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1From the Departments of Anatomy and Cell Biology and 3Ophthalmology, Wayne State University, Detroit, Michigan; and 2Max-Planck-Institut für Experimental Endokrinologie, Hannover, Germany.
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METHODS. Retinal oxygenation response (
PO2) was measured using functional magnetic resonance imaging (MRI) and either carbogen or 100% oxygen inhalation challenge in C57BL/6J control (C) mice. Retinal PO2 during carbogen breathing was also measured in PKCß knockout (C57BL6-Prkcb1; [KO]), 4 month C57BL/6J diabetic (D), and 4-month diabetic PKCß KO (D+KO) mice. Retinal PKCß protein expression was assessed by Western analysis.
RESULTS. In C mice, retinal PO2 during carbogen breathing was significantly greater (P < 0.05) than during oxygen breathing. In D mice, PO2 during carbogen breathing was significantly lower than normal in the superior, but not the inferior, hemiretina and was normal (P > 0.0 5) in the KO group. In the D+KO mice PO2 was normal (P > 0.05) only at distances less than 1.5 mm from the optic nerve head. PKCß expression was elevated in the retina in diabetes (P < 0.05), but was significantly decreased (P < 0.05) in D+KO mice.
CONCLUSIONS. The present study confirms that, in the mouse, retinal PO2 patterns during different inhalation challenges or in the presence of diabetes are similar to what has been reported in rats. Diabetes-induced elevation of PKC appears to contribute only focally to subnormal retinal PO2. This raises the possibility that PKC inhibition therapy may be only regionally effective in treating retinal pathophysiology associated with diabetic retinopathy.
We have developed a novel functional magnetic resonance imaging (MRI) method for measuring the retinal oxygenation response to a hyperoxic inhalation challenge in the newborn and adult rat, rabbit, cat, and human.6 7 8 9 10 11 In this technique, hyperoxia-induced increases in vitreous partial oxygen pressure over the room-air value (PO2) are measured as increased signal intensity. Good agreement is found between the MRI-measured response and that derived from oxygen electrode data in normal rat retina under similar conditions.12 13 In control rats, during a carbogen challenge, oxygen from the choroid is mostly consumed by the inner plexiform layer and does not enter the preretinal vitreous space to a substantial degree.12 14 A number of interrelated hemodynamic parameters can modulate oxygen supply during a carbogen challenge, including retinal vessel autoregulation, perfusion reserve, ocular perfusion pressure, plasma pH, and vascular density. In normal adult and newborn rats, carbogen breathing oxygenates the retina significantly better than pure oxygen breathing.13 Carbogen, a gas mixture of carbon dioxide (5%) and oxygen (95%), is used clinically instead of 100% oxygen to minimize the vasoconstrictive effects of pure O2 on retinal blood flow and oxygenation.
Application of this functional MRI method in rat models of diabetic retinopathy has revealed that the superior hemiretina exhibits a lesser (i.e., subnormal) oxygenation response than is found in control (i.e., normal) animals and that this abnormality appears several months before retinal histopathology becomes apparent.7 15 In principle, during a carbogen challenge, reduced oxygen supplied by the retinal circulation and/or increased retinal oxygen consumption could produce a subnormal response. However, a subnormal oxygenation response does not appear to represent retinal hypoxia.16 Because retinal oxygen demand may decrease in experimental diabetes,17 18 our working hypothesis is that measurement of a subnormal retinal PO2 is primarily a measure of a decrease in oxygen supplied by the retinal circulation.6 7 8 15
Aminoguanidine (AMG), a compound shown to inhibit retinal PKC activity in diabetes, preserved the retinal oxygenation response at the 3-month time point and prevented the development of retinal histopathology at 15 months of diabetes in rats.2 15 19 In contrast, AMG treatment of galactose-fed rats did not protect the retinal oxygenation response at 3 months, nor did it prevent the development of retinal lesions at 15 months.7 15 19 20 Galactose-fed rats treated with an aldose reductase inhibitor also preserved the retinal oxygenation response and prevented development of retinal lesions.15 Taken together, these data underscore the use of retinal PO2 measurements as a potential surrogate marker of the efficacy of treatment in diabetic retinopathy.
