in vivo? Does the distribution of senescent RPE cells in vivo vary as a function of age and posterior pole topography in a fashion consistent with a role in the pathogenesis of AMD?
2. In what ways does cellular senescence alter the function of lysosomes?
3. Does senescence alter the RPE cell's ability to respond to oxidative stress or to synthesize and secrete trophic factors for neighboring cell types?
4. Does oxidative stress (or photooxidative stress) represent a viable mechanism for the induction of RPE senescence in vivo? Given that RPE cells are thought to be postmitotic after birth, what other mechanisms could account for the apparent decrease of proliferative potential observed for RPE cells in the macula versus the periphery of the human eye?
5. In what ways are senescence, oxidative stress, the presence of aging pigments, and the alteration of lysosomal function related? Can cause and effect be determined among these aspects of the aging RPE cell?
6. Would RPE senescence represent a cause or a contributing factor for AMD? How would the role of RPE senescence in age related maculopathy (ARM) be related to or distinct from its role in AMD?
Supported by NIH grant EYO6473 and an unrestricted grant from Research to Prevent Blindness, Inc. L. M. Hjelmeland is the recipient of a Senior Scientist Award from Research to Prevent Blindness, Inc.
1. Hayflick L, Moorhead PS. The serial cultivation of human diploid cell strains. Exp Cell Res 1961; 25:585-621.
2. Hayflick L. The limited in vitro lifetime of human diploid cell strains. Exp Cell Res 1965; 37:614-36.
3. Martin GM, Sprague CA, Epstein CJ. Replicative life-span of cultivated human cells. Effects of donor's age, tissue, and genotype. Lab Invest 1970; 23:86-92.
4. Schneider EL, Mitsui Y. The relationship between in vitro cellular aging and in vivo human age. Proc Natl Acad Sci U S A 1976; 73:3584-8.
5. West MD. The cellular and molecular biology of skin aging. Arch Dermatol 1994; 130:87-95.
6. Bladier C, Wolvetang EJ, Hutchinson P, de Haan JB, Kola I. Response of a primary human fibroblast cell line to H2O2: senescence-like growth arrest or apoptosis? Cell Growth Differ 1997; 8:589-98.
7. Chen Q, Ames BN. peroxide in human diploid fibroblast F65 cells. Senescence-like growth arrest induced by hydrogen peroxide in human diploid fibroblast F65 cells. Proc Natl Acad Sci U S A 1994; 91:4130-4.
8. Serrano M, Lin AW, McCurrach ME, Beach D, Lowe SW. Oncogenic ras provokes premature cell senescence associated with accumulation of p53 and p16INK4a. Cell 1997; 88:593-602.
9. Saito H, Hammond A, Moses R. The effect of low oxygen tension on the in vitro-replicative life span of human diploid fibroblast cells and their transformed derivatives. Exp Eye Res 1995; 217:272-9.
10. von Zglinicki T, Saretzki G, Docke W, Lotze C. Mild hyperoxia shortens telomeres and inhibits proliferation of fibroblasts: a model for senescence? Exp Eye Res 1995; 220:186-93.
11. Burke JM. Cytochrome oxidase activity in bovine and human retinal pigment epithelium: topographical and age-related differences. Curr Eye Res 1993; 12:1073-9.
12. Burke JM, McKay BS. In vitro aging of bovine and human retinal pigment epithelium: number and activity of the Na/K ATPase pump. Exp Eye Res 1993; 57:51-7.
13. Burke JM, Soref C. Topographical variation in growth in cultured bovine retinal pigment epithelium. Invest Ophthalmol Vis Sci 1988; 29:1784-8.
14. Del Monte MA, Maumenee IH. In vitro culture of human retinal pigment epithelium for biochemical and metabolic study. Vision Res 1981; 21:137-42.
15. Flood MT, Gouras P, Kjeldbye H. Growth characteristics and ultrastructure of human retinal pigment epithelium in vitro. Invest Ophthalmol Vis Sci 1980; 19:1309-20.
16. Newsome DA. Retinal pigmented epithelium culture: current applications. Trans Ophthalmol Soc U K 1983; 103:458-66.
17. Sheedlo HJ, Wordinger RJ, Fan W, Turner JE. A transformed neonatal rat retinal pigment epithelial cell line: secreted protein analysis and fibroblast growth factor and receptor expression. Curr Eye Res 1997; 16:116-26.
