May 2009
Volume 50, Issue 5
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Visual Neuroscience  |   May 2009
Functional and Structural Changes Resulting from Strain Differences in the Rat Model of Oxygen-Induced Retinopathy
Author Affiliations
  • Allison Lindsay Dorfman
    From the Departments of Pharmacology and Therapeutics and
    Ophthalmology/Neurology–Neurosurgery, McGill University-Montreal Children’s Hospital Research Institute, Montreal, Quebec, Canada; and the
  • Anna Polosa
    Ophthalmology/Neurology–Neurosurgery, McGill University-Montreal Children’s Hospital Research Institute, Montreal, Quebec, Canada; and the
  • Sandrine Joly
    Departments of Biology,
  • Sylvain Chemtob
    Ophthalmology/Neurology–Neurosurgery, McGill University-Montreal Children’s Hospital Research Institute, Montreal, Quebec, Canada; and the
    Pediatrics,
    Ophthalmology, and
    Pharmacology, University of Montreal, Montreal Quebec, Canada.
  • Pierre Lachapelle
    Ophthalmology/Neurology–Neurosurgery, McGill University-Montreal Children’s Hospital Research Institute, Montreal, Quebec, Canada; and the
Investigative Ophthalmology & Visual Science May 2009, Vol.50, 2436-2450. doi:10.1167/iovs.08-2297
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      Allison Lindsay Dorfman, Anna Polosa, Sandrine Joly, Sylvain Chemtob, Pierre Lachapelle; Functional and Structural Changes Resulting from Strain Differences in the Rat Model of Oxygen-Induced Retinopathy. Invest. Ophthalmol. Vis. Sci. 2009;50(5):2436-2450. doi: 10.1167/iovs.08-2297.

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      © ARVO (1962-2015); The Authors (2016-present)

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Abstract

purpose. Results of studies that compared the racial incidence of retinopathy of prematurity (ROP) suggested that ocular pigmentation might offer protection against the development of severe ROP. The structural and functional consequences of postnatal hyperoxia (oxygen-induced retinopathy; OIR) were compared in albino Sprague-Dawley (SD) and pigmented Long-Evans (LE) rats to verify this finding.

methods. Newborn rats were exposed to 80% O2 during selected postnatal day intervals. The severity of the OIR was determined by examining retinal flatmounts (retinal vasculature assessment), protein level quantification and cellular localization of fibroblast growth factor (FGF)-2 and ciliary neurotrophic factor (CNTF; Western blot analysis and immunohistochemistry, respectively), retinal histology, and photopic and scotopic electroretinograms (ERGs).

results. Irrespective of the parameter considered, structural and functional deficits resulting from postnatal hyperoxia were significantly more pronounced in LE rats. Although FGF-2 levels in LE rats had a tendency to increase after hyperoxia compared with normoxic littermates, it did not reach statistical significance. A similar finding was observed in SD rats. Of interest, however, baseline levels of FGF-2 were approximately four to five times higher in SD rats than in LE rats. There was a similar, hyperoxia-induced increase in CNTF levels between SD and LE rats.

conclusions. The findings suggest an increased susceptibility of newborn LE rats to postnatal hyperoxia in comparison with SD rats. Whether a pro-oxidant rather than antioxidant role of melanin or other genetic factors can explain these differences in oxygen susceptibility of the animal model of this retinopathy, remains to be determined.

