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1From the Lions Eye Institute, The Centre for Ophthalmology and Visual Science, The University of Western Australia, Perth, Australia; the 2Retinal Dystrophy Research Centre, Department of Anatomy and Histology and Institute for Biomedical Research, University of Sydney, Sydney, Australia; and the 3Research School of Biological Science, Australian National University, Canberra, Australia.
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Abstract |
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METHODS. Heterozygote P23H-3 (Line 3) rats were studied. Photoreceptor death rates were assessed with the TUNEL technique for detection of fragmenting DNA, in a developmental series from postnatal day (P)16 to P105 (adult). In adult retinas, trophic factor status was assessed with immunohistochemistry, intraretinal oxygen environment with O2-sensing electrodes, and photoreceptor function by the flash-evoked, dark-adapted electroretinogram (ERG), recorded in anesthetized animals.
RESULTS. Photoreceptor death begins by P16; peaks at P25, when the frequency of TUNEL+ profiles exceeds 70/mm of retina; and then declines to low (<5/mm) adult rates. Compared with that in nondegenerative Sprague-Dawley (SD) rats, the rate of photoreceptor death is abnormally high from P16 and remains several-fold higher than normal into young adulthood. In addition, the outer nuclear layer is reduced to approximately half of control thickness, and the levels of ciliary neurotrophic factor (CNTF), glial fibrillary acidic protein (GFAP), fibroblast growth factor (FGF)-2, and FGF-2/FGFR1 colocalization are markedly upregulated. O2 tension and uptake are relatively normal in the inner retina, but uptake is considerably reduced, and O2 tension is significantly raised in the outer retina. Surviving photoreceptors generate an a-wave with normal peak latency but sharply reduced amplitude.
CONCLUSIONS. Excess photoreceptor degeneration in the P23H-3 retina begins just after eye opening, peaks in early postnatal life, and then slows, but persists into adulthood. In the adult retina, surviving photoreceptors operate in an environment that is chronically hyperoxic (and therefore toxic) and in which protective factors (CNTF, FGF-2) are chronically upregulated. The net result, slow degeneration and degraded function in an environment that is both toxic and protective, may be representative of adult photoreceptor status in a number of human retinal degenerations. Hyperoxia-induced photoreceptor death may be a self-reinforcing factor that increases oxidative stress in surviving photoreceptors.
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Strains and Rearing Conditions
P23H-3 homozygous animals were obtained from the UCSF School of Medicine, Beckman Vision Centre. Those used in the present experiments were heterozygotes, the offspring of mating P23H-3 homozygotes with SD control animals. All rats were born and bred in the University of Sydney animal facility under dim cyclic light (12 hours at <5 lux, 12 hours in the dark). Rats used for intraretinal oxygen measurements were reared until P60 to P80 as above, when they were transferred to the Lions Eye Institute in Perth, where they were kept in brighter conditions (12 hours at 
50 lux, 12 hours in the dark) until they were used for studies of intraretinal oxygen levels. The animals were fed standard laboratory rat chow with water provided ad libitum.
TUNEL Labeling Immunohistochemistry
Eyes were immersion fixed in 4% paraformaldehyde in PBS buffer at pH 7.4 and 4°C for 1 to 3 hours. After three rinses in 0.1 M PBS the eyes were left overnight in a 15% sucrose solution to provide cryoprotection. Eyes were embedded in mounting medium by snap freezing in liquid nitrogen and were cryosectioned at 20 µm. Sections were labeled with the TUNEL technique8 to identify the fragmentation of DNA characteristic of dying cells, after protocols published previously.9 Adjacent sections were labeled with a mouse monoclonal antibody against bovine basic FGF (Type I; Upstate Biotechnology, Lake Placid, NY), with mouse monoclonal antibodies to rod opsin (Rho4D2, a gift from Robert Molday, University of British Columbia, Vancouver, BC, Canada), with a rabbit polyclonal antibody raised against a peptide mapping at the carboxyl terminus of the human Flg (FGFR1) receptor (Santa Cruz Biotechnology Inc., Santa Cruz, CA); with a polyclonal antibody to ciliary neurotrophic factor (CNTF, cat AB 1499P; Chemicon International Inc., Temecula, CA), and/or with a polyclonal antibody to glial fibrillary acidic protein (GFAP; Dako, Carpinteria, CA). Protocols for the use of these have been published previously.1 10 11 To demonstrate the general cellular structure of the retina, many sections were also labeled with the DNA-specific dye bisbenzamide (Calbiochem, La Jolla, CA). Sections were incubated for 2 minutes at room temperature in a 1:10,000 dilution of bisbenzamide in 0.1 M PBS.