In this study, we examined whether in the mouse retinal PO2 patterns during breathing of carbogen and oxygen as well as in diabetes were similar to those in the rat.7 13 15 In addition, we began to test the hypothesis that diabetes-induced elevation of PKCß in the retina contributes to the development of an early subnormal PO2. Retinal PO2 was compared in control, diabetic, and diabetic PKCß KO mice. Diabetic mice exhibited early biochemical changes, including elevation of PKC activity, similar to those in diabetic rats.1 Furthermore, in long-term (>15 months) experimental diabetes in mice, retinal histopathology (acellular capillaries and pericyte ghosts) developed that is similar to that observed in lesions in long-term diabetes in rats and early in the course of diabetes in humans.21 22 23
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Animal Models
Control (C57BL/6J, C group) mice were purchased from Jackson Laboratories (Bar Harbor, ME) and PKC mice with targeted disruption of the PKCß gene (KO group) with a C57J background (C57BL6-Prkcb1) were provided by one of the authors (ML).24 The KO mice have a similar lifespan as the C57BL/6J (Leitges M, personal communication, December 2000).
Diabetes was induced in the C57BL/6J and KO mice (starting weight, 20–24 g) by streptozotocin (65 mg/kg; 50 mM citrate buffer [pH 4.5]) intraperitoneal injection once a day for 5 days. Body weight, blood glucose, and urine ketones levels were monitored weekly. Insulin (neutral protamine Hagedorn [NPH]) was administered to allow slow weight gain while maintaining hyperglycemia (blood glucose levels above 400 mg/dL). Normal rodent chow and water were provided ad libitum. After 4 months of diabetes, only those mice with fasting blood glucose levels higher than 400 mg/dL (D group) were studied by functional MRI.
Western Blot Analysis
Retinal tissue was homogenized in 30 mM Tris-HC buffer containing 10 mM EGTA, 5 mM EDTA, 1% Triton X-100, and 250 mM sucrose containing 1 mM NaF, 1 mM phenylmethylsulfonyl fluoride, 15 µg/mL aprotinin, 5 µg/mL leupeptin, 5 µg/mL pepstatin, and 1 mM Na3VO4. The homogenate was centrifuged at 5000g for 5 minutes at 4°C to remove cell debris. The protein concentration was determined with the bicinchoninic acid assay (BCA; Sigma-Aldrich, St. Louis, MO) with bovine serum albumin used as the standard. Retinal protein (30–40 µg) was separated on a 10% reducing polyacrylamide gel and then transferred to nitrocellulose membranes. The membranes were blocked in the wash buffer (10 mM Tris-HCl [pH 7.5], 100 mM NaCl and 0.1% Tween 20), containing 5% milk, followed by the incubation with a polyclonal antibody against PKCß (catalog no. sc-210; Santa Cruz Biotechnology, Santa Cruz, CA) at a dilution of 1:1000. The membranes were washed three times (10 minutes each), and incubated for 1 hour at room temperature with horseradish-peroxidase–linked anti-rabbit Ig (Amersham Pharmacia Biotech, Amersham, UK; 1:2500 dilution), and developed with a Western blot kit (ECL-Plus; Amersham Pharmacia Biotech).
To ensure equal loading among lanes, the expression of the intrinsic protein ß-actin was quantified in each lane. The intensities of PKCßII band and ß-actin band in each lane were quantified with an automated gel digitizing system (Un-Scan-It; Silk Scientific, Orem, UT). After blots were transferred to the membranes for detection of PKC, the membranes were incubated with stripping buffer (62.5 mM Tris-HCl [pH 6.8], 100 mM mercaptoethanol, and 2% sodium dodecyl sulfate) at 50°C for 30 minutes, washed, and incubated with antibody against ß-actin (Sigma-Aldrich). The secondary antibody used was anti-mouse IgG horseradish-peroxidase–conjugated antibody, and the membranes were developed with the Western blot kit (ECL-Plus; Amersham). Each sample was analyzed at least in duplicate.