Despite many advances in the clinical monitoring of oxygen administration in neonatal intensive care units, retinopathy of prematurity (ROP) continues to be a major cause of morbidity affecting prematurely born infants. Similar to other types of proliferative retinopathies, such as diabetic retinopathy and sickle cell retinopathy, ROP is characterized by retinal neovascularization. 1 2 3 4 It is this abnormal growth of new retinal vessels that can lead to the most devastating consequences of the disease, namely retinal detachment and eventually blindness. Through years of research, it has become increasingly evident that ROP is a very complex, multifactorial disease. Some of the risk factors known to be associated with the incidence of ROP include low birthweight and low gestational age 5 6 7 and, notably, relative hyperoxia (compared with oxygen tension in utero). 1 Other studies have also shown that race is a significant factor that influences the epidemiology of ROP 8 9 ; for example, a greater number of Caucasian than African-American infants have been shown to reach threshold ROP. 8 10 These studies suggest that melanin pigmentation offers protection against the development of severe ROP, suggesting that fundus pigmentation may play an instrumental role in this protection. 6 8 On the other hand, other studies have suggested that an even greater incidence in the onset of threshold ROP exists in another darkly pigmented race, namely native Alaskans, when compared with that in other racial groups. 11 12 Despite the fact that several of these studies have controlled for birth weight, gestational age, length of oxygen therapy, and ethnic group in multivariate analysis, the influence of race on the rate and severity of ROP is evidently an area that requires further clarification. 
The rat model of oxygen-induced retinopathy (OIR) mimics what occurs in the human form of ROP and continues to provide insight into the pathogenesis of this disease. Findings of ours have shown that in addition to the vasculopathy that develops in the rat model of OIR, permanent retinal cytoarchitectural and functional anomalies can also be documented. 13 14 15 16 17 The most striking changes in retinal ultrastructure that we have evidenced in the albino SD rat model of OIR include outer plexiform layer (OPL) thinning and a decrease in horizontal cell count. Attenuation in the amplitude of the electroretinogram (ERG) b-wave with relative sparing of the a-wave are also features that are characteristic of this model. 13 14 15 16 17 Previously published studies that compared pigmented and albino rats have shown that the former are less susceptible to ischemic retinal injury than are the latter, 18 whereas others have demonstrated that certain pigmented strains develop more severe retinal neovascularization. 19 20 Consequently, the purpose of this study was to determine whether similar strain differences might also extend to retinal vasculature (flatmounts), structure (histology), and function (ERG), in addition to neurotrophic factor expression. In the present study, we therefore sought to explore these factors in two rat models of OIR: the albino SD and the pigmented LE strains. 
Methods
Animals
All experiments were approved by the McGill University-Montreal Children’s Hospital Research Institute according to the guidelines of the Canadian Council on Animal Care and were conducted in accordance with the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research. Within a 24-hour period after birth, newborn litters of Sprague-Dawley (SD) and Long Evans (LE) rats (Charles River Laboratories, St. Constant, Québec, Canada) were placed, with their mothers, in room air (21% O2) or in a hyperoxic (80% O2) environment (mixture of medical grade 100% O2 and room air measured with an oxygen meter [MaxO2 Ceramatec, model OM25-ME; Medicana Inc., Montréal, Québec, Canada]). A controlled cyclic light environment was maintained throughout the exposure (12 hours light–dark). Exposure to hyperoxia lasted for 22.5 hours daily and was interrupted for three 0.5-hour periods of normoxia (21% O2), as per previously published protocols. 13 14 15 16 17 Casualties were more frequently observed throughout hyperoxic exposure of the pigmented LE strain, a finding that might be explained by interstrain differences in the rat genome that contribute to variations in ventilation, metabolism, and ventilatory responses to inspired levels of O2 and CO2. 21 22 Based on previous studies in SD rats that suggested unequal susceptibilities to hyperoxia throughout different developmental time points, 15 16 the hyperoxic cohort of each strain was divided at birth into seven different exposure protocols: from birth through postnatal day 6 ([P0–P6]: LE, n = 24 [P30], n = 5 [P60]; SD, n = 6 [P30], n = 6 [P60]); birth through postnatal day 9 ([P0–P9]: LE, n = 11 [P30], n = 5 [P60]); birth through postnatal day 12 ([P0–P12]: LE, n = 7 [P30], n = 6 [P60]); and birth through postnatal day 14 ([P0–P14]: LE, n = 21 [P30], n = 6 [P60]; SD, n = 11 [P30], n = 7 [P60]0; or from postnatal day 6, 9, or 12 through postnatal day 14 (P6–P14: LE, n = 22 [P30], n = 6 [P60]; SD, n = 8 [P30], n = 8 [P60]; P9–P14: LE, n = 17 [P30], n = 9 [P60]; and P12–P14: LE, n = 15 [P30], n = 7 [P60]). Select time points within the first and second weeks of life were taken for comparing SD rats to LE rats, as we had documented the effects of postnatal hyperoxia in the former strain elsewhere. 15 16 After hyperoxic exposure, animals were returned to room air (21% O2) until P30 when retinal function was first assessed. Data were compared with respective age-matched control litters raised in normoxic conditions (LE, n = 21; SD, n = 22). 
Malondialdehyde Assay
Malondialdehyde (MDA), a product of peroxidation, was determined as previously described. 23 24 LE and SD rats were killed immediately after exposure to hyperoxia (P0–P6 and P6–P9, n = 3 to 4 per group), given that levels (relative to normoxia) have been shown to subside as the hyperoxic period is extended for longer periods. 23 Each retina was harvested less than 1 minute later, ground twice for 30 seconds each with a homogenizer (Omni International, Marietta, GA) in a buffer containing butyl-hydroxytoluene (BHT; 5 mM Tris, 0.02% ASA, 0.5 mM EGTA), and centrifuged at 1000g for 10 minutes. To each sample was added 0.33% 2-thiobarbituric acid (TBA) in water mixed with glacial acetic acid and then the sample was heated at 95°C for 60 minutes. HPLC-grade n-butanol was added, and samples were vortexed and spun down at 1000g for 10 minutes. The upper phase was read at 532 nm with a spectrophotometer. 
Retinal Flatmounts
Groups of animals killed on P6 and P14 after exposure from P0 to P6 (SD, n = 3; LE, n = 3) and P6 to 14 (SD, n = 3; LE, n = 3), respectively. After enucleation and dissection of the cornea and lens, eye cups were collected and fixed overnight at 4°C in 4% vol/vol formalin. Retinas were isolated, and flatmounts were prepared for adenosine diphosphatase (ADPase) staining, according to a method that was previously described. 17 25 26  
Sequential, overlapping images of the entire retina were subsequently collected and arranged to construct a montage of the complete retina (Photoshop, ver. 6.0; Adobe Systems Inc., San Jose, CA). Vascularized areas were manually outlined and measured as a percentage of the total retinal area to assess vascular extent. Density analysis obtained from oxygen-exposed groups was compared with that from animals raised in normoxia, for which values were set at 100% (ImagePro Plus, ver. 4.1; Media Cybernetics, Silver Spring, MD). 
Retinal Immunohistochemistry
Other groups of animals were assigned to be killed on postnatal day 6 after exposure from P0 to P6 (LE, n = 3; SD, n = 3) and on P14 and P60 after exposure from P6 to P14 (P14: LE, n = 3; SD, n = 3; P60: LE, n = 3), respectively, for detection of endogenous levels of FGF-2 and CNTF neurotrophic factors that were found to be upregulated in another model of oxidative stress—namely, light-induced retinopathy (LIR). 27 Animals were euthanatized immediately after removal from hyperoxia and were perfused intracardially with 4% paraformaldehyde (PFA) in 0.2 M phosphate buffer (PB; pH 7.4). After removal of the cornea and lens as described, eye cups were immersed in 4% PFA in 0.2 M PB (pH 7.4; 2 hours) and were cryoprotected in sequential sucrose solution gradients (10%, 20%, and 30% in PB for 30 minutes, 1 hour, and overnight, respectively) at 4°C. Eye cups were subsequently embedded in optimal cutting temperature (OCT) compound (Tissue-Tek; Miles Laboratories, Elkhart, IN) and were frozen in a bath containing 2-methylbutane/liquid nitrogen. Fourteen-micrometer-thick cryosections were collected on polylysine-S-coated slides for immunohistochemical analysis. To block any nonspecific binding of the antibodies, we incubated retinal sections in 3% normal goat serum, 0.3% Triton X-100 (Sigma-Aldrich, St. Louis, MO) in 0.1 M PBS (pH 7.4) for 30 minutes at room temperature. The sections were then incubated overnight at 4°C with the following primary antibodies: monoclonal anti-human FGF-2 (type II, clone bFM-2, 10 μg/mL; Upstate Biotechnology, Lake Placid, NY), monoclonal anti-mouse CNTF (5 μg/mL; Chemicon, Temecula, CA), and polyclonal anti-rabbit GFAP (1:100; Sigma-Aldrich). Normally, GFAP expression in Müller cells is very limited 28 29 but increases after neuronal cell death. In fact, studies conducted in the rat model of OIR have suggested that increasingly intense GFAP reactivity is observed once animals are returned to room air after 14 days of hyperoxia, the most intense of which is seen at approximately 10 weeks of age. 30 Consequently, we chose to conduct our colocalization studies with GFAP in retinas from rats exposed to hyperoxia from P0 to 14, collected at P60. The next day, sections were incubated with the appropriate secondary antibody for 1 hour at room temperature, washed in PBS, and mounted with an antifade reagent (SlowFade; Invitrogen-Molecular Probes, Eugene, OR). Photographs were taken with a microscope (40× objective, Axiophot; Carl Zeiss Meditec, Oberkochen, Germany). 
Western Blot
Retinas from right and left eyes of LE and SD control and oxygen-exposed rats (80% from P0 to P6, P6 to P14, and P0 to P14) were isolated (n = 3 to 4 per group), independently homogenized with an electric pestle (Kontes, Vineland, NJ) in ice-cold lysis buffer (0.1% sodium dodecyl sulfate, 20 mM Tris [pH 8.0], 135 mM sodium chloride, 1% NP-40, 10% glycerol supplemented with protease inhibitors), and then pooled. Retinal lysates were kept on ice for 30 minutes and then centrifuged at 14,000 rpm for 15 minutes, after which the supernatant was collected. Protein concentration was determined by the Lowry method (Bio-Rad Life Science, Mississauga, ON, Canada). For each sample, 100 μg of protein was resolved by electrophoresis on 15% sodium dodecyl sulfate polyacrylamide gels (SDS-PAGE) and transferred to nitrocellulose membranes (Bio-Rad Life Science), as previously described. 27 After nonspecific signal blocking in 10 mM Tris (pH 8.0), 150 mM NaCl, 0.2% Tween 20 (TBST), and 5% lyophilized skim milk for 1 hour at room temperature, the membranes were incubated overnight at 4°C with the primary antibodies used according to the following conditions: monoclonal anti-human FGF-2 (type II, clone bFM-2, 2.5 μg/mL; Upstate Biotechnology) and monoclonal anti-mouse CNTF (5 μg/mL; Chemicon). The membranes were subsequently washed with TBST and incubated with anti-mouse peroxidase-linked secondary antibodies (Amersham Pharmacia, Baie d’Urfé, Québec, Canada) for 1 hour at room temperature. A chemiluminescent reagent (ECL; GE Healthcare, Piscataway, NJ) was used for detection of protein signals, followed by the exposure of the membranes to autoradiograph film (X-OMAT; Eastman Kodak, Rochester, NY). 
The membranes were incubated in a stripping solution (200 mM glycine [pH 2.8], 500 mM NaCl, and 0.7% β-mercaptoethanol) at 55°C for 1 hour and then reprobed with an anti-actin polyclonal antibody (1:10,000; generous gift from J. Chloe Bulinski) followed by incubation with an anti-rabbit peroxidase-linked secondary antibody (Amersham Pharmacia Biotech). 
The γ-actin protein was chosen as a reference instead of β-actin, given that the latter isoform patterns vary with age. 31 32 33 Ponceau-S (Sigma-Aldrich) was used to ensure equal protein loading. Densitometry analysis was performed on scanned autoradiographic films and quantified according to pixel intensity (Quantity One 4.1.0 software; Bio-Rad Laboratories). The densitometry values obtained for each neurotrophic factor were subsequently normalized to the γ-actin level in the same blot. 
Electroretinography
ERGs and oscillatory potentials (OPs) were recorded (AcqKnowledge Data Acquisition System, Biopac MP 100WS; BIOPAC Systems Inc., Goleta, CA) as previously described 13 14 15 16 17 from control and hyperoxic cohorts at postnatal days 30 and 60. Briefly, after 12 hours of dark adaptation, rats were anesthetized, under a dim red light illumination, with an intramuscular injection of ketamine (85 mg/kg) and xylazine (5 mg/kg). The pupils were subsequently dilated with 1 to 2 drops of 1% cyclopentolate hydrochloride, and the rat was placed on its side in a recording chamber that included both the flash stimulator and background light. A DTL fiber electrode (27/7 X-Static silver-coated conductive nylon yarn; Sauquoit Industries, Scranton, PA) that served as the active electrode was placed over the corneal surface and was kept in place with 2% methylcellulose (Gonioscopic solution; Alcon Laboratories, Fort Worth, TX), which also helped to prevent corneal desiccation. Grass disc (model E6GH; Grass Instruments, Quincy, MA) and subdermal needle electrodes (model E2; Grass Instruments) were placed in the mouth as the reference and inserted in the tail as the ground, respectively. Broadband ERGs (10,000×, 1–1000 zHz bandwidth, 6 dB of attenuation; P511 analog preamplifiers; Grass Instruments) and OPs (50,000×, 100–1000 Hz, 6 dB of attenuation; P511 analog preamplifiers) were recorded simultaneously with the data acquisition system (AcqKnowledge; Biopac MP 100 WS; Biopac System Inc.). Scotopic luminance–response functions were evoked by flashes of white light spanning a 6-log-unit range in 0.3-log increments (maximal intensity, 0.6 log cd · s · m−2, average, two to five flashes, interstimulus interval, 9.60 seconds). Photopic (cone-mediated) signals were evoked to flashes of white light of 0.9 log cd · s · m−2that were delivered after more than 15 minutes of light adaptation to a background of 30 cd · m−2 (average, 20 flashes; interstimulus interval, 1 second) to avoid the previously described light adaptation effect. 34  
Retinal Histology
