1 Department of Anatomy and Cell Biology, Wayne State University, Detroit, Michigan
2 Department of Ophthalmology, Wayne State University, Detroit, Michigan
3 Pharmacia Corporation, St. Louis, Missouri
4 Department of Medicine, Case Western Reserve University, Cleveland, Ohio
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ABSTRACT |
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Nitric oxide (NO), a potent regulator of retinal vascular function, is elevated in the vitreous humor of patients with proliferative diabetic retinopathy and with tractional retinal detachment, as well as in the retina of rodents, 24 months after the induction of diabetes (14). NO synthase (NOS) converts L-arginine to NO and L-citrulline. Excessive NO production by the inducible isoform of NOS (iNOS) in particular has been implicated in the pathogenesis of various ocular diseases (5,6). iNOS is induced in the retina in diabetes, but it is not yet known if iNOS regulates aspects of retinal circulatory pathophysiology associated with diabetes (4).
Previously, we 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 (710). In this technique, hyperoxia increases vitreous partial oxygen pressure over room-air values (PO2). Because oxygen is paramagnetic, this
PO2 will produce an increase in the vitreous signal intensity on a T1-weighted image. Furthermore, good agreement is found between the MRI-measured response and that determined by other investigators (11) using an oxygen electrode in normal rat retina under similar conditions. In normal adult and newborn rats, carbogen breathing oxygenated the retina significantly better than pure oxygen breathing (11). Carbogen is a gas mixture of carbon dioxide (5%) and oxygen (95%) that has been used clinically, instead of 100% oxygen, to minimize the vasoconstrictive effects of pure O2 on retinal blood flow and oxygenation.
The usefulness of the functional MRI approach has been tested in experimental diabetes models that reliably develop retinal histopathology (e.g., acellular capillaries and pericyte ghosts) after 15 months of hyperglycemia. In these models, 34 months after the induction of diabetes (before the appearance of retinal lesions), a subnormal retinal PO2 during carbogen breathing was observed (7,9). Aminoguanidine (AMG) treatment initiated with the induction of diabetes preserved the retinal oxygenation response at the 3-month time point and prevented the development of retinal histopathology at 15 months (9). Consistent with these results, 2 months of treatment with AMG, a relatively selective inhibitor of iNOS activity, also significantly inhibited nitrative stress in diabetic rat retina (4). In contrast, AMG treatment of galactosemic rats did not preserve the retinal oxygenation response at 3 months or prevent the development of retinal lesions at 15 months (7,9). Together these results suggest that correction of the early subnormal retinal
PO2 is a useful surrogate marker of treatment efficacy.
The present study tests the hypothesis that iNOS contributes to the early development of a subnormal PO2 in diabetic rodents. We compared the effect of both pharmacological and genetic iNOS inhibition on the retinal
PO2 in diabetic rat and mouse models, respectively.
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RESEARCH DESIGN AND METHODS |
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Animal model.
Two experimental models of diabetic retinopathy were studied: the diabetic rat and mouse. Each diabetic group started with 1520 animals. Animals were fed normal rat chow (5001; Ralston Purina, Richmond, IN) and water ad libitum. Rats were maintained in three groups for 3 months: controls (CR; n = 9), untreated diabetes (DR; n = 5), and diabetes treated with the prodrug L-N(6)-(1-iminoethyl)lysine 5-tetrazole amide (12) (100 ppm; D+PRO; n = 6). Prodrug was administered in the chow (i.e., 100 mg compound/kg of chow). The average daily dose depends on the amount of chow consumed (e.g., a 200-g rat consuming 20 g of chow per day will receive a dose of
10 mg/kg body wt).
Diabetes was induced in rats (starting weight 200220 g) with an intraperitoneal injection of streptozotocin (55 mg/kg, 0.01 mol/l citrate buffer, pH 4.5) after a 24-h fast and verified 24 h later by the presence of plasma hyperglycemia and glucosuria in nonfasted rats. Rat body weight, average food consumption, and blood glucose levels were monitored weekly. Subtherapeutic levels of insulin (04 units of neutral protamine Hagedorn [NPH] insulin administered subcutaneously up to 7 days per week, as needed) were administered to allow slow weight gain, yet maintain hyperglycemia and glucosuria. The diabetic animals produced in this study had mean plasma glucose levels of >400 but <550 mg/dl (Table 1).
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Measurement of NO.