Quantifying TUNEL-Labeling, Layer Thickness
Counts of TUNEL+ profiles (apoptotic cells) were made using a calibrated 20x objective and an eyepiece graticule. Each section was scanned from the superior to inferior edge in 400-µm steps, and the number of TUNEL+ profiles was recorded for each 400-µm length of the section. Separate counts were recorded for the INL and ONL. Sections adjacent to those through the optic nerve head were used, to minimize variations in retinal length and position. Counts were averaged from at least two sections per animal.
Oxygen Measurements
Animal Preparation.
Eleven P23H rats aged 15 to 29 weeks, and 11 SD control animals matched for age were used for intraretinal oxygen measurements. The rats were housed two per cage on sawdust. They were fed standard laboratory rat chow with water ad libitum. On the day of the experiment the rat was anesthetized with an intraperitoneal injection of 100 mg/kg 5-ethyl-5- (1'-methyl-propyl)-2-thiobarbiturate (Inactin; Sigma-Aldrich, St. Louis, MO). Atropine sulfate (20 µg) was administered intramuscularly to minimize salivation. The trachea was cannulated for mechanical ventilation, the left internal jugular vein for venous infusion, and the femoral artery for continuous blood pressure monitoring and occasional aspiration of arterial blood (60 µL) for blood gas analysis (CIBA-Corning 238; Corning, NY). The rat was then mounted prone in a modified stereotaxic apparatus and the head fixed in position. The rat was artificially respired (rodent respirator, model 683; Harvard Apparatus, Holliston, MA) with a ventilation rate of 90 breaths per minute and a tidal volume appropriate to ensure normal arterial pCO2 levels. Rectal temperature was monitored and maintained at 37.5°C by a homeothermic blanket (Harvard Apparatus). Experiments usually lasted 8 hours, after which the rat was killed with an anesthetic overdose.
Ocular Surgery.
The left eye was used for all oxygen experiments. The pupil was dilated with 1% tropicamide (Mydriacyl; Alcon Laboratories, French’s Forest, NSW, Australia). The upper eyelid was partially removed, and an eye ring was sutured to the conjunctiva at the limbus and fixed to the stereotaxic framework. A small incision was made in the superior nasal quadrant with a diamond knife, just posterior to the limbus to allow entry of the microelectrode. Damage to the larger choroidal vessels or posterior lens capsule was avoided. A planoconcave contact lens was placed on the cornea to allow the vitreous and the fundus to be visualized with an operating microscope during all intraocular manipulations.
Intraretinal Oxygen Profiles.