Retinal histology was performed at P60. After the completion of ERG recordings, the rats were euthanatized with an anesthetic overdose followed by intracardiac perfusion with 3.5% glutaraldehyde in 0.1 M phosphate buffer (pH 7.4). The eyes were enucleated and postfixed in 3.5% glutaraldehyde for 3 hours. The corneas were then dissected, the lenses were removed as described earlier, and the eyes were subsequently placed in 3.5% glutaraldehyde overnight (12 hours). The eye cups were then immersed in a 4% osmium tetroxide solution with 0.1 M phosphate buffer. After sequential dehydrations in ethanol baths (50%, 70%, 90%, 95%, and 100% ethanol) and propylene oxide, the specimens were embedded in epoxy resin (Durcupan ACM Fluka epoxy resin kit; Sigma-Aldrich). Once polymerization was completed (48 hours at 58°C), the tissue was cut in 0.7-μm ultrathin sections that were mounted on glass slides and stained with 0.1% toluidine blue. Images were obtained with a digital camera (RR slider spot; Diagnostic Instruments Inc., Sterling Heights, MI) that was mounted on a microscope (Axiophot, 40×; Carl Zeiss Meditec). Three retinas per group were analyzed, and an average of 10 measurements was obtained from each. 
Data Analysis
Retinal histology and ERGs were obtained from the same group of animals, whereas retinal flatmounts, MDA assay, and neurotrophic factor analysis through Western blot analysis and immunofluorescence were all performed independently. Amplitudes of ERG components along with their peak times were measured as previously described. 13 14 15 16 17 Briefly, the a-wave amplitude was measured from baseline to the most negative trough, and the b-wave amplitude was measured from the trough of the a-wave to the peak of the b-wave. Similarly, oscillatory potentials were measured from the preceding trough to the peak of the OP under evaluation, with the exception of OP2, which was measured from the baseline to its peak. OPs were reported as the sum of OPs (SOP = OP2 + OP3 + OP4 + OP5). 
Scotopic luminance–response function curves were derived for each animal by plotting b-wave amplitudes against flash intensities. A sigmoidal intensity–response regression curve was then applied to fit the data (Prism 3.00 software; GraphPad, San Diego, CA), from which the rod V max (maximal rod response) could be calculated. Furthermore, the ERG response evoked to the brightest flash delivered in scotopic conditions (the mixed rod-cone response) was also analyzed. To determine the effect of the different oxygen regimens on retinal structure and function, we used one-way analysis of variance (ANOVA; P < 0.05) and the Dunnett multiple comparison as a post hoc test. Functional data obtained from each exposure regimen at each recording session (P30 and P60) in addition to retinal vascular findings and protein analysis (immunoblotting) were compared by Student’s t-test (P < 0.05). Finally, Student’s t-test (P < 0.05) was also used to compare values obtained at P30 and P60 from LE and SD rats from preselected oxygen exposure regimens. 
Results
Effect of Postnatal Hyperoxia on MDA Concentration
Given the effects of the redox state on peroxidation 35 36 we measured levels of malondialdehyde (MDA), a product of peroxidation. Although a 31.9% increase in MDA levels was observed in SD rats after early hyperoxia (P0–P6, 3.06 ± 0.34 [80%] compared with 2.32 ± 0.09 [21%]; P < 0.05), a strikingly greater difference in levels was found in LE rats after the same regimen, resulting in a 91.5% increase (P0–P6, 3.63 ± 0.13 [80%] compared with 1.90 ± 0.25 [21%]; P < 0.05). After hyperoxic exposures that extended into the second week of life (P6–P9), MDA levels were reduced to near normal levels (1.99 ± 0.35 [80%] compared with 2.50 ± 0.65 [21%] in SD rats and 2.47 ± 0.40 [80%] compared with 3.37 ± 0.69 [21%] in LE rats; P > 0.05). 
Effect of Postnatal Hyperoxia on Retinal Vasculature
Retinal vasculature in the neonatal rat undergoes significant development from birth through P14, beginning in the central retina and moving outward to the periphery. 13 Retinal flatmounts reveal that normoxia-raised LE and SD rats are similarly vascularized at P6 (78.50% ± 8.11% and 71.67% ± 2.89%, respectively; Figs. 1A 1E ), whereas vascular growth to the periphery is complete (100% vascularized) by P14 (Figs. 1C 1G) . On the other hand, hyperoxia (P0–P6 exposure) significantly attenuated the postnatal vascular growth, reducing the vascularized area to 28.7% (P < 0.05) and 57.2% (P < 0.05) of control in LE and SD rats, respectively (Figs. 1B 1F 2A) . This attenuation in vascular growth was accompanied by a marked reduction in vascular density for the same exposure regimen, resulting in 51.03% ± 4.25% (P < 0.05) and 66.70% ± 3.96% (P > 0.05) of normal density in LE and SD rats, respectively (Fig. 2C)
Similarly, exposure from P6 to 14 resulted in a vascularized area that was also reduced in LE rats to 57.90% ± 28.15% of normal coverage (P < 0.05), whereas SD rats were not significantly affected (94.76% ± 5.61% of normal, P > 0.05; Figs. 1D 1H 2B ). As shown in Figure 2D , this trend was also reflected in vascular density measurements (57.03% ± 18.72% and 113.36% ± 16.46% of normal in LE [P < 0.05]) and SD [P > 0.05] rats, respectively). 
Effect of Postnatal Hyperoxia on Retinal Function
Figure 3shows representative scotopic (Figs. 3A 3B 3C 3D 3E 3F)and photopic (Figs. 3G 3H)ERGs (Figs. 3A 3B 3C 3D 3E 3G)and oscillatory potentials (Figs. 3F 3H)recorded from control (21% O2) and hyperoxic rats (80% O2) at P60. Exposure durations are indicated in postnatal day intervals. Group data are graphically reported in Figure 4 . Although parallel experiments for LE and SD rats were performed, retinal function characteristics of SD rats have been reported by us 13 14 15 16 17 ; hence, a more elaborate description of retinal function parameters will be focused on LE rats, and comparisons concluded for strain differences. 
In normal LE rats, the use of a progressively brighter flash stimulus (from bottom to top) resulted in a gradual increase in the a- and b-wave parameters, a trend also observed in rats that were exposed postnatally to each of the hyperoxic regimens. 
A comparison between the scotopic (a-wave, rod V max, rod-cone b-wave, and SOPs) and photopic (b-wave and SOPs) ERG parameters of LE and SD rats obtained at P30 and P60 is graphically represented in Figure 4 . Group data along with statistical analysis for LE and SD rats are shown in Tables 1 and 2 , respectively. 
Effect of Postnatal Hyperoxia on Scotopic Function
In all instances, there is a gradual decline in the amplitude of ERG parameters at P30 and P60, culminating in maximum attenuation after exposures to hyperoxia from P0 to P14 and P6 to P14. After hyperoxia initiated at birth, a dose–response effect was observed between the duration of hyperoxia and the amplitude of the resulting a-wave amplitude in LE rats (Fig. 4A)for measurements obtained at P30 and P60. Although a tendency for a decline in a-wave amplitude was observed on exposure to hyperoxia in the first postnatal week (Fig. 4A) , a more pronounced attenuation was observed when hyperoxic exposure extended into the second week of life; this culminated in a decline to 51.9% and 45.6% of control values at P30 and P60, respectively (P < 0.05). Regimens that were limited to the second week of life also resulted in an attenuation of the a-wave amplitude to 63.4% and 42.9% of P30 and P60 control values, respectively (P < 0.05). Notably, the a-wave is considerably more affected in LE than SD rats exposed to hyperoxia, where a decline to 79.7% and 65.7% of control values, respectively, was observed at P30 and P60 in SD rats, compared with 51.9% and 45.6% of control values at P30 and P60, in LE rats (Fig. 4G)
A similar dose–response correlation between the duration of hyperoxia and the amplitude of the rod V max in LE rats was also observed in P30 and P60 recording sessions (Fig. 4B) . The amplitude of the rod V max was already attenuated by hyperoxia from P0 to P6 to 84.9% (P30) and 74.4% (P60) of control; this parameter continued to decline when oxygen exposure was extended into the second week of life. Similar findings were also observed when hyperoxia was initiated in the second week resulting in attenuations to 18.2% (P30) and 13.0% (P60) of control (P < 0.05). As seen for the a-wave, the rod V max attenuation after maximum hyperoxia was notably less severe in SD rats (Fig. 4H)than in LE rats. 
A comparable trend was also observed for the rod-cone b-wave of LE rats (Fig. 4C) , where significant attenuations in amplitude were observed after hyperoxic exposure from P0 to P6 to 68.7% of control at P30 (P < 0.05) and 59.9% of control at P60 (P < 0.05). Exposures from P0 to P14 further reduced the amplitude of the rod-cone b-wave. When initiated later in life, hyperoxia caused similar amplitude attenuations that were most pronounced after exposure from P6 to P14 (reductions to 21.7% of control at P30 and 17.6% of control at P60; P < 0.05). 
Finally, given that there was no specific effect of hyperoxia on any particular oscillatory potential (OP), we are reporting them collectively as the sum of OPs (SOPs). Scotopic SOPs (Fig. 4E)in LE rats were all significantly attenuated after hyperoxia, irrespective of the exposure regimen. Attenuations in SOP amplitude were observed in exposures from P0 to P6 (59% of control [P30], and 68.7% of control [P60]; P < 0.05). After maximum exposure from P0 to P14, SOP amplitudes were significantly (P < 0.05) reduced to 6.2% and 12.7% of control amplitude at P30 and P60, respectively. Hyperoxia resulted in an almost equivalent attenuation after P6 to P14 as it did after a 2-week oxygen exposure. 
Again, the rod-cone b-wave (Fig. 4I)along with the scotopic SOPs (Fig. 4K)were less attenuated after postnatal hyperoxia in SD rats where maximum rod-cone b-wave attenuations were 36.9% (P30) and 40.3% (P60) of control compared with 18.3% (P30) and 20.5% (P60) of control for the scotopic SOPs (P < 0.05). 
Effect of Postnatal Hyperoxia on Photopic Function
The photopic b-wave and SOPs of LE rats raised in a hyperoxic environment were also markedly affected. A striking decline in the photopic b-wave amplitude (Fig. 4D) , was observed after P0 to P6 exposure to 53.0% and 51.6% of control at P30 and P60, respectively (P < 0.05); this trend continued during extended hyperoxia (P0–P14). Oxygen exposure regimens initiated in the second week of life also resulted in significant amplitude attenuations. Photopic SOPs were similarly affected after postnatal hyperoxia (Fig. 4F) . As described for the scotopic condition, the cone-mediated function of the SD rats also appeared to be less susceptible to the hyperoxic insult than was that of the LE rats. Along these lines, after hyperoxia from P0 to P14, the photopic b-wave of the SD rats (Fig. 4J)was attenuated to 27.3% and 32.7% of control at P30 and P60, respectively, compared with 11.9% and 20.3% in the LE rats. 
Effect of Postnatal Hyperoxia on Retinal Cytoarchitecture
The most striking ultrastructural consequences of postnatal hyperoxia, reported in the neonatal SD rat model included the near disappearance of the OPL after maximum exposure to approximately 20.0% of control along with a significant reduction in horizontal cell count to approximately 20.0% of the control, both of which were most obvious after regimens that extended hyperoxia into the second week of life 13 14 16 17 (Figs. 5F 5G) . Added to this, postnatal hyperoxia in LE rats also damaged other retinal structures as exemplified with representative histologic sections shown in Figures 5A 5B 5C 5D 5E . These are characterized by a marked thinning of the inner retina as well as outer retinal damage limited to the outer nuclear layer (ONL). Group data are graphically presented in Figure 6 . As shown in Figures 5B and 6 , early exposure from P0 to P6 resulted not only in a reduction in OPL thickness to 65.6% of control (P < 0.05), but also in a significantly thinner ONL, inner nuclear layer (INL), and inner plexiform layer (IPL; reduced to 61.5%, 80.2%, and 70.9% of the respective controls). Progressive increases in duration of oxygen exposure, particularly when encompassing the second week of postnatal life, resulted in even greater structural anomalies. As shown in Figures 5C and 6 , exposures within the first 2 weeks of life resulted in maximum thinning of the ONL, OPL, INL, and IPL to 67.6%, 0.7%, 26.0%, and 45.8% of control, respectively (P < 0.05). When hyperoxia was initiated in the second postnatal week (P6–P14 and P12–P14; Figs. 5D 5E 6 ), the thinning of the retinal layers was more important than that observed after exposures in the first postnatal week, but not as severe as that observed during 2 weeks of hyperoxia (P0–P14). 
Neurotrophic Factor Expression after Postnatal Hyperoxia
Previous studies have suggested that mechanical injury to the retina, 37 38 39 in addition to light-induced damage, 40 41 42 can lead to the upregulation of neurotrophic factors. Furthermore, studies conducted in our laboratory have suggested that an increase in endogenous levels of some of these factors, such as FGF-2 and CNTF for example, plays a role in the neuroprotection of juvenile rat retinas against another environmental oxidative stress—namely, light-induced damage. 27 To determine whether a similar protective mechanism might explain the difference between SD and LE rats, we examined the levels of retinal FGF-2 and CNTF (Fig. 7)by Western blot analysis. LE and SD rat retinas were collected immediately after early oxygen exposure (P0–P6) at P6 and late oxygen exposure (P6–P14) at P14 and were compared with retinas collected from age-matched control rats at P6 and P14, respectively. Although FGF-2 levels in LE rats had a tendency to increase after exposures from P0 to P6, P6 to P14, and P0 to 14 compared with their respective controls (Fig. 7A) , results did not reach statistical significance. Similarly, FGF-2 levels also tended to increase in SD rats after postnatal hyperoxia (Fig. 7B)after each exposure regimen compared with control. It is interesting to note, however, that control SD rats appeared to have approximately four to five times higher endogenous levels of FGF-2 than did the LE rats. This tendency extended during exposure to hyperoxia. 
In all hyperoxic cohorts of LE and SD rats studied, CNTF levels appeared higher compared with their corresponding controls (Figs. 7C 7D)
Retinal immunohistochemistry was performed to determine the cellular localization of FGF-2 and CNTF proteins in LE and SD rats after exposure to hyperoxia. Figures 8A and 8Breveal that, as described above, FGF-2 immunoreactivity was more pronounced during exposure to hyperoxia compared with the control. Immunofluorescence in both strains was limited to the INL and ganglion cell layer (GCL). Basal levels of CNTF staining were observed in the INL and GCL of the control animals and were markedly increased after exposure to hyperoxia in both LE and SD rats (Figs. 8C 8D) . The localization of CNTF suggested that these were Müller cells (GFAP positive, Fig. 9 ). 
Discussion
Previous studies showed that postnatal hyperoxia in the albino SD rat model of OIR resulted in severe structural and functional damage, including the permanent loss of the OPL, significant reduction in the horizontal cell count, and permanent attenuation in retinal function, as observed with the scotopic and photopic ERGs. 13 14 15 16 17 Results from the present study suggest an even greater susceptibility of the pigmented LE rat to postnatal hyperoxia, as evidenced with higher vascular dropout throughout exposure, more severely depressed retinal function and permanently altered retinal structure that was not only limited to the loss of the OPL and horizontal cells as in SD rats, but also extended into the inner retina, resulting in significant loss of the INL and IPL. 