In CR (n = 7 rats), DR (3 months, n = 6 rats), and D+PRO (3 months, n = 6 rats), both eyes were enucleated and retinas rapidly removed from urethane-anesthetized animals and frozen. As previously described, NO levels were indirectly determined from the concentration of stable metabolites of NO (nitrate plus nitrite) and measured using a fluorimetric assay kit (Cayman Chemical, Ann Arbor, MI) (4). Briefly, retinal homogenates were first filtered (10 kDa) and then incubated again with nitrate reductase for 2 h. Fluorescence generated by nitrite reaction with 2,3-diaminonaphthalene was measured. In all cases, samples were assayed simultaneously.
MRI exam.
On the day of the experiment, anesthesia was induced by a single intraperitoneal injection of urethane (36% solution, 0.083 ml/20 g body wt, prepared fresh daily; Aldrich, Milwaukee, WI). Each animal was gently positioned on an MRI-compatible homemade 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, pulse, and hemoglobin oxygen saturation (data not shown) were continuously monitored while the animal was inside the magnet, as previously described (9).
MRI data were acquired on a 4.7 T system using a two-turn transmit/receive surface coil (1.5-cm diameter) placed over the eye. Images were acquired using an adiabatic spin-echo imaging sequence (repetition time [TR] 1 s, echo time [TE] 22.7 ms [the shortest echo time allowed with this sequence], number of acquisitions [NA] 1, matrix size 128 x 256, slice thickness 1 mm, field of view 32 x 32 mm2 [for rats], 18 x 18 mm2 [for mice], sweep width 25,000 Hz, and 2 min/image) (14). This resulted in an in-plane resolution of 250 x 125 µm2 (rat) and 141 x 70 µm2 (mouse). 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. We reason that the retinal circulation is often required to maintain an adequate oxygenation level under different conditions (e.g., standing up versus sitting down), and the hyperoxic challenge acts as an acute "stress" test of the retinal circulations ability to oxygenate. Other stress tests (e.g., response to hypoxia and/or reduced blood pressure) can also be envisioned. However, relatively smaller signal intensity changes are expected from these challenges compared with that found during carbogen breathing. This would likely make analysis of the data difficult to detect/interpret. In the present study, we chose to use a carbogen challenge because large signal intensity changes are produced in both rat (11) and mouse (data not shown), allowing for the detection of subtler pathophysiology.
The MRI data were collected as follows. Sagittal localizer images were first collected and used to position a single 1-mm 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-min image. This slice thickness resulted in some partial volume averaging so that the final image contained superior and inferior hemiretina with some relatively minor contribution from temporal and nasal hemiretina. A capillary tube (1.5-mm inner diameter) filled with distilled water was used as the external standard (not shown in Fig. 1). It is important to note that steady state (room air) vitreous oxygen tension cannot be measured using 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.
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Data analysis.
To be included in this study, the animal must have demonstrated the following traits. 1) There had to be minimal eye movement during the MRI exam. Movement artifacts (typically seen in the phase-encode direction) will confound interpretation of the vitreous signal intensity changes produced during the hyperoxic challenge. 2) The animal had to have a nongasping respiratory pattern before and after the MRI exam. If the animal is gasping, which occurred <1% of the time, the anesthetic was likely improperly administrated (e.g., not intraperitoneally). This could produce a change in systemic oxygenation unrelated to the retinal changes. 3) Rectal temperatures in the range 35.536.5°C had to be observed. Preliminary experiments (data not shown) found a strong association between core temperature and PaCO2 and PaO2 levels. The effect of this correlation on the precision of the measurements was minimized by using a relatively tight range of temperatures. 4) Finally, PaO2 >350 mmHg and PaCO2 between 46 and 65 mmHg during the carbogen challenge had to be observed. Previously, we found (15) that arterial oxygen levels >350 mmHg during a hyperoxic challenge were needed to produce a consistently large preretinal vitreous oxygenation response. The range of acceptable arterial carbon dioxide levels lie within the array of values in the literature measured under carbogen-breathing conditions. In addition, tight control over the acceptable blood gas value range is needed to ensure adequate quality control of each sample. 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. In general, PO2 data were collected
60 min (rats) or 90 min (mice) after urethane injection to avoid potential errors due to variable time under anesthesia. The above acceptance criteria are needed to critically compare the retinal oxygenation response in these spontaneously breathing normal and sick animals while minimizing systemic differences. Because such tight criteria are used, only
50% of the animals that started the study were used in the final analysis. Based on our previous experience in rats, n
5 is sufficient to draw statistical conclusions.