The microelectrode techniques were similar to those reported in our earlier publications.12 Recessed oxygen sensitive microelectrodes were manufactured and calibrated in our own laboratory. The microelectrode entered the eye through the entry hole, which was also the locus of rotation of our microsurgical system, such that rotation of the positioning system pivots the electrode about the entry point.13 The small size of the electrode tip (1 µm) coupled with electrode beveling techniques and the high acceleration piezoelectric translation of the electrode produced highly reproducible measurements of intraretinal oxygen distribution. Intraretinal oxygen profiles were measured in the inferior retina, approximately two to three disc diameters from the disc margin. The electrode tip was initially placed at the surface of the chosen area of retina under microscope observation. The electrode was stepped through the retina in 10-µm increments, under computer control, until a peak oxygen level within the choroid was reached. The measurement was repeated during stepwise withdrawal of the electrode. Although very close agreement between the insertion and withdrawal profiles was routinely achieved, the withdrawal profiles were used for data analysis, as they tended to be less influenced by artifacts associated with mechanical stress on the electrode tip during penetration. The oxygen tension measured by the microelectrode and systemic conditions such as arterial blood pressure, were recorded continuously on an eight-channel chart recorder (model LR8100; Yokogawa, Tokyo, Japan). The readings of each channel were also accessed every 2 seconds through a computer interface (GPIB-IEEE) and the data logged directly to a spreadsheet along with the relative position of the microelectrode. All microelectrode measurements were performed in photopic conditions.
Statistics.
All average values for oxygen tension are stated as means ± SE. Significant differences were determined using Student’s t-test, with P < 0.05 accepted as significant. All statistical testing was performed on computer (SigmaStat; SPSS Science, Chicago, IL).
Electroretinogram Recording
Electrophysiological recordings were taken from four SD and four P23H-3 animals at P120. Animals were dark adapted overnight (minimum of 12 hours) and set up under dim red illumination. Anesthesia was achieved with intramuscular injections of ketamine (60 mg/kg) and xylazine (10 mg/kg; Lyppard; Castle Hill, NSW, Australia). Pupils were dilated with 1 drop of tropicamide (Mydriacyl 0.5%; Alcon Laboratories). Corneal hydration was maintained through the duration of recordings with synthetic tears (Viscotears; Carbomer 940 2 mg/g; CIBA Vision, Baulkham Hills, NSW, Australia), which also aided in maintaining electrical contact with the corneal electrode. Body temperature was maintained close to 37°C with an electric blanket controlled by feedback from a rectal temperature probe (Harvard Apparatus). The ERG was recorded between a Pt wire touching the cornea and an Ag/AgCl pellet (Clarke electrode E206; SDR Clinical Technology, Middle Cove, NSW, Australia) in the mouth.
The animal was positioned with the head approximately in the center of a 60-cm diameter Ganzfeld, with the flash source positioned centrally at approximately 45° above the animal’s head. A further 10 minutes of dark adaptation was allowed before commencement of recording. The flash stimulus was provided by a (model 70; Metz GmbH, Zirndorf, Germany) flash unit and flash intensity was attenuated over a 7-log-unit range with neutral density filters. To minimize the cone contribution to the ERG, all stimulus flashes were delivered through a filter (Wratten 47A; Kodak, Rochester, NY) in place. Stimuli were controlled, recorded, and displayed on a computer workstation (MacLab/200 system and Scope software; ADInstruments, Castle Hill, NSW, Australia). Responses were band-pass filtered at 0.3 to 500 Hz. A 50-Hz notch filter was used to minimize mains noise. With attenuations between 1.4 and 7.0 log units, two to three responses were averaged with an interstimulus interval of between 20 (neutral density [ND] 7.0) and 120 (ND 1.4) seconds. At the lowest attenuation (i.e., the brightest flash) a single response was recorded, after an interval of at least 2 minutes from any prior flash.
The flash source was calibrated at the CSIRO National Measurement Laboratory (Lane Cove, NSW, Australia), using a photometer (SD2; Hagner). Conversion from illuminance to irradiance units involved adjustment for the spectral distribution of the flash source (spectral curve provided by Metz), duration of the light pulse (4 ms), and transmissive properties of the filter (Wratten-47; Eastman Kodak). The conversion from flash intensity at the cornea to photoisomerizations per rod per flash (
) was based on previously published methods,14 15 with modifications accounting for the efficiency of photon capture in rodent scotopic vision. The final estimate of flash output (with a Wratten 47 filter) in photoisomerizations per rod per second was 1.09 x 107. In practice, the brightest flash used was with attenuation by the 0.7 neutral density filter (
= 2.18 x 106). This was of sufficient intensity to elicit saturated a-wave responses.