These alterations in retinal cytoarchitecture are likely to be at the root of the functional anomalies, such as the severe attenuation of the ERG b-wave we are reporting. The most critical, and most permanent, sequelae of OIR in the SD rat model occurred in the more inner part of the retina. However, the tendency for the shortening of inner segments of the photoreceptors which is accompanied by a significantly attenuated a-wave in LE rats after hyperoxia is suggestive of a photoreceptor involvement as well. Therefore, the short- and long-term consequences of postnatal hyperoxia on the retinal structure and function were significantly more pronounced in LE rats than in SD rats. 
Free radicals that are generated through lipid peroxidation have been thought to play an important role in the pathogenesis of ROP and its animal counterpart, OIR. 43 It has been suggested that the formation of free radicals in the retina may result from the exposure of premature infants to a hyperoxic environment, particularly due to their deficiency in antioxidant defenses, such as superoxide dismutase, α-tocopherol, catalase, and glutathione peroxidase, for example. 44 45 Reactive oxygen species (ROS) have also been shown to contribute to the vaso-obliterative phase that is associated with OIR 46 47 48 49 either through direct cytotoxic effects on endothelial cells or through an initial phase of vasoconstriction which ultimately results in reduced tissue perfusion. Of interest, treatment with vitamin E, a free-radical scavenger, has been shown to protect against the loss of superoxide dismutase in cats exposed to hyperoxia 50 and has successfully attenuated the severity of vaso-obliteration in the rat model of OIR. 49 51 A previous study of ours revealed that administration of the water-soluble antioxidant Trolox C to SD rats prevents only some of the structural and functional sequelae associated with OIR, suggesting that free radical formation may not be the sole cause of the damage. 14 Similarly, our current results suggest that melanin would be insufficient in having any beneficial protective effect under similar circumstances; and moreover, it appeared as though it might even potentiate the effect of ROS. Melanin is produced and stored in the retinal pigment epithelium (RPE) cytoplasm and melanocytes, and continuous exposure of the RPE to environmental stressors such as light and oxygen have been thought to dampen its antioxidant capacity. 52 Furthermore, melanin has been known to take on the role of a pro-oxidant under these conditions, leading to cytotoxic activity such as photoreceptor damage and even age-related macular degeneration. 53 54 This would explain the severity of OIR in LE rats, including a significantly injured photoreceptor layer, given its proximity with the RPE. Our findings of increased MDA levels in LE compared with SD rats (after hyperoxic exposure initiated at birth throughout the first week of life) support this inference. Recently, we showed that despite the damage to retinal vasculature that is observed within the first week or so of exposure of SD rats (e.g., P0–P6 and P0–P9), the retinal vascular growth process appears to be able to fight this oxidative stress to achieve nearly full coverage while remaining under hyperoxic exposure throughout the first 2 weeks of life, 13 a finding that is further corroborated by the results in the present study. A different picture emerges in the pigmented rat, however, where vascular dropout and extent are so severely compromised that they never achieve a normal appearance (Fig 1) . Taken together, these anatomic characteristics are inconsistent with a protective role of melanin in the retinal vasculature and inner structures. This result suggests that other mechanisms partake in the increased vulnerability of LE compared with SD rats, as observed in the present study. It is of interest to note that the complex series of events that characterize retinal development occur differently in albino and pigmented animals, where retinal maturation is delayed in the absence of melanin. 55 For example, studies have revealed a temporal delay in the central-to-peripheral pattern of cell production 56 57 along with a delay in cell death of the INL and GCL in the albino rat retina. 57 One wonders whether the more “mature” retina of the LE rat might result in a greater difficulty in overcoming hyperoxic stress, whereas the more plastic immature retina of the albino rat is better able to overcome a similar stress. Variations in the responses of different rat strains to hyperoxia, including the increased susceptibility of the Dark Agouti and Hooded Wistar rats compared with several albino strains, 58 as well as that of the pigmented offspring of Fischer 344 X Dark Agouti compared with the albino progeny, 59 further support the association between ocular pigmentation and vulnerability in OIR. Furthermore, although no difference in the time course of retinal vascular development was observed between age-matched control Brown Norway and SD rats in a study by Gao et al., 19 those raised in hyperoxia showed significant differences in vascular permeability and retinal neovascularization. For example, retinal artery constriction along with large avascular regions in the retina were observed in Brown Norway rats at P12—that is, 1 day after their removal from the hyperoxic environment. In contrast, SD rats studied under the same conditions revealed a lower degree of retinal artery constriction and larger areas of vascular perfusion throughout the retina. 19 Increased vascular permeability followed by a more severe neovascularization in the Brown Norway strain 19 60 61 was also observed. This effect was characterized by tortuosity and dilatation of retinal vessels, neovascular tufts, and hemorrhage when compared with the SD rats, which were lacking neovascular tufts and evidence of microaneurysms. 19 Moreover, the Brown Norway model of OIR revealed higher retinal VEGF levels and more severe retinal vascular leakage than did the SD strain. 62 Taken together with our current results and considering that the secondary neovascular phase of OIR is thought to be a mirror image of the initial vasoconstriction and vaso-obliteration which takes place, these findings suggest that retinal vascular development along with the neovascularization phase that ensues is significantly more affected by hyperoxia in pigmented than albino SD rats. Variations in the regulation of the antiangiogenic pigment epithelium–derived factor (PEDF) and the angiogenic vascular endothelial growth factor (VEGF) among strains are thought to contribute to these discrepancies, which may be genetically regulated, 19 not to mention other strain-dependant factors that may be involved in regulating the effects of oxygen tension on retinal angiogenesis, such as glial cell sensitivity to hyperoxia, for example. 63  
Evidently, despite using identical oxygen exposure protocols, the pathophysiological consequences of OIR were much more severe in the pigmented LE model than in the albino SD rat model. Not only did the hypothesized free-radical–scavenging role of melanin fail to protect the LE cohorts that were exposed to hyperoxia postnatally, but significantly greater levels of the MDA product of peroxidation were also evidenced in the LE model. We therefore sought to further understand the mechanisms by which SD rats remained significantly more resistant to a hyperoxic stress by exploring the potential involvement of specific neurotrophic factors throughout the hyperoxia. In a previous study, we showed that the relative (compared with adult) resistance of juvenile SD rats exposed after birth to a bright luminous environment (not combined with oxygen), another oxidative stress, might be explained by the upregulation of neurotrophic factors such as FGF-2 and CNTF. In adult retinas, FGF-2 and CNTF levels are also upregulated after mechanical injury. Furthermore, intravitreous injection of the above-mentioned factors in addition to brain-derived neurotrophic factor (BDNF) and glia-derived neurotrophic factor have all been shown to delay photoreceptor cell death in animal models of retinal degeneration. 64 65 66 67 68 69 We therefore hypothesized that the expression of similar survival factors might explain the relative resistance of SD rats to (1) photoreceptor damage (as evidenced by the relatively intact a-wave) and to (2) more severe thinning of the inner retinal layers. Of note, our results suggest that although only limited variations in FGF-2 occur in both LE and SD models after hyperoxic exposure, higher levels (approximately twofold) are observed for each exposure regimen in SD rats. Furthermore, although CNTF levels are only significantly higher after exposure regimens that include the second week of life in LE rats, they are already significantly greater in SD rats after exposure within the first week (e.g., from P0–P6). These results would therefore suggest that SD rats are better equipped with survival factors in the face of a hyperoxic challenge. The involvement of CNTF and FGF-2 may explain the preservation of photoreceptor function in SD rats as well as the relative resistance to functional and structural damage that follows hyperoxia from P0 to P6. Moreover, previous studies have shown that, in the adult retina, light-induced damage provokes Müller cell gliosis, 70 which in turn releases several neurotrophic factors. 38 39 40 71 72 73 Microglia have been thought to play an integral role in this process as they have been shown to secrete factors that diffuse to the outer retina. 74 Given the relatively intact inner retina that is observed in SD rats after hyperoxic exposure, the above may indeed be a plausible mechanism of photoreceptor survival in this model. The severely compromised inner retina of LE rats exposed to the same conditions, however, would make it quite difficult for a similar protective mechanism to take place and may consequently explain their increased susceptibility, particularly where photoreceptor function is concerned. 
Conflicting results have been obtained using gene transfer strategy in Prph2Rd2/Rd2 mice receiving subretinal injections with adenoviral (Ad)-vectored CNTF that resulted in the preservation of ERG amplitudes 75 76 and with adenoassociated virus (AAV)-vectored CNTF that resulted in accelerated deterioration of ERG amplitudes, 77 suggesting that therapeutics of this nature still require further study. More recently, however, Dong et al. 78 demonstrated that double transgenic mice with inducible glial-derived neurotrophic factor (GDNF) provides protection from loss of retinal function (a- and b-waves of the ERG) generated by three different sources of oxidative stress, including one where mice were exposed to 75% oxygen for 3 weeks. It may therefore also be worthwhile to explore the potential antioxidant activity (or other) that GDNF exerts on pigmented and albino models of OIR and in other models of disease in which oxidative damage plays a role. 78  
As described earlier, there is an apparent discrepancy between some clinical studies of human ROP which have shown, for example, that a greater number of Caucasian infants reach threshold ROP compared with African-American infants 8 10 and the animal model, which suggests a positive correlation between pigmentation and the severity of OIR. 
Of interest, however, the greater susceptibility of the SD strain to vascular attenuation after hyperoxia compared with other albino strains such as the Fischer 344, Wistar-Furth, and Lewis strains would further support a role for factors other than ocular pigmentation. 55 Given that genetic factors appear to influence the angiogenic response of different rat strains, it would not be surprising that they also play a role in the increased incidence and severity of the retinopathy observed on a functional and structural level. Of interest, studies have shown that ventilatory responses are thought to be genetically as well as environmentally regulated 21 22 resulting in varying responses to changes in O2 and CO2 levels, for example. 22 In fact, our findings of an increased mortality rate of Brown Norway rats after exposure to the same hyperoxic protocol used to obtain our current results may be explained by these strain-related differences, which suggests an unequal susceptibility of different rat strains. It is also of interest to mention that differences in the incidence and severity of OIR were observed among Sprague-Dawley rats obtained from independent commercial sources, suggesting an important role of genetics, even within a given strain (Kitzmann AS, et al. IOVS 2002;43:ARVO E-Abstract 1256). Of course, studies aimed at further elucidating the genetic and racial factors affecting the incidence of ROP in humans are even further complicated by several confounding variables including sex, socioeconomic status, geographical areas, and environment, although a deeper understanding of these factors may explain the relative sparing of some ethnicities with respect to the development of severe ROP versus others that seem more susceptible. Although a connection between racial variation and the incidence and severity of diseases such as ROP 8 9 10 and age-related macular degeneration (ARMD) 79 80 have been documented, it is important to take into consideration that very little is understood regarding the functional mechanisms of differences in pigmentation 81 and that current studies are under way that are aimed at better understanding the molecular composition of melanosomes, along with their structure and function in the eye. 82 83 Other studies have shown that melanin may not act alone as the substance responsible for the protective effect, 84 85 but may instead be a confounding variable that is associated with certain genes that are more or less prevalent in certain populations. Genetic influences in the pathogenesis of these diseases may play an important role in the heterogeneity in risk of their progression. 
The underlying genetic differences that could explain the differential susceptibility between rat models of OIR as well as humans of different ethnicities remain to be explored. 
In conclusion, whereas some clinical studies suggest that the presence of increased melanin content confers added protection to infants that are postnatally exposed to hyperoxia as observed in African-Americans that appear to be protected against the development of severe ROP, 8 10 other studies have concluded that fundus pigmentation may play only a limited role in this protection. 11 12 Present animal findings suggest an increased susceptibility of pigmented LE rats to postnatal hyperoxia in comparison with the albino SD rat. Given the discrepancy between findings, future studies warrant careful investigation of other factors, including the contribution of genetics, which may help to elucidate the differences between LE and SD rats (as well as others) and to further our understanding of the pathogenesis of OIR and ROP, to potentially facilitate the derivation of therapeutic modalities. 
 