To correct for any movement in the slice plane, a warp affine image coregistration was performed on each animal using software written in-house. This was done for all of the animals used in the final analysis but was only necessary in about half of them regardless of group (e.g., subtle shifting of the animals position occurred during the experiment due to settling on the gauze packing). Because the slice thickness (1 mm) is relatively large compared with the diameter of the eye (3 mm), partial volumes will be similar if the eye subtly moves out of the imaging plane, and so the data analysis results were not expected to be substantially affected. After coregistration, the MRI data were transferred to a Power Mac G4 computer and analyzed using the program NIH IMAGE (a freeware program available at http://rsb.info.nih.gov/nih-image). 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-min 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 (11). For each pixel, the fractional signal enhancement E was calculated as E = [S(t) - S0]/S0, where S(t) is the pixel signal intensity at time (t) after starting the gas inhalation and S0 is the control signal intensity (measured from the average of the three control images) at the same pixel spatial location. E values were converted into
PO2 using a theory that has been validated in the rat (16):
PO2 = E/(R1 x Tk), where R1 is the oxygen relaxivity (s-1 mmHg-1), and Tk = Tr x exp(-Tr/T10), where Tr is the repetition time and T10 is the T1 in the absence of oxygen. Using a Tr of 1 s, and assuming a vitreous T10 of 4 s, Tk = 3.52. This T10 value is based on our previous measurement of the proton spin-lattice relaxation time in the rabbit vitreous (4 s), reported values in human vitreous (3.3 s), and in cerebral spinal fluid (4.3 s), which has a similar high water content as vitreous (11,1719). An R1 of 2 x 10-4 s-1 mmHg-1 was used. This R1 was previously measured in a saline phantom, which is assumed to be a reasonable model of vitreous (98% water) (11). A similar R1 value was found for plasma, suggesting that relatively low protein levels do not substantially contribute to oxygen relaxivity (20). Note that an E of 0.01 (i.e., a 1% signal intensity change) corresponds to a
PO2 of 14 mmHg. There did not appear to be any significant changes in vitreous T10 or R1 in the animals of this study, based on the similar inferior hemiretinal oxygenation response found in all of the groups in this study.
The PO2 parameter image was analyzed as follows. First, the pixel values along a 1-pixel thick line drawn at the boundary of the retina/choroid and vitreous were set to 255 (black). We estimated that the thickness of this line, based on the in-plane resolution (e.g., in the mouse, an 18 x 18 mm2 field of view was sampled by 128 x 256 datapoints) of 140 x 70 µm2, is
100 µm. The values in another 1-pixel thick line drawn in the preretinal vitreous next to the black pixels were then extracted (21). This procedure minimized retinal/choroid pixel values from potentially contaminating ("pixel bleed") those used in the final analysis and insured that similar preretinal vitreous space was sampled for each animal. In addition, spatial averaging over these 100-µm regions of interest will tend to minimize the contribution from the very local preretinal oxygenation gradients next to the retinal surface (22). In addition, an average
PO2 band was constructed based on the within-group mean for each pixel.
Statistical analysis.
The physiological parameters (i.e., blood gas values, rectal temperatures, and blood glucose data) were normally distributed and are presented as mean ± SE. Comparisons were performed using an unpaired t test. Comparison of retinal PO2 between control and experimental groups was performed using a generalized estimating equation approach. This method performs a general linear regression analysis using all of the pixels in each subject and accounts for the within-subject correlation between adjacent pixels. In all cases, a P
0.05 was considered significant.
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RESULTS |
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Drug treatment.
Plasma prodrug levels at midstudy (1.5 month) and at the end of study (3 months) were 15 µmol/l, which is above the IC50 for iNOS (
5 µmol/l) and below the IC50 for endothelial NOS (>100 µmol/l) and neuronal NOS (50 µmol/l). The retinal NO level was evaluated using nitrite and nitrate, the stable metabolites of NO. Compared with CR, the retinal concentration of nitrite plus nitrate in DR was elevated
1.5-fold (5.3 ± 0.6 vs. 8.1 ± 1.2 pmol/mg protein, P < 0.05, respectively), and in D+PRO animals (2.7 ± 0.28 pmol/mg protein, P < 0.05), the concentration was significantly less than that of DR. These data are consistent with selective inhibition of iNOS in the D+PRO group.
Functional MRI.