18. Song MK, Lui GM. Propagation of fetal human RPE cells: preservation of original culture morphology after serial passage. J Cell Physiol 1990; 143:196-203.
19. Guillonneau X, Tassin J, Berrou E, Bryckaert M, Courtois Y, Mascarelli F. In vitro changes in plasma membrane heparan sulfate proteoglycans and in perlecan expression participate in the regulation of fibroblast growth factor 2 mitogenic activity. J Cell Physiol 1996; 166:170-87.
20. Tombran-Tink J, Shivaram SM, Chader GJ, Johnson LV, Bok D. Expression, secretion, and age-related downregulation of pigment epithelium-derived factor, a serpin with neurotrophic activity. J Neurosci 1995; 15:4992-5003.
21. Wyszynski RE, Bruner WE, Cano DB, Morgan KM, Davis CB, Sternberg P. A donor-age-dependent change in the activity of alpha-mannosidase in human cultured RPE cells. Invest Ophthalmol Vis Sci 1989; 30:2341-7.
22. Dimri GP, Lee X, Basile G, Acosta M, Scott G, Roskelley C, Medrano EE, Linskens M, Rubelj I, Pereira-Smith O, et al. A biomarker that identifies senescent human cells in culture and in aging skin in vivo. Proc Natl Acad Sci U S A 1995; 92:9363-7.
23. Matsunaga H, Handa JT, Aotaki-Keen A, Sherwood S, West MD, Hjelmeland LM. Beta-galactosidase histochemistry and telomere loss in senescent retinal pigment epithelial cells. Invest Ophthalmol Vis Sci 1999; 40:197-202.
24. Mishima K, Aotaki-Keen AE, Lutty GA, Handa JT, Mores LS, Hjelmeland LM. Histochemical staining of the rhesus retina for senescence related beta-galactosidase. Invest Ophthalmol Vis Sci 1998; 39:S827.
25. Cristofalo VJ, Pignolo RJ. Molecular markers of sensecence in fibroblast-like cultures. Exp Gerontol 1996; 31:111-23.
26. Linskens MH, Feng J, Andrews WH, Enlow BE, Saati SM, Tonkin LA, Funk WD, Villeponteau B. Cataloging altered gene expression in young and senescent cells using enhanced differential display. Nucleic Acids Res 1995; 23:3244-51.
27. Brown PO, Hartwell L. Genomics and human disease--variations on variation. Nat Genet 1998; 18:91-3.
28. Schena M, Shalon D, Davis RW, Brown PO. Quantitative monitoring of gene experession patterns with a complementary DNA microarray. Science 1995; 270:467-70.
29. Boulton M, Moriarty P, Jarvis-Evans J, Marcyniuk B. Regional variation and age-related changes of lysosomal enzymes in the human retinal pigment epithelium. Br J Ophthalmol 1994; 78:125-9.
30. Cingle KA, Kalski RS, Bruner WE, O'Brien CM, Erhard P, Wyszynski RE. Age-related changes of glycosidases in human retinal pigment epithelium. Curr Eye Res 1996; 15:433-8.
31. el-Hifnawi E, Kuhnel W, el-Hifnawi A, Laqua H. Localization of lysosomal enzymes in the retina and retinal pigment epithelium of RCS rats. Anat Anz 1994; 176:505-13.
32. Hayasaka S. Lysosomal enzymes in ocular tissues and diseases. Surv Ophthalmol 1983; 27:245-58.
33. Rakoczy PE, Baines M, Kennedy CJ, Constable IJ. Correlation between autofluorescent debris accumulation and the presence of partially processed forms of cathepsin Din cultured retinal pigment epithelial cells challenged with rod outer segments. Exp Eye Res 1996; 63:159-67.
34. Rakoczy PE, Lai CM, Baines M, Di Grandi S, Fitton JH, Constable IJ. Modulation of cathepsin D activity in retinal pigment epithelial cells. Biochem J 1997; 324:935-40.
35. Wiederanders B, Oelke B. Accumulation of inactive cathepsin D in old rats. Mech Ageing Dev 1984; 24:265-71.
36. Wilcox DK. Vectorial accumulation of cathepsin D in retinal pigmented epithelium: effects of age. Invest Ophthalmol Vis Sci 1988; 29:1205-12.
37. Yamada T, Hara S, Tamai M. Immunohistochemical localization of cathepsin D in ocular tissues. Invest Ophthalmol Vis Sci 1990; 31:1217-23.