The a-wave amplitude was measured from baseline to the a-wave trough and implicit time (latency) was measured to the trough peak. The b-wave amplitude was determined from a-wave trough to b-wave peak.
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4/mm) but remained several-fold higher than in the SD control (1–2/mm).
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Expression and Sites of Action of Stress-Inducible Factors
In SD control rats, GFAP expression was confined to astrocytes at the inner surface of the retina (Fig. 3A , red). In the P23H-3 retina, GFAP expression was prominent also in the processes of Müller cells, which cross the retina radially (Fig. 3B) . In SD control rats, FGF-2 was prominent only in Müller cells in the INL (Figs. 3A 3C) with some expression in ganglion cell cytoplasm and astrocyte nuclei (Fig. 3C ; for a more detailed description see Ref. 3 ). In the P23H-3 retina FGF-2 expression was markedly upregulated, particularly in the ONL (Figs. 3B 3D 3E 3F 3G) . At higher power (Figs. 3F 3G) the FGF-2 was seen to concentrate in the cytoplasm of photoreceptor somas. In the SD control retina, CNTF was present in astrocytes at the inner surface and was detectable at low levels across all layers of the retina (Fig. 3C ; for more detail see Ref. 3 ). In the P23H-3 retina, CNTF levels were upregulated in Müller cell processes crossing the retina (Figs. 3D 3E) . At higher magnification CNTF was apparent in the cytoplasm of Müller cells in the INL (Figs. 3F 3H) , at the OLM (Figs. 3F 3G , arrows) and in the processes, which extend radially from the inner and outer poles of Müller cell somas in the INL (Fig. 3H) .
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The Oxygen Status of the P23H Retina
Figure 3K is a fundus photograph from a 15-week-old P23H-3 rat, with an oxygen-sensing microelectrode with the tip positioned in the inferior retina. In these animals, the choroidal vessels were much more easily seen than in normal SD rats, a phenomenon presumably related to retinal thinning.16 Typical intraretinal oxygen profiles for a 17-week-old P23H-3 rat are shown in Figure 4B . Data for an age-matched SD control rat are shown in Figure 4A . In the control rat, the intraretinal oxygen distribution reflected the uptake of oxygen in both the inner and outer retina. This was evident in the rapid changes in oxygen gradient at 170- and 330-µm track distances from the retinal surface. In the P23H rat the oxygen-consuming zone in the outer retina was much less evident, and the choroidal peak in oxygen tension was reached at a much reduced track distance, indicating retinal thinning.
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Discussion |
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Hyperoxia in the Degenerating Retina
Models of oxygen flow in the retina5 18 have been developed that relate retinal oxygen tension to oxygen consumption, over a wide range of conditions. When, as in the P23H-3 retina, the population of photoreceptors is depleted, these models assume a corresponding reduction in oxygen consumption by photoreceptors and predict an increase in oxygen tension in the outer retina. This increase goes largely uncompensated because the choriocapillaris autoregulates only poorly (reviewed in Refs. 2 ,19 ). This prediction has now been confirmed in three models of retinal degeneration, the RCS rat,4 the Abyssinian cat (Linsenmeier RA, et al. IOVS 2000;41:ARVO Abstract 1268) and, in the present study, in the P23H-3 rat. It is intriguing that inner retinal oxygen levels and inner retinal oxygen metabolism are largely sustained after photoreceptor degeneration. This was found in the RCS rat4 even after almost complete degeneration of the outer retina. Contrasting results were found in the urethane model of retinal degeneration in which almost all inner retinal oxygen metabolism was lost after photoreceptor degeneration.16
Less direct evidence that the photoreceptor-depleted retina is hyperoxic comes from observations on the status of retinal vessels. Hyperoxia is known to cause the constriction and obliteration of vessels, effects that are particularly marked in the retina because of the poor autoregulation of the choriocapillaris (reviewed in Refs. 19 ,20 ). The mechanisms of vascular thinning relate to the regulation by oxygen of angiogenic factors.21 In the late stages of both human RP22 and rodent models,23 retinal vessels thin, suggesting hyperoxia of the inner retina. Further, it has been shown in the rodent models, though not yet in humans, that hypoxia reverses the depletion-induced thinning of retinal vessels.