MRI provides a clear image of rodent ocular anatomy in vivo, including the retina/choroid complex (seen as a white line in the posterior region of the eye) and the nucleus and cortex of the lens (Fig. 1) (7).
No significant (P > 0.05) differences were found in average inferior hemiretinal PO2 comparing any of the rat or mouse groups with their respective normal, nondiabetic controls (CR 169 ± 6 mmHg, DR 121 ± 6, D+PRO 214 ± 9, CM 129 ± 4, DM 142 ± 6, KO 171 ± 6, and D+KO 147 ± 9; mean ± SE). The mean superior hemiretinal
PO2 of the iNOS knockout mice was not significantly (P > 0.05) different from that of controls (Fig. 2). As expected, the average superior hemiretinal
PO2 of the 3-month diabetic rats and 4-month diabetic mice were significantly (P < 0.05) subnormal (9) (Fig. 2). In treated diabetic rats and in the diabetic iNOS knockout mice, the mean superior hemiretinal
PO2 values were not different (P > 0.05) from that of their respective controls (Figs. 2 4). To examine the regional effect of iNOS inhibition in diabetic rats and mice, we compared plots of the average superior hemiretinal pixel
PO2 values versus pixel position (i.e., distance) from the optic nerve head. As seen in Figs. 3 and 4, there is clear evidence that diabetic iNOS-inhibited rodents experienced a pansuperior retinal correction in
PO2.
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DISCUSSION |
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The subnormal superior hemiretinal PO2 found in 3-month diabetic rats and 4-month diabetic mice appears to be an early event in the sequence leading to histopathology in diabetes. Furthermore, treatments that prevented retinal lesion formation in long-term diabetic and galactosemic rats also prevented the development of a subnormal superior hemiretinal
PO2 in more acute studies (9). These observations strongly suggest that subnormal superior hemiretinal
PO2 is an early functional lesion that predicts the development of the structural lesions. The vascular lesions of diabetic retinopathy in rats or mice take longer to develop than the 3- to 4-month duration used in the present study (7,23,24), so we cannot currently evaluate the relation of NO to the development of histopathology. It was beyond the scope of this study to determine why the superior hemiretinal
PO2 becomes subnormal and the inferior hemiretinal does not. Nonetheless, based on the above, it appears that subnormal retinal
PO2 occurs before the appearance of experimental diabetic retinopathy.
Previous studies have shown that in diabetic rats, AMG, which is a relatively selective inhibitor of iNOS activity (among other actions), significantly inhibited diabetes-induced nitrative stress by 2 months, corrected the development of a subnormal superior hemiretinal PO2 by 3 months, and prevented the development of significant retinal histopathology by 15 months (4,9,25). However, based on these data, the association between diabetes-induced iNOS and the development of a subnormal
PO2 is indirect. The major finding in this study is that the inhibition of iNOS activity, either pharmacologically or genetically, prevented development of a subnormal superior hemiretinal
PO2 in diabetic rodents. One possible implication of the present study is that diabetes induces a greater increase in iNOS activity (and presumably NO level) in the superior hemiretina than in the inferior hemiretina. More work is needed to test this hypothesis.
The exact mechanism(s) by which iNOS produces a subnormal PO2 remains unclear. In diabetic rats and mice, chronic hyperglycemia produces an elevated retinal iNOS expression resulting in increased NO levels (3,4). Such nitrative stress may produce chronic vasodilatation that, coupled with other complications of diabetes, such as increased advanced glycated end products, could prevent the retinal vessel endothelium from responding appropriately to a carbogen challenge. Experiments are ongoing in the laboratory to further evaluate this speculation. It remains to be determined whether diabetic rodents treated with a specific inhibitor of iNOS activity will have subsequent retinal histopathology. Nonetheless, the present data establish that diabetes-induced increased iNOS activity as a key step in the development of early diabetic pathophysiology.
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ACKNOWLEDGMENTS |
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We are grateful to Dr. Tom Hohman for his careful reading of this manuscript.
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FOOTNOTES |
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Address correspondence and reprint requests to Bruce A. Berkowitz, PhD, Department of Anatomy and Cell Biology, Wayne State University School of Medicine, 540 E. Canfield, Detroit, MI 48201. E-mail: baberko{at}med.wayne.edu
Received for publication October 21, 2002 and accepted in revised form October 3, 2003
AMG, aminoguanidine; iNOS, inducible nitric oxide synthase; MRI, magnetic resonance imaging; NO, nitric oxide; NOS, NO synthase
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REFERENCES |
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