Figure 1.
 
Representative retinal vascular flatmounts stained with ADPase from LE (top) and SD (bottom) rats, raised in room air and euthanatized at P6 (A, E, respectively) or P14 (C, G, respectively), or raised in hyperoxia and euthanatized immediately after 6 days (P0–P6; B, F, respectively) or P14 (P0–P14; D, H, respectively) of oxygen exposure. Magnification, ×40.
Figure 1.
 
Representative retinal vascular flatmounts stained with ADPase from LE (top) and SD (bottom) rats, raised in room air and euthanatized at P6 (A, E, respectively) or P14 (C, G, respectively), or raised in hyperoxia and euthanatized immediately after 6 days (P0–P6; B, F, respectively) or P14 (P0–P14; D, H, respectively) of oxygen exposure. Magnification, ×40.
Figure 2.
 
Graphic representation of retinal vasculature extent (A, B) and vascular density (C, D) (ordinate) as a function of the oxygen exposure regimen (days, abscissa) in the normoxic and hyperoxic LE and SD cohorts at P6 (SD, n = 3; LE, n = 3) or after 6 days of hyperoxia (P0–P6; A, C, respectively; SD, n = 3; LE, n = 3) and at P14 or after 14 days of hyperoxia (P0–P14; B, D, respectively; SD, n = 3; LE, n = 3). Results are expressed as a percentage of vascular extent or vascular density of control retinas at P6 and P14, respectively, ±1 SD. *Statistically significant differences from the control at a given age found with the Student’s t-test (P < 0.05); •, significant differences from SD rats for the same exposure regimen (P < 0.05).
Figure 2.
 
Graphic representation of retinal vasculature extent (A, B) and vascular density (C, D) (ordinate) as a function of the oxygen exposure regimen (days, abscissa) in the normoxic and hyperoxic LE and SD cohorts at P6 (SD, n = 3; LE, n = 3) or after 6 days of hyperoxia (P0–P6; A, C, respectively; SD, n = 3; LE, n = 3) and at P14 or after 14 days of hyperoxia (P0–P14; B, D, respectively; SD, n = 3; LE, n = 3). Results are expressed as a percentage of vascular extent or vascular density of control retinas at P6 and P14, respectively, ±1 SD. *Statistically significant differences from the control at a given age found with the Student’s t-test (P < 0.05); •, significant differences from SD rats for the same exposure regimen (P < 0.05).
Figure 3.
 
Representative scotopic (mixed rod-cone, flash intensity: −6.3 through 0.60 log cd · s · m−2; AE) and photopic (flash intensity: 0.9 log cd · s · m−2, background: 30 cd · m−2; (G) ERGs along with scotopic (F) and photopic (H) oscillatory potentials obtained at P60 from animals raised in normoxia (21% O2) and hyperoxia (80% O2) from P0 to P6, P0 to P14, P6 to P14, and P12 to P14. Calibrations: horizontal, 20 ms; vertical, 400 μV for scotopic mixed rod-cone ERG responses, 200 μV for photopic ERG responses and 40 μV for scotopic and photopic oscillatory potentials, respectively. All tracings include a 20-ms prestimulus baseline; arrows, flash onset.
Figure 3.
 
Representative scotopic (mixed rod-cone, flash intensity: −6.3 through 0.60 log cd · s · m−2; AE) and photopic (flash intensity: 0.9 log cd · s · m−2, background: 30 cd · m−2; (G) ERGs along with scotopic (F) and photopic (H) oscillatory potentials obtained at P60 from animals raised in normoxia (21% O2) and hyperoxia (80% O2) from P0 to P6, P0 to P14, P6 to P14, and P12 to P14. Calibrations: horizontal, 20 ms; vertical, 400 μV for scotopic mixed rod-cone ERG responses, 200 μV for photopic ERG responses and 40 μV for scotopic and photopic oscillatory potentials, respectively. All tracings include a 20-ms prestimulus baseline; arrows, flash onset.
Figure 4.
 
Graphic representation of a-wave (A, G), rod V max (B, H), rod-cone b-wave (C, I), photopic b-wave (D, J), scotopic SOP (E, K), and photopic SOP (F, L) amplitudes (ordinate in microvolts) obtained at P30 and P60, respectively (abscissa in days of exposure), from LE and SD rats. The hyperoxic cohort of each strain was divided at birth into seven different exposure protocols: from birth through postnatal day 6 (P0–P6: LE, n = 24 [P30], n = 5 [P60]; SD, n = 6 [P30], n = 6 [P60]); birth to postnatal day 9 (P0–P9: LE, n = 11 [P30], n = 5 [P60]); birth to postnatal day 12 (P0–P12: LE, n = 7 [P30], n = 6 [P60]); and birth to postnatal day 14 (P0–14, LE, n = 21 [P30], n = 6 [P60]; SD, n = 11 [P30], n = 7 [P60]) or from P6, P9, or P12 through P14 (P6–P14: LE, n = 22 [P30], n = 6 [P60]; SD, n = 8 [P30], n = 8 [P60]; P9–P14: LE, n = 17 [P30], n = 9 [P60]; P12–14: LE, n = 15 [P30], n = 7 [P60]). Data were compared with results for respective age-matched control litters raised in normoxia (LE, n = 21; SD, n = 22).
Figure 4.
 