Functional MRI Examination
On the day of the experiment, anesthesia was induced by a single intraperitoneal injection of urethane (36% solution, 0.083 mL/20 g animal weight, prepared fresh daily; Aldrich, Milwaukee, WI). Each mouse was gently positioned on an MRI-compatible custom-made holder with its nose placed in a plastic nose cone. Animals were allowed to breathe spontaneously during the experiment. To maintain the core temperature, a recirculating heated water blanket was used. Rectal temperature was continuously monitored while the animal was inside the magnet.15
MRI data were acquired on a 4.7-T system using a two-turn transmit-and-receive surface coil (1.0 cm diameter) placed over the eye, as previously described6 (repetition time [TR], 1 second; echo time [TE], 22.7 ms; number of acquisitions [NA], 1; matrix size, 128 x 256; slice thickness, 1 mm; field of view, 18 x 18 mm; sweep width, 25,000 Hz; 2 minutes per image). Examining the initial retinal oxygenation response to a hyperoxic challenge requires careful attention to experimental timing and makes high demands on the animals’ physiology, compared with steady state measurements.
The functional MRI data were collected as follows. Sagittal localizer images were first collected and used to position a single 1-mm-thick transverse slice through the center of the eye. The 1-mm slice thickness was needed to obtain an adequate signal-to-noise ratio in a 2-minute image. This slice thickness resulted in some partial volume averaging, so that the final image contained superior and inferior hemiretina, with a relatively minor contribution from the temporal and nasal hemiretina. A capillary tube (1.5-mm inner diameter) filled with distilled water was used as the external standard.
It is important to note that steady state (room air) vitreous oxygen tension cannot be measured by this method, because many factors affect the preretinal vitreous water signal and its relaxation properties. In other words, simply obtaining an image of the eye during room air breathing alone cannot be used to measure retinal oxygenation. In addition, detailed inferences regarding oxygen consumption and delivery in individual retinal layers cannot be made, because functional MRI measures vitreal oxygenation and so reflects general treads in intraretinal oxygenation.14 Nonetheless, as discussed in the introduction, retinal PO2 have shown promise as a biomarker of treatment efficacy in diabetic retinopathy.
In general, PO2 data were collected approximately 90 minutes after urethane injection to avoid potential errors due to variable time under urethane anesthesia. Three images were acquired while the animal breathed room air and one image during the inhalation of carbogen or 100% oxygen. The hyperoxic gas exposure was started at the end of the third image. Animals were returned to room air for 5 minutes to allow recovery from the inhalation challenge and were removed from the magnet. A second 2-minute inhalation challenge was performed outside the magnet with care taken to avoid altering the spatial relationship between the animal head and nose cone. At exactly 2 minutes, arterial blood from the abdominal aorta was collected as described previously.8 This blood was analyzed for glucose, percentage of glycated hemoglobin, PaO2, PaCO2, and pH. However, because the length of time that the blood was stored before the percent glycated hemoglobin assay, the data produced were not reproducible and so are not reported. Note that this second inhalation challenge (outside the magnet) is needed because it is not feasible to obtain routinely an arterial blood sample from inside the magnet (>40 cm away from the magnet opening) from 25 g mice. In all cases, after the blood collection, the mice were killed with an intracardiac potassium chloride injection and eyes enucleated for Western blot analysis.
Data Analysis
To be included in this study, the animal must have demonstrated (1) minimal eye movement during the MRI examination, (2) a nongasping respiratory pattern before the MRI examination, (3) rectal temperature in the range of 35.5 to 36.5°C, and (4) PaO2 higher than 350 mm Hg and PaCO2 between 46 and 65 mm Hg during the carbogen challenge. Occasionally, the blood gas machine was not able to read a sample (e.g., due to a clot or excessive air in the capillary tube). In this case, the MRI data were also excluded. Between 5 to 20 animals in each of the different treatment groups satisfied the fourth inclusion criterion.