38. Boulton M, Dontsov A, Jarvis-Evans J, Ostrovsky M, Svistunenko D. Lipofuscin is a photoinducible free radical generator. J Photochem Photobiol B 1993; 19:201-4.
39. Dorey CK, Wu G, Ebenstein D, Garsd A, Weiter JJ. Cell loss in the aging retina. Relationship to lipofuscin accumulation and macular degeneration. Invest Ophthalmol Vis Sci 1989; 30:1691-9.
40. Eldred GE, Lasky MR. Retinal age pigments generated by self-assembling lysosomotropic detergents. Nature 1993; 361:724-6.
41. Feeney-Burns L, Eldred GE. The fate of the phagosome: conversion to 'age-pigment' and impact in human retinal pigment epithelium. Trans Ophthalmol Soc U K 1983; 103:416-21.
42. Feeney-Burns L, Hilderbrand ES, Eldridge S. Aging human RPE: morphometric analysis of macular, equatorial, and peripheral cells. Invest Ophthalmol Vis Sci 1984; 25:195-200.
43. Katz ML, Parker KR, Handelman GJ, Bramel TL, Dratz EA. Effects of antioxidant nutrient deficiency on the retina and retinal pigment epithelium of albino rats: a light and electron microscopic study. Exp Eye Res 1982; 34:339-69.
44. Katz ML, Robison WG Jr. Age-related changes in the retinal pigment epithelium of pigmented rats. Exp Eye Res 1984; 38:137-151.
45. Katz ML, Robison WG Jr, Dratz EA. Potential role of autoxidation in age changes of the retina and retinal pigment epithelium of the eye. In: Armstrong D, Sohal RS, Cutler RG, Slater TF, editors. Free radicals in molecular biology, aging, and disease. New York: Raven Press. 1984; p. 63-180.
46. Katz ML, Robison WG Jr. Nutritional influences on autoxidation, lipofuscin accumulation, and aging. In: Johnson JE Jr, Walford R, Harmon D, Miquel J, editors. Free radicals, aging, and degenerative diseases. New York: Liss; 1986. p. 221-59.
47. Katz ML, Drea CM, Eldred GE, Hess HH, Robison WG Jr. Influence of early photoreceptor degeneration on lipofuscin in the retinal pigment epithelium. Exp Eye Res 1986; 43:561-73.
48. Katz ML, Robison WG Jr, Drea CM. Factors influencing lipofuscin accumulation in the retinal pigment epithelium of the eye. In: Totaro EA, Glees P, Pisanti FA, editors. Advances in age pigments research: proceedings of the First International Workshop on "Age Pigments, Biological Markers in Aging and Environmental Stress?"; 1985 May 29-Jun 1; Vico Equense (Napoli), Italy. Oxford: Pergamon Press; 1987. p. 111-31.
49. Eldred GE, Katz ML. Fluorophores of the human retinal pigment epithelium: separation and spectral characterization. Exp Eye Res 1988; 47:71-86.
50. Katz ML, Eldred GE. Failure of vitamin E to protect the retina against damage resulting from bright cyclic light exposure. Invest Ophthalmol Vis Sci 1989; 30:29-36.
51. Eldred GE, Katz ML The autofluorescent products of lipid peroxidation may not be lipofuscin-like. Free Radic Biol Med 1989; 7:157-63.
52. Katz ML, Robison WG Jr. Senescent alterations in the retina and retinal pigment epithelium: evidence for mechanisms based on nutritional studies. In: Armstrong D, Marmor MF, Ordy JM, editors. The effects of aging and environment on vision. New York: Plenum Press; 1991. p. 195-208.
53. Katz ML. Age-related alterations in the retina. In: Lutjen-Drecoll E, editor. Basic Aspects of Glaucoma Research III: international symposium held at the Department of Anatomy, University of Erlangen-Nürnberg, FRG, September 23-25, 1991. Stuttgart: F. K. Schattauer Verlag; 1993. p. 133-152.
54. Porta EA. Advances in age pigment research. Arch Gerontol Geriatr 1991; 12:303-20.
55. Robison WG Jr, Kuwabara T, Bieri JG. Deficiencies of vitamins E and A in the rat. Retinal damage and lipofuscin accumulation. Invest Ophthalmol Vis Sci 1980; 19:1030-7.