Graphic representation of a-wave (A, G), rod V max (B, H), rod-cone b-wave (C, I), photopic b-wave (D, J), scotopic SOP (E, K), and photopic SOP (F, L) amplitudes (ordinate in microvolts) obtained at P30 and P60, respectively (abscissa in days of exposure), from LE and SD rats. The hyperoxic cohort of each strain was divided at birth into seven different exposure protocols: from birth through postnatal day 6 (P0–P6: LE, n = 24 [P30], n = 5 [P60]; SD, n = 6 [P30], n = 6 [P60]); birth to postnatal day 9 (P0–P9: LE, n = 11 [P30], n = 5 [P60]); birth to postnatal day 12 (P0–P12: LE, n = 7 [P30], n = 6 [P60]); and birth to postnatal day 14 (P0–14, LE, n = 21 [P30], n = 6 [P60]; SD, n = 11 [P30], n = 7 [P60]) or from P6, P9, or P12 through P14 (P6–P14: LE, n = 22 [P30], n = 6 [P60]; SD, n = 8 [P30], n = 8 [P60]; P9–P14: LE, n = 17 [P30], n = 9 [P60]; P12–14: LE, n = 15 [P30], n = 7 [P60]). Data were compared with results for respective age-matched control litters raised in normoxia (LE, n = 21; SD, n = 22).
Table 1.
 
Group Data Obtained in Scotopic and Photopic Conditions from LE Rats Raised in Normoxia and Hyperoxia
Table 1.
 
Group Data Obtained in Scotopic and Photopic Conditions from LE Rats Raised in Normoxia and Hyperoxia
Age Control P0–P6 P0–P9 P0–P12 P0–P14 P6–P14 P9–P14 P12–P14
Scotopic parameters
 a-Wave (μV) 30 d 264.34 ± 87.25 256.41 ± 85.60 223.34 ± 77.82 166.76 ± 67.68* 137.18 ± 44.29* 167.65 ± 64.47* 223.42 ± 89.78 282.08 ± 130.44
60 d 220.29 ± 58.84 185.80 ± 59.51 182.15 ± 56.72 175.00 ± 100.36 100.49 ± 27.73 94.57 ± 61.72* , † 217.62 ± 121.44 202.48 ± 153.34
 Rod V max (μV) 30 d 497.58 ± 124.50 422.54 ± 123.10 202.82 ± 96.03* 79.90 ± 74.54* 81.63 ± 59.71* 90.32 ± 58.76* 173.61 ± 105.87* 341.60 ± 96.67*
60 d 544.61 ± 178.97 405.44 ± 109.12 209.85 ± 138.10* 186.29 ± 127.92* 148.79 ± 98.49* 70.99 ± 42.08* 135.44 ± 119.43* 308.85 ± 101.74*
 Rod-cone b-wave (μV) 30 d 864.19 ± 193.40 593.61 ± 200.20* 277.20 ± 137.83* 133.00 ± 91.77* 139.84 ± 58.11* 187.31 ± 68.79* 362.09 ± 269.4* 680.97 ± 304.62*
60 d 806.76 ± 257.07 483.50 ± 217.97 281.00 ± 204.02* 184.08 ± 153.75* 200.32 ± 122.22* 142.08 ± 46.91* 468.23 ± 350.81* 598.27 ± 423.08*
 SOPs (μV) 30 d 388.82 ± 100.64 229.23 ± 91.31* 99.52 ± 48.02* 51.05 ± 40.36* 24.28 ± 20.95* 51.28 ± 45.03* 93.99 ± 88.76* 277.80 ± 177.02*
60 d 395.69 ± 94.98 271.81 ± 142.43 155.79 ± 121.60* 81.75 ± 78.20* 50.14 ± 47.45* 44.91 ± 37.73* 139.43 ± 108.68* 233.38 ± 185.07*
Photopic parameters
 b-Wave (μV) 30 d 209.61 ± 72.44 111.19 ± 40.21* 49.06 ± 35.97* 31.41 ± 13.51* 25.02 ± 10.79* 42.87 ± 27.22* 67.41 ± 53.26* 126.59 ± 71.18*
60 d 190.29 ± 52.24 98.22 ± 40.65* 63.65 ± 58.33* 42.42 ± 23.21* 38.68 ± 21.13* , † 35.50 ± 23.65* 89.44 ± 64.71* 147.44 ± 102.81
 SOPs (μV) 30 d 55.88 ± 21.24 42.89 ± 16.92 19.42 ± 17.35* 7.71 ± 7.99* 4.20 ± 6.28* 11.11 ± 14.34* 28.97 ± 32.93* 43.67 ± 25.46
60 d 62.93 ± 17.79 36.51 ± 20.08* 34.08 ± 20.46* 22.8 ± 11.96* , † 13.37 ± 10.57* , † 9.87 ± 6.67* 25.67 ± 15.67* 38.91 ± 30.77
Table 2.
 
Group Data Obtained in Scotopic and Photopic Conditions from SD Rats Raised in Normoxia and Hyperoxia
Table 2.
 
Group Data Obtained in Scotopic and Photopic Conditions from SD Rats Raised in Normoxia and Hyperoxia
Age Control P0–P6 P0–P14 P6–P14
Scotopic parameters
 a-Wave (μV) 30 d 361.07 ± 109.65* 315.30 ± 50.70 287.65 ± 69.38* 305.27 ± 70.41*
60 d 326.35 ± 57.68* 337.68 ± 85.41* 214.32 ± 82.09* 357.36 ± 96.43*
 Rod V max (μV) 30 d 496.25 ± 146.35 366.03 ± 66.63, † 123.15 ± 92.98, † 201.19 ± 138.68* , †
60 d 529.24 ± 85.92 392.08 ± 90.25 138.08 ± 137.25, † 232.96 ± 114.33* , †
 Rod-cone b-wave (μV) 30 d 879.35 ± 226.06 656.83 ± 122.92, † 324.79 ± 116.96* , † 430.05 ± 164.67* , †
60 d 766.74 ± 107.27, ‡ 661.67 ± 160.81 309.18 ± 248.22, † 472.25 ± 102.94* , †
 SOPs (μV) 30 d 574.01 ± 297.05* 353.63 ± 81.25* 104.86 ± 69.00* , † 225.95 ± 133.51* , †
60 d 470.19 ± 237.24 329.01 ± 92.99 96.44 ± 89.80, † 244.39 ± 54.7* , †
Photopic parameters
 b-Wave (μV) 30 d 209.25 ± 72.92 128.05 ± 33.09, † 57.18 ± 34.09* , † 87.99 ± 42.61* , †
60 d 160.49 ± 30.08, ‡ 143.67 ± 38.34 52.43 ± 44.79, † 91.72 ± 19.58* , †
 SOPs (μV) 30 d 73.05 ± 18.91* 51.91 ± 7.57 19.15 ± 25.35* , † 34.81 ± 25.96, †
60 d 59.47 ± 40.31 55.94 ± 18.96 9.64 ± 7.66, † 41.28 ± 23.73*
Figure 5.
 
Representative retinal histologic sections (central retina) from LE rats obtained at P60 from the normoxic cohort (A), and from hyperoxic cohorts including P0 to P6 (B), P0 to P14 (C), P6 to P14 (D), and P12 to P14 (E). Retinal sections obtained from SD rats raised in normoxia (F) and maximum hyperoxia (P0–P14, 80%; G) are also shown for comparative purposes. RPE, retinal pigment epithelium; OS, outer segment; IS, inner segment. Scale bar, 50 μm.
Figure 5.
 
Representative retinal histologic sections (central retina) from LE rats obtained at P60 from the normoxic cohort (A), and from hyperoxic cohorts including P0 to P6 (B), P0 to P14 (C), P6 to P14 (D), and P12 to P14 (E). Retinal sections obtained from SD rats raised in normoxia (F) and maximum hyperoxia (P0–P14, 80%; G) are also shown for comparative purposes. RPE, retinal pigment epithelium; OS, outer segment; IS, inner segment. Scale bar, 50 μm.
Figure 6.
 
Graphic representation of the mean retinal layer thicknesses (in micrometers) and total retinal thickness from control (P0–P14, 21%) and from hyperoxic LE cohorts, including P0 to P6, P0 to P14, P6 to P14, and P12 to P14 (n = 3 per group). Measurements were taken at P60. Significant differences within normoxic or hyperoxic cohorts were identified with a one-way ANOVA followed by the Dunnett multiple comparison test comparing data obtained from each regimen with control; *significantly different from control (P < 0.05). Results are given as the mean thickness ± 1 SD.
Figure 6.
 
Graphic representation of the mean retinal layer thicknesses (in micrometers) and total retinal thickness from control (P0–P14, 21%) and from hyperoxic LE cohorts, including P0 to P6, P0 to P14, P6 to P14, and P12 to P14 (n = 3 per group). Measurements were taken at P60. Significant differences within normoxic or hyperoxic cohorts were identified with a one-way ANOVA followed by the Dunnett multiple comparison test comparing data obtained from each regimen with control; *significantly different from control (P < 0.05). Results are given as the mean thickness ± 1 SD.
Figure 7.
 