To correct for possible subtle shifting of the animals’ position during the experiment (usually for reasons unrelated to the physiology of the animal, such as settling on the gauze packing) a warp affine image coregistration was performed on each animal with software written in-house. After coregistration, the MRI data were transferred to a computer (Power Mac G4; Apple Computer, Cupertino, CA) and analyzed with the software program Image (available by ftp at zippy.nimh.nih.gov/ or at http://rsb.info.nih.gov/nih-image; developed by Wayne Rasband, National Institutes of Health, Bethesda, MD). Images obtained during room air breathing were averaged to improve the signal-to-noise ratio. All pixel signal intensities in the average-room-air image and the 2-minute carbogen image were then normalized to the external standard intensity. Signal intensity changes during carbogen breathing were calculated and converted to PO2 values, on a pixel-by-pixel basis, as follows.13 For each pixel, the fractional signal enhancement (E) was calculated
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PO2 according to a previously established theory13
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, where Tr is the repetition time, and T10 is the T1 in the absence of oxygen. Using a Tr of 1 second and assuming a vitreous T10 of 4 seconds, Tk = 3.52. This T10 value is based on our previous measurement of the proton spin–lattice relaxation time in the rabbit vitreous (4 seconds) and reported values in human vitreous (3.3 seconds) and in cerebral spinal fluid (4.3 seconds), which has a high water content similar to that of vitreous.13 25 An R1 of 2 x 10-4 sec-1 · mm Hg-1 was used.13 This R1 was previously measured in a saline phantom, which is assumed to be a reasonable model of vitreous (98% water).13 A similar R1 value was found for plasma, suggesting that relatively low protein levels do not substantially contribute to oxygen relaxivity.26 Note that an E of 0.01 (i.e., a 1% signal intensity change) corresponds to a PO2 of 14 mm Hg.
The PO2 parameter image was analyzed as follows: First, the pixel value along a 1-pixel-thick line at the boundary of the retina-choroid and vitreous was set to 255 (black). We estimate that the thickness of this line, based on the in-plane resolution (18 x 18 mm2 field of view was sampled by 128 x 256 data points) of 140 x 70 µm, is approximately 100 µm. The values in another 1-pixel-thick line drawn in the preretinal vitreous next to the black pixels were then extracted.6 This procedure minimized retina–choroid pixel values from potentially contaminating those used in the final analysis ("pixel bleed") and insured that similar preretinal vitreous space was sampled for each animal. In addition, spatial averaging over these 100-µm regions of interest tends to minimize the contribution from the very local preretinal oxygenation gradients next to the retinal surface.27 An average PO2 band was constructed based on the within-group mean for each pixel. Note that due to the size of our in-plane pixels (approximately 100 µm) and slice thickness, changes in oxygen from regions less than 100 µm, such as a particular retinal artery, cannot be measured. However, inhomogeneous distributions of retinal oxygenation responses on a scale of more than 100 µm are readily observed, when present.
Statistical Analysis
The physiological parameters (i.e., blood gas values, rectal temperatures, blood glucose data) were normally distributed and are presented as the mean ± SEM. Comparisons were performed with an unpaired t-test. Separate comparisons of retinal PO2 between the C and D groups and between the KO and D+KO groups were performed with a generalized estimating equation. This method performs a general linear regression analysis using all the pixels in each subject and accounts for the within-subject correlation between adjacent pixels. When multiple comparisons were made, the likelihood of finding a significant impact for all the comparisons by chance alone is P 
0.0167 (0.05/(number of comparisons made) or 0.05/3 = 0.0167). When two samples were compared, P 0.05 was considered significant.
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PO2 during carbogen breathing (Fig. 2) better than the small PaO2 difference (approximately 10%). Compared with C-group mice, all D-group animals had significantly (P < 0.05) elevated blood glucose levels. There were no significant (P > 0.05) differences in plasma glucose levels between the D and D+KO groups. The PaO2, pH, and core temperature were not significantly (P > 0.05) different among any of the carbogen-breathing mouse groups (Table 1) .
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PO2 = 142 ± 4, n = 20) produced a larger (P < 0.05) oxygenation response than during oxygen inhalation (panretinal PO2 = 78 ± 5, n = 6; Fig. 2 ). Further experiments in the present study were performed with carbogen gas to best detect subtle differences in retinal oxygenation between groups.
Figure 3 summarizes the retinal PO2 data in all groups. No significant (P > 0.05) differences were found between the superior and inferior hemiretinal PO2 of the control and PKC KO mice (i.e., no hemiretinal asymmetry) or between the inferior hemiretinal PO2 in all the groups. A significantly (P < 0.05) subnormal average superior hemiretinal PO2 was found only in the D group.
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PO2 versus distance from the optic nerve head for the C, D, KO, and D+KO groups. No significant differences (P > 0.05) were found between the inferior hemiretinal PO2 in any of these groups (data not shown). Although the pansuperior hemiretinal PO2 of the D+KO mice was not subnormal compared with the KO group (P > 0.05, Fig. 3 ), retinal PO2 was normal only in a region less than 1.5 mm from the optic nerve head.