Toxicity of Hyperoxia in the Degenerating Retina
The question arises (reviewed in Ref. 2 ) whether chronic outer retinal hyperoxia contributes to the stress to which the depleted retina appears subject. It is well known that lack of oxygen, or anoxia, is a common environmental challenge. However, there is evidence to demonstrate that oxidative stress may not only be caused by hypoxia or anoxia, but also by hyperoxia.24 25 Deviation of oxygen tension above or below normal physiological levels may dramatically change the intracellular redox equilibrium and may alter gene expression patterns in the manifestation of an adaptive stress response. The oxygen consumption of the retina on a per-gram basis has been described as higher than that of the that brain.26 27 Since the brain consumes a highly disproportionate share of the total body’s oxygen uptake,28 the retina is one of the highest oxygen-consuming tissues in the body.5 29 Like the brain, the retina has low levels of the antioxidant enzyme catalase and is rich in iron, which can be a potent catalase for hydroxyl radical formation. These characteristics make the brain and retina particularly sensitive to oxidative stress. The roles of oxidative stress in neurodegenerative diseases such as retinal degeneration should not be underestimated.
In the rabbit,30 and mouse,31 hyperoxia has been shown to be directly and specifically toxic to photoreceptors. It is possible then that depletion-induced hyperoxia is a factor in making many retinal degenerations relentlessly progressive.
It is a clinical feature of human retinal degenerations that many begin with some specificity, affecting rods but not cones or vice versa, and then lose their specificity. This loss of specificity is common, even in degenerations caused by mutations specific to rods, such as the rhodopsin mutations. Depletion-induced hyperoxia could be a factor in this loss of specificity. If confirmed, this would suggest that oxygen management could provide some reduction of stress to depleted retinas, which might slow the rate of progression of the degeneration.
Factors Determining Photoreceptor Death Rates in the Degenerating Retina
The present suggestion that depletion-induced hyperoxia of the ONL makes the late stages of photoreceptor degeneration both progressive and nonspecific also predicts that degenerations should accelerate, because oxygen levels increase as depletion progresses,4 and then should stop suddenly, when the population of photoreceptors is exhausted. Empirically, by contrast, many degenerations slow with age and continue through adult life, as reported herein and previously6 in the P23H transgenic rat. This deceleration may result from the upregulation of protective mechanisms (such as the protective factors FGF-2 and CNTF) by oxidative stress. The rate of photoreceptor death occurring in any retina is presumably determined not simply by the levels of stress experienced, but is the net outcome of the lethal effects of stress and the protective effects of the retina’s defense mechanisms.
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Acknowledgements |
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for achromatic illumination. |
Footnotes |
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Submitted for publication August 6, 2003; revised January 1, 2004; accepted January 5, 2004.
Disclosure: D.-Y. Yu, None; S. Cringle, None; K. Valter, None; N. Walsh, None; D. Lee, None; J. Stone, None
The publication costs of this article were defrayed in part by page charge payment. This article must therefore be marked "advertisement" in accordance with 18 U.S.C. 
1734 solely to indicate this fact.
Corresponding author: Jonathan Stone, Research School of Biological Sciences, The Australian National University, GPO Box 475, Canberra City, ACT 2601, Australia; stone{at}rsbs.anu.edu.au.
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