Western blot analysis was used to detect FGF-2 and CNTF protein levels in LE (A, C, respectively) and SD (B, D, respectively) rats. Retinas were collected from normoxia-raised rats at P6 and P14 and from rats exposed to hyperoxia from P0 to P6, P6 to P14, and P0 to P14 (n = 3–4 per group). Neurotrophic factor levels were normalized to γ-actin (ordinate: ratio/γ-actin, mean ± 1 SD) for quantification. Right: molecular mass (in kilodaltons) of each band (18 kDa for FGF-2, and 19.4 kDa for CNTF). *Statistically significant differences (P < 0.05) obtained with a Student’s t-test comparing exposed animals from the P0 to P6, P6 to P14, and P0 to P14 groups to age-matched control rats.
Figure 7.
 
Western blot analysis was used to detect FGF-2 and CNTF protein levels in LE (A, C, respectively) and SD (B, D, respectively) rats. Retinas were collected from normoxia-raised rats at P6 and P14 and from rats exposed to hyperoxia from P0 to P6, P6 to P14, and P0 to P14 (n = 3–4 per group). Neurotrophic factor levels were normalized to γ-actin (ordinate: ratio/γ-actin, mean ± 1 SD) for quantification. Right: molecular mass (in kilodaltons) of each band (18 kDa for FGF-2, and 19.4 kDa for CNTF). *Statistically significant differences (P < 0.05) obtained with a Student’s t-test comparing exposed animals from the P0 to P6, P6 to P14, and P0 to P14 groups to age-matched control rats.
Figure 8.
 
Retinal cellular localization of FGF-2 (A, B) and CNTF (C, D) in representative control retinas at P6 and P14 and in exposed retinas after hyperoxia from P0 to P16, P6 to P14, and P0 to P14 in LE (A, C) and SD (B, D) rats. Upregulation of FGF-2 and CNTF staining was observed in the INL and GCL by fluorescence microscopy after hyperoxia regimens for both strains. Scale bars, 100 μm.
Figure 8.
 
Retinal cellular localization of FGF-2 (A, B) and CNTF (C, D) in representative control retinas at P6 and P14 and in exposed retinas after hyperoxia from P0 to P16, P6 to P14, and P0 to P14 in LE (A, C) and SD (B, D) rats. Upregulation of FGF-2 and CNTF staining was observed in the INL and GCL by fluorescence microscopy after hyperoxia regimens for both strains. Scale bars, 100 μm.
Figure 9.
 
Colocalization of CNTF immunostaining (A) with anti-GFAP labeling (B) specific for active Müller glial cells, revealed that Müller cells actively participate in upregulating CNTF levels after hyperoxia (shown in retinas of LE rats after maximum exposure from P0 to P14 collected at P60; C). Scale bar, 100 μm.
Figure 9.
 
Colocalization of CNTF immunostaining (A) with anti-GFAP labeling (B) specific for active Müller glial cells, revealed that Müller cells actively participate in upregulating CNTF levels after hyperoxia (shown in retinas of LE rats after maximum exposure from P0 to P14 collected at P60; C). Scale bar, 100 μm.
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Figure 1.
 
Representative retinal vascular flatmounts stained with ADPase from LE (top) and SD (bottom) rats, raised in room air and euthanatized at P6 (A, E, respectively) or P14 (C, G, respectively), or raised in hyperoxia and euthanatized immediately after 6 days (P0–P6; B, F, respectively) or P14 (P0–P14; D, H, respectively) of oxygen exposure. Magnification, ×40.
Figure 1.
 
Representative retinal vascular flatmounts stained with ADPase from LE (top) and SD (bottom) rats, raised in room air and euthanatized at P6 (A, E, respectively) or P14 (C, G, respectively), or raised in hyperoxia and euthanatized immediately after 6 days (P0–P6; B, F, respectively) or P14 (P0–P14; D, H, respectively) of oxygen exposure. Magnification, ×40.
Figure 2.
 
Graphic representation of retinal vasculature extent (A, B) and vascular density (C, D) (ordinate) as a function of the oxygen exposure regimen (days, abscissa) in the normoxic and hyperoxic LE and SD cohorts at P6 (SD, n = 3; LE, n = 3) or after 6 days of hyperoxia (P0–P6; A, C, respectively; SD, n = 3; LE, n = 3) and at P14 or after 14 days of hyperoxia (P0–P14; B, D, respectively; SD, n = 3; LE, n = 3). Results are expressed as a percentage of vascular extent or vascular density of control retinas at P6 and P14, respectively, ±1 SD. *Statistically significant differences from the control at a given age found with the Student’s t-test (P < 0.05); •, significant differences from SD rats for the same exposure regimen (P < 0.05).
Figure 2.
 
Graphic representation of retinal vasculature extent (A, B) and vascular density (C, D) (ordinate) as a function of the oxygen exposure regimen (days, abscissa) in the normoxic and hyperoxic LE and SD cohorts at P6 (SD, n = 3; LE, n = 3) or after 6 days of hyperoxia (P0–P6; A, C, respectively; SD, n = 3; LE, n = 3) and at P14 or after 14 days of hyperoxia (P0–P14; B, D, respectively; SD, n = 3; LE, n = 3). Results are expressed as a percentage of vascular extent or vascular density of control retinas at P6 and P14, respectively, ±1 SD. *Statistically significant differences from the control at a given age found with the Student’s t-test (P < 0.05); •, significant differences from SD rats for the same exposure regimen (P < 0.05).
Figure 3.
 
Representative scotopic (mixed rod-cone, flash intensity: −6.3 through 0.60 log cd · s · m−2; AE) and photopic (flash intensity: 0.9 log cd · s · m−2, background: 30 cd · m−2; (G) ERGs along with scotopic (F) and photopic (H) oscillatory potentials obtained at P60 from animals raised in normoxia (21% O2) and hyperoxia (80% O2) from P0 to P6, P0 to P14, P6 to P14, and P12 to P14. Calibrations: horizontal, 20 ms; vertical, 400 μV for scotopic mixed rod-cone ERG responses, 200 μV for photopic ERG responses and 40 μV for scotopic and photopic oscillatory potentials, respectively. All tracings include a 20-ms prestimulus baseline; arrows, flash onset.
Figure 3.
 
Representative scotopic (mixed rod-cone, flash intensity: −6.3 through 0.60 log cd · s · m−2; AE) and photopic (flash intensity: 0.9 log cd · s · m−2, background: 30 cd · m−2; (G) ERGs along with scotopic (F) and photopic (H) oscillatory potentials obtained at P60 from animals raised in normoxia (21% O2) and hyperoxia (80% O2) from P0 to P6, P0 to P14, P6 to P14, and P12 to P14. Calibrations: horizontal, 20 ms; vertical, 400 μV for scotopic mixed rod-cone ERG responses, 200 μV for photopic ERG responses and 40 μV for scotopic and photopic oscillatory potentials, respectively. All tracings include a 20-ms prestimulus baseline; arrows, flash onset.
Figure 4.
 
Graphic representation of a-wave (A, G), rod V max (B, H), rod-cone b-wave (C, I), photopic b-wave (D, J), scotopic SOP (E, K), and photopic SOP (F, L) amplitudes (ordinate in microvolts) obtained at P30 and P60, respectively (abscissa in days of exposure), from LE and SD rats. The hyperoxic cohort of each strain was divided at birth into seven different exposure protocols: from birth through postnatal day 6 (P0–P6: LE, n = 24 [P30], n = 5 [P60]; SD, n = 6 [P30], n = 6 [P60]); birth to postnatal day 9 (P0–P9: LE, n = 11 [P30], n = 5 [P60]); birth to postnatal day 12 (P0–P12: LE, n = 7 [P30], n = 6 [P60]); and birth to postnatal day 14 (P0–14, LE, n = 21 [P30], n = 6 [P60]; SD, n = 11 [P30], n = 7 [P60]) or from P6, P9, or P12 through P14 (P6–P14: LE, n = 22 [P30], n = 6 [P60]; SD, n = 8 [P30], n = 8 [P60]; P9–P14: LE, n = 17 [P30], n = 9 [P60]; P12–14: LE, n = 15 [P30], n = 7 [P60]). Data were compared with results for respective age-matched control litters raised in normoxia (LE, n = 21; SD, n = 22).
Figure 4.
 
Graphic representation of a-wave (A, G), rod V max (B, H), rod-cone b-wave (C, I), photopic b-wave (D, J), scotopic SOP (E, K), and photopic SOP (F, L) amplitudes (ordinate in microvolts) obtained at P30 and P60, respectively (abscissa in days of exposure), from LE and SD rats. The hyperoxic cohort of each strain was divided at birth into seven different exposure protocols: from birth through postnatal day 6 (P0–P6: LE, n = 24 [P30], n = 5 [P60]; SD, n = 6 [P30], n = 6 [P60]); birth to postnatal day 9 (P0–P9: LE, n = 11 [P30], n = 5 [P60]); birth to postnatal day 12 (P0–P12: LE, n = 7 [P30], n = 6 [P60]); and birth to postnatal day 14 (P0–14, LE, n = 21 [P30], n = 6 [P60]; SD, n = 11 [P30], n = 7 [P60]) or from P6, P9, or P12 through P14 (P6–P14: LE, n = 22 [P30], n = 6 [P60]; SD, n = 8 [P30], n = 8 [P60]; P9–P14: LE, n = 17 [P30], n = 9 [P60]; P12–14: LE, n = 15 [P30], n = 7 [P60]). Data were compared with results for respective age-matched control litters raised in normoxia (LE, n = 21; SD, n = 22).
Figure 5.
 
Representative retinal histologic sections (central retina) from LE rats obtained at P60 from the normoxic cohort (A), and from hyperoxic cohorts including P0 to P6 (B), P0 to P14 (C), P6 to P14 (D), and P12 to P14 (E). Retinal sections obtained from SD rats raised in normoxia (F) and maximum hyperoxia (P0–P14, 80%; G) are also shown for comparative purposes. RPE, retinal pigment epithelium; OS, outer segment; IS, inner segment. Scale bar, 50 μm.
Figure 5.
 
Representative retinal histologic sections (central retina) from LE rats obtained at P60 from the normoxic cohort (A), and from hyperoxic cohorts including P0 to P6 (B), P0 to P14 (C), P6 to P14 (D), and P12 to P14 (E). Retinal sections obtained from SD rats raised in normoxia (F) and maximum hyperoxia (P0–P14, 80%; G) are also shown for comparative purposes. RPE, retinal pigment epithelium; OS, outer segment; IS, inner segment. Scale bar, 50 μm.
Figure 6.
 