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PO2) after 4 months of diabetes in the mouse that was similar to that in the rat with 3 months of experimental diabetes.7 15 This pathophysiology occurs earlier than the reported development of retinal vascular lesions (acellular capillaries and pericyte ghosts appear at later times [>15 months] in experimental diabetes in mice).21 22 23 The mechanism(s) underlying the early superior pathophysiology are not known. It is possible that the present retinal oxygenation differences between the C and D groups were the result of systemic variations between these groups. However, no significant differences in core temperature and blood gas parameters were found (Table 1) . Although we did not measure other systemic physiologic parameters, such as blood pressure, it seems reasonable to expect that systemic physiologic differences would similarly affect both the inferior and superior hemiretinal oxygenation response. The lack of differences between the inferior hemiretinal PO2 in the C and D groups does not support this possibility. Based on these considerations, it appears that the inability of the superior hemiretina to oxygenate in the D group is due to local (i.e., retinal) and not systemic factors.
We have not yet identified any mechanisms that can explain why this early pathophysiology appears in superior but not inferior hemiretina of diabetic rats. In control and KO mice, no superior-inferior hemiretinal PO2 asymmetry was detected (Fig. 3) . One possibility is that early histopathology may preferentially occur in superior hemiretina of diabetic rats. However, to the best of our knowledge, there are no reports of early changes in histology in the superior hemiretina preferentially. However, the relatively small number of lesions in the diabetic retina at more than 15 months prevents finding a statistical answer to this question (Kern T, unpublished data, 2000). Also, superior–inferior differences in choroidal circulation have not been reported, although in the rat, electrode data suggest that oxygen from the choroid is mostly consumed by the photoreceptors and does not enter the preretinal vitreous space.12 The rodent superior hemiretina is reported to contain a greater number of ganglion cells than the inferior hemiretina and so would be expected to have a relatively greater metabolic demand.28 We speculate that this greater metabolic demand may make the superior hemiretina more susceptible to a hyperglycemic insult than the inferior hemiretina. Because control and diabetic animals are handled identically throughout the data collection and analysis aspects of the experiment, factors, such as differences in geometry, are similar for both groups of animals. Note that after 15 months of galactose feeding in rats, both superior and inferior hemiretina are subnormal, suggesting that the early superior hemiretinal subnormal response indicates that perhaps the superior region is more susceptible to diabetes-induced damage.7 The present data in mice, and results from our previous studies in diabetic and galactose-fed rats, highlights a defect in the superior hemiretinal PO2 as an early marker of diabetes-related histopathology.
Defining the physiology underlying retinal PO2 remains an area of on-going research. To help clarify interpretation of altered PO2, we note first that during steady state (room air breathing) conditions, retinal oxygen demand and supply are matched so that all the oxygen delivered to the retina is consumed and none is dumped into the preretinal vitreous. In this case, inner retinal PO2 is normally maintained by appropriate retinal vasoreactivity over a broad range of ocular perfusion pressures and retinal oxygen consumption rates. However, the retinal circulation is not usually at steady state. Instead, retinal oxygen supply and demand constantly adapt during daily activities. PO2 is believed to be a measure of how well retinal oxygen supply works. If baseline PO2 is either higher or lower than normal but PO2 is normal, the implication is that the circulation will be able to compensate for the baseline abnormality. A subnormal PO2 (i.e., an inability to match oxygen supply with demand) is expected to have long-term negative consequence for the health of the retina.
In this study, we investigated the contribution of a specific diabetes-induced biochemical abnormality, increased PKCß activity, to the PO2 differences between control and D groups.1 The results of the current work, and the results of others,2 indicate that retinal PKC expression is elevated before the appearance of retinal histopathology in diabetic and galactose-fed rats and mice (Fig. 1) .1 Increased PKCß activity can affect retinal circulatory physiology. For example, drug inhibition of PKCß has been reported to prevent the decreased retinal perfusion measured in the first couple of weeks after conversion to diabetes,5 attenuate the increase in leukocyte entrapment in the retinal microcirculation during the first 4-week diabetic period in rats,29 and inhibit preretinal and optic nerve head neovascularization in a nondiabetic branch retinal vein occlusion pig model.30 As discussed previously, retinal PO2 is expected to reflect the contributions of a variety of vascular physiological processes, such as vessel autoregulation, that govern the functional retinal oxygenation response during the carbogen challenge.6 The relationship between decreased steady state perfusion of the retina (as measured using dye dilution methods, for example) and a subnormal functional response to an inhalation provocation has not yet been investigated. We hypothesized that in 4-month D mice, elevated PKCß contributed to the development of a subnormal PO2 in experimental diabetic retinopathy.
The results of this study only partially supported this hypothesis. The reported elevated retinal PKCß activity appears to be important to the development of a subnormal PO2 in central retina of D mice. To the best of our knowledge there is no literature that maps PKC activity across the mouse retina in normal and KO mice and more work is needed to chart regional PKC in the mouse eye. Our Western blot data show that the protein expression of PKCß in the retina was decreased by 60% in the KO mice and 50% in the D+KO mice compared with the control mice. This could suggest that the deletion of the gene for PKCß was incomplete in the retina, or that the polyclonal antibody used for PKCß was reacting with some other proteins in the retina. However, given that these mice were homozygous for a targeted disruption,24 the possibility of incomplete PKC knockout in the retina seems unlikely. It remains to be determined whether or not chronic inhibition of PKCß in nongenetically altered rodents will correct central retina oxygenation response defects or, more important, prevent long-term retinal histopathology. Nonetheless, these considerations raise the possibility, for the first time, that specific PKCß inhibition treatment may only be regionally effective in treating retinal pathophysiology associated with diabetic retinopathy.
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Acknowledgements |
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Submitted for publication September 11, 2003; revised November 21, 2003; accepted December 1, 2003.
Disclosure: H. Luan, None; M. Leitges, None; R.R. Gupta, None; D. Pacheco, None; A. Seidner, None; J. Liggett, None; Y. Ito, None; R. Kowluru, None; B.A. Berkowitz, 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: Bruce A. Berkowitz, Department of Anatomy and Cell Biology, Wayne State University School of Medicine, 540 E. Canfield, Detroit, MI 48201; baberko{at}med.wayne.edu.
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PO2 mapping. Invest Ophthalmol Vis Sci. 1996;37:2089–2098.PO(2) predicts therapeutic efficacy in experimental diabetic retinopathy. Invest Ophthalmol Vis Sci. 2001;42:2964–2969.This article has been cited by other articles:
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  S. H. Hardarson, A. Harris, R. A. Karlsson, G. H. Halldorsson, L. Kagemann, E. Rechtman, G. M. Zoega, T. Eysteinsson, J. A. Benediktsson, A. Thorsteinsson, et al. Automatic Retinal Oximetry Invest. Ophthalmol. Vis. Sci., November 1, 2006; 47(11): 5011 - 5016. [Abstract] [Full Text] [PDF]
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R. Roberts, H. Luan, and B. A. Berkowitz {alpha}-Lipoic Acid Corrects Late-Phase Supernormal Retinal Oxygenation Response in Experimental Diabetic Retinopathy. Invest. Ophthalmol. Vis. Sci., September 1, 2006; 47(9): 4077 - 4082. [Abstract] [Full Text] [PDF]
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G. L. Trick, P. Edwards, U. Desai, and B. A. Berkowitz Early supernormal retinal oxygenation response in patients with diabetes. Invest. Ophthalmol. Vis. Sci., April 1, 2006; 47(4): 1612 - 1619. [Abstract] [Full Text] [PDF]
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D.-Y. Yu and S. J. Cringle Oxygen distribution in the mouse retina. Invest. Ophthalmol. Vis. Sci., March 1, 2006; 47(3): 1109 - 1112. [Abstract] [Full Text] [PDF]
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H. Luan, R. Roberts, M. Sniegowski, D. J. Goebel, and B. A. Berkowitz Retinal Thickness and Subnormal Retinal Oxygenation Response in Experimental Diabetic Retinopathy Invest. Ophthalmol. Vis. Sci., January 1, 2006; 47(1): 320 - 328. [Abstract] [Full Text] [PDF]
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B. A. Berkowitz, R. Roberts, H. Luan, J. Peysakhov, D. L. Knoerzer, J. R. Connor, and T. C. Hohman Drug Intervention Can Correct Subnormal Retinal Oxygenation Response in Experimental Diabetic Retinopathy Invest. Ophthalmol. Vis. Sci., August 1, 2005; 46(8): 2954 - 2960. [Abstract] [Full Text] [PDF]
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