Graphic representation of the mean retinal layer thicknesses (in micrometers) and total retinal thickness from control (P0–P14, 21%) and from hyperoxic LE cohorts, including P0 to P6, P0 to P14, P6 to P14, and P12 to P14 (n = 3 per group). Measurements were taken at P60. Significant differences within normoxic or hyperoxic cohorts were identified with a one-way ANOVA followed by the Dunnett multiple comparison test comparing data obtained from each regimen with control; *significantly different from control (P < 0.05). Results are given as the mean thickness ± 1 SD.
Figure 6.
 
Graphic representation of the mean retinal layer thicknesses (in micrometers) and total retinal thickness from control (P0–P14, 21%) and from hyperoxic LE cohorts, including P0 to P6, P0 to P14, P6 to P14, and P12 to P14 (n = 3 per group). Measurements were taken at P60. Significant differences within normoxic or hyperoxic cohorts were identified with a one-way ANOVA followed by the Dunnett multiple comparison test comparing data obtained from each regimen with control; *significantly different from control (P < 0.05). Results are given as the mean thickness ± 1 SD.
Figure 7.
 
Western blot analysis was used to detect FGF-2 and CNTF protein levels in LE (A, C, respectively) and SD (B, D, respectively) rats. Retinas were collected from normoxia-raised rats at P6 and P14 and from rats exposed to hyperoxia from P0 to P6, P6 to P14, and P0 to P14 (n = 3–4 per group). Neurotrophic factor levels were normalized to γ-actin (ordinate: ratio/γ-actin, mean ± 1 SD) for quantification. Right: molecular mass (in kilodaltons) of each band (18 kDa for FGF-2, and 19.4 kDa for CNTF). *Statistically significant differences (P < 0.05) obtained with a Student’s t-test comparing exposed animals from the P0 to P6, P6 to P14, and P0 to P14 groups to age-matched control rats.
Figure 7.
 
Western blot analysis was used to detect FGF-2 and CNTF protein levels in LE (A, C, respectively) and SD (B, D, respectively) rats. Retinas were collected from normoxia-raised rats at P6 and P14 and from rats exposed to hyperoxia from P0 to P6, P6 to P14, and P0 to P14 (n = 3–4 per group). Neurotrophic factor levels were normalized to γ-actin (ordinate: ratio/γ-actin, mean ± 1 SD) for quantification. Right: molecular mass (in kilodaltons) of each band (18 kDa for FGF-2, and 19.4 kDa for CNTF). *Statistically significant differences (P < 0.05) obtained with a Student’s t-test comparing exposed animals from the P0 to P6, P6 to P14, and P0 to P14 groups to age-matched control rats.
Figure 8.
 
Retinal cellular localization of FGF-2 (A, B) and CNTF (C, D) in representative control retinas at P6 and P14 and in exposed retinas after hyperoxia from P0 to P16, P6 to P14, and P0 to P14 in LE (A, C) and SD (B, D) rats. Upregulation of FGF-2 and CNTF staining was observed in the INL and GCL by fluorescence microscopy after hyperoxia regimens for both strains. Scale bars, 100 μm.
Figure 8.
 
Retinal cellular localization of FGF-2 (A, B) and CNTF (C, D) in representative control retinas at P6 and P14 and in exposed retinas after hyperoxia from P0 to P16, P6 to P14, and P0 to P14 in LE (A, C) and SD (B, D) rats. Upregulation of FGF-2 and CNTF staining was observed in the INL and GCL by fluorescence microscopy after hyperoxia regimens for both strains. Scale bars, 100 μm.
Figure 9.
 
Colocalization of CNTF immunostaining (A) with anti-GFAP labeling (B) specific for active Müller glial cells, revealed that Müller cells actively participate in upregulating CNTF levels after hyperoxia (shown in retinas of LE rats after maximum exposure from P0 to P14 collected at P60; C). Scale bar, 100 μm.
Figure 9.
 
Colocalization of CNTF immunostaining (A) with anti-GFAP labeling (B) specific for active Müller glial cells, revealed that Müller cells actively participate in upregulating CNTF levels after hyperoxia (shown in retinas of LE rats after maximum exposure from P0 to P14 collected at P60; C). Scale bar, 100 μm.
Table 1.
 
Group Data Obtained in Scotopic and Photopic Conditions from LE Rats Raised in Normoxia and Hyperoxia
Table 1.
 
Group Data Obtained in Scotopic and Photopic Conditions from LE Rats Raised in Normoxia and Hyperoxia
Age Control P0–P6 P0–P9 P0–P12 P0–P14 P6–P14 P9–P14 P12–P14
Scotopic parameters
 a-Wave (μV) 30 d 264.34 ± 87.25 256.41 ± 85.60 223.34 ± 77.82 166.76 ± 67.68* 137.18 ± 44.29* 167.65 ± 64.47* 223.42 ± 89.78 282.08 ± 130.44
60 d 220.29 ± 58.84 185.80 ± 59.51 182.15 ± 56.72 175.00 ± 100.36 100.49 ± 27.73 94.57 ± 61.72* , † 217.62 ± 121.44 202.48 ± 153.34
 Rod V max (μV) 30 d 497.58 ± 124.50 422.54 ± 123.10 202.82 ± 96.03* 79.90 ± 74.54* 81.63 ± 59.71* 90.32 ± 58.76* 173.61 ± 105.87* 341.60 ± 96.67*
60 d 544.61 ± 178.97 405.44 ± 109.12 209.85 ± 138.10* 186.29 ± 127.92* 148.79 ± 98.49* 70.99 ± 42.08* 135.44 ± 119.43* 308.85 ± 101.74*
 Rod-cone b-wave (μV) 30 d 864.19 ± 193.40 593.61 ± 200.20* 277.20 ± 137.83* 133.00 ± 91.77* 139.84 ± 58.11* 187.31 ± 68.79* 362.09 ± 269.4* 680.97 ± 304.62*
60 d 806.76 ± 257.07 483.50 ± 217.97 281.00 ± 204.02* 184.08 ± 153.75* 200.32 ± 122.22* 142.08 ± 46.91* 468.23 ± 350.81* 598.27 ± 423.08*
 SOPs (μV) 30 d 388.82 ± 100.64 229.23 ± 91.31* 99.52 ± 48.02* 51.05 ± 40.36* 24.28 ± 20.95* 51.28 ± 45.03* 93.99 ± 88.76* 277.80 ± 177.02*
60 d 395.69 ± 94.98 271.81 ± 142.43 155.79 ± 121.60* 81.75 ± 78.20* 50.14 ± 47.45* 44.91 ± 37.73* 139.43 ± 108.68* 233.38 ± 185.07*
Photopic parameters
 b-Wave (μV) 30 d 209.61 ± 72.44 111.19 ± 40.21* 49.06 ± 35.97* 31.41 ± 13.51* 25.02 ± 10.79* 42.87 ± 27.22* 67.41 ± 53.26* 126.59 ± 71.18*
60 d 190.29 ± 52.24 98.22 ± 40.65* 63.65 ± 58.33* 42.42 ± 23.21* 38.68 ± 21.13* , † 35.50 ± 23.65* 89.44 ± 64.71* 147.44 ± 102.81
 SOPs (μV) 30 d 55.88 ± 21.24 42.89 ± 16.92 19.42 ± 17.35* 7.71 ± 7.99* 4.20 ± 6.28* 11.11 ± 14.34* 28.97 ± 32.93* 43.67 ± 25.46
60 d 62.93 ± 17.79 36.51 ± 20.08* 34.08 ± 20.46* 22.8 ± 11.96* , † 13.37 ± 10.57* , † 9.87 ± 6.67* 25.67 ± 15.67* 38.91 ± 30.77
Table 2.
 
Group Data Obtained in Scotopic and Photopic Conditions from SD Rats Raised in Normoxia and Hyperoxia
Table 2.
 
Group Data Obtained in Scotopic and Photopic Conditions from SD Rats Raised in Normoxia and Hyperoxia
Age Control P0–P6 P0–P14 P6–P14
Scotopic parameters
 a-Wave (μV) 30 d 361.07 ± 109.65* 315.30 ± 50.70 287.65 ± 69.38* 305.27 ± 70.41*
60 d 326.35 ± 57.68* 337.68 ± 85.41* 214.32 ± 82.09* 357.36 ± 96.43*
 Rod V max (μV) 30 d 496.25 ± 146.35 366.03 ± 66.63, † 123.15 ± 92.98, † 201.19 ± 138.68* , †
60 d 529.24 ± 85.92 392.08 ± 90.25 138.08 ± 137.25, † 232.96 ± 114.33* , †
 Rod-cone b-wave (μV) 30 d 879.35 ± 226.06 656.83 ± 122.92, † 324.79 ± 116.96* , † 430.05 ± 164.67* , †
60 d 766.74 ± 107.27, ‡ 661.67 ± 160.81 309.18 ± 248.22, † 472.25 ± 102.94* , †
 SOPs (μV) 30 d 574.01 ± 297.05* 353.63 ± 81.25* 104.86 ± 69.00* , † 225.95 ± 133.51* , †
60 d 470.19 ± 237.24 329.01 ± 92.99 96.44 ± 89.80, † 244.39 ± 54.7* , †
Photopic parameters
 b-Wave (μV) 30 d 209.25 ± 72.92 128.05 ± 33.09, † 57.18 ± 34.09* , † 87.99 ± 42.61* , †
60 d 160.49 ± 30.08, ‡ 143.67 ± 38.34 52.43 ± 44.79, † 91.72 ± 19.58* , †
 SOPs (μV) 30 d 73.05 ± 18.91* 51.91 ± 7.57 19.15 ± 25.35* , † 34.81 ± 25.96, †
60 d 59.47 ± 40.31 55.94 ± 18.96 9.64 ± 7.66, † 41.28 ± 23.73*
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