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Ionizing irradiation results in significant alterations in hippocampal neurogenesis that are associated with cognitive impairments. Such effects are influenced, in part, by alterations in the microenvironment within which the neurogenic cells exist. One important factor that may affect neurogenesis is oxidative stress, and this study was done to determine if and how the extracellular isoform of superoxide dismutase (SOD3, EC-SOD) mediated radiation-induced alterations in neurogenic cells. Wild type (WT) and EC-SOD knock out (KO) mice were irradiated with 5 Gy and acute (8–48 hr) cellular changes and long-term changes in neurogenesis were quantified. Acute radiation responses were not different between genotypes suggesting that the absence of EC-SOD did not influence mechanisms responsible for acute cell death after irradiation. On the other hand, the extent of neurogenesis was decreased by 39% in non-irradiated KO mice relative to WT controls. In contrast, while neurogenesis was decreased by nearly 85% in WT mice after irradiation, virtually no reduction in neurogenesis was observed in KO mice. These findings show that after irradiation, an environment lacking EC-SOD is much more permissive in the context of hippocampal neurogenesis. This finding may have a major impact in developing strategies to reduce cognitive impairment after cranial irradiation.
Radiation-related brain injury has a variable character, involving multiple regions and cell and tissue types with a large number of physical and biological factors influencing its extent [1, 2]. While overt tissue injury generally occurs after relatively high doses [3, 4], less severe morphologic changes can occur after lower doses, resulting in differing degrees of cognitive impairment. This latter change often involves deficits in hippocampal-dependent functions of learning and memory, including spatial information processing [5–9]. The underlying mechanisms responsible for radiation-induced cognitive impairment have remained elusive, although recently it has been suggested that changes in neuronal precursor cells and altered neurogenesis in the dentate subgranular zone (SGZ) of the hippocampus may be involved [10–14].
While radiation induced changes in neurogenesis involve a persistent decrease in precursor cells, there are data also showing that the ability of surviving cells to differentiate into mature cell phenotypes may involve an altered neurogenic environment [10, 13, 15–18]. One of the microenvironmental factors that may have a significant effect on neurogenesis is oxidative stress, a biochemical mechanism that has been shown to regulate the fate of neural precursor cells . The central nervous system (CNS) is inherently susceptible to oxidative injury, and because there are relatively low levels of endogenous antioxidants in the CNS [20, 21] that sensitivity has been implicated as playing a causative or contributory role in a number of neurodegenerative conditions, aging, and in ischemic, traumatic and excitotoxic damage [22–26]. The cellular changes observed in the CNS after irradiation are very similar to changes observed after other types of injury , and may involve increased production of reactive oxygen species (ROS), which can contribute to the spread and ultimate expression of tissue injury . Recent studies show that the radiation response of hippocampal neural precursor cells in culture is characterized by early apoptosis, specific cell cycle blocks, and activation of functional cell cycle checkpoints; these changes are coupled with elevated levels of ROS that persist for weeks after irradiation . Indications of persistent oxidative stress after irradiation have also been demonstrated in the brains of mice  and rats . Taken together, these data suggest that ROS may constitute critical environmental cues to control precursor cell survival and differentiation, and may play an important role in altered neurogenesis and perhaps cognitive impairment after irradiation .
There are several pathways that mitigate the physiological and pathological effects of ROS in mammalian cells . One of the pathways involves the antioxidant enzyme, superoxide dismutase (SOD), which exists as three genetically and geographically distinct isoenzymes . The SODs convert superoxide anions to hydrogen peroxide which is then enzymatically removed by catalase and glutathione peroxidase. Although the physiological roles of various isoforms of SOD in mammals are not completely understood, the extracellular isoform (EC-SOD, SOD3), was recently shown to be associated with certain cognitive functions [33, 34], and its removal was shown to interfere with signaling cascades critical for learning . Given the reported relationship between the effects of irradiation on SGZ precursor cells and changes in redox status within the hippocampal formation, along with recent evidence for a significant role of EC-SOD in learning and memory, we were interested in determining if there was a relationship between EC-SOD expression and hippocampal neurogenesis. Understanding how EC-SOD expression affects radiation-induced changes in neurogenesis may provide an important base for developing strategies to reduce cognitive impairment after cranial irradiation.
Congenic EC-SOD KO mice  on the C57BL/6J (B6) background were initially obtained from Dr. James Crapo at the National Jewish Medical and Research Center, Denver, Colorado. The colony is maintained by backcrossing to B6 mice purchased from the Jackson Laboratory (Bar Harbor, ME). Homozygous KO and their wild type littermate controls were generated from the intercrosses of heterozygous KO, and two month old male mice (WT, n = 50; KO, n = 52) were used in all experiments. All animal handling procedures were done according to institutional IACUCs. Mice were kept in a temperature and light-controlled environment with a 12 hr light/dark cycle, and provided food and water ad libitum.
For irradiation, animals were anesthetized with a mixture of ketamine hydrochloride (60 mg/kg, Abbott Laboratories, North Chicago, IL) and medetomidine hydrochloride (0.25 mg/kg, Orion Corp., Espoo, Finland), administered by intraperitoneal (i.p.) injection. Irradiations were done using a Phillips orthovoltage X-ray system as previously described . Briefly, animals were exposed to a single dose of 5 Gy using a special positioning jig so 4 animals could be irradiated simultaneously; the heads were centered in a 5 × 6 cm treatment field. The beam was directed down onto the head and the body was shielded with lead. Dosimetry was done using a Keithley electrometer ionization chamber calibrated using lithium fluoride thermal luminescent dosimeters. The corrected dose rate was approximately 175 cGy/min at a source to skin distance of 21 cm. Mice were irradiated at 8 weeks of age and were sacrificed, along with age-matched non-irradiated controls, at different times for various studies described below.
To determine if the major antioxidant enzymes were altered in EC-SOD KO mice, cortices and hippocampi were collected from irradiated and non-irradiated WT and KO mice one month after irradiation, which is the time when the neurogenesis study was carried out. For tissue collection, mice were anesthetized with i.p. injection of a mixture of ketamine hydrochloride (120 mg/kg) and medetomidine hydrochloride (0.5 mg/kg) and decapitated. Brains were removed and dissected on ice, and the hippocampus and cortex were isolated, frozen in dry ice and stored at −80°C. Cortex and hippocampus were homogenized separately in four volumes (weight to volume ratio 1:4) of PBS, pH 7.4, containing Complete® protease inhibitor cocktail (Roche, Switzerland), followed by 3 short pulses of sonication (3×5 sec). To ensure complete disruption of mitochondrial membranes, 3 rounds of freeze-thaw cycles between liquid nitrogen and room-temperature water were carried out. The samples were centrifuged at 20,800 g at 4°C for 5 minutes, and the supernatants were stored in 20 μl aliquots at −80°C. Protein concentration of each sample was measured in triplicates using the BCA Protein Assay Reagent (Pierce, Rockford, IL).
CuZnSOD (SOD1) and MnSOD (SOD2) activities were determined by Ampholine PAGplate (Amersham Pharmacia Biotech, Inc. Piscataway, NJ), pH 3.5 – 9.5, as described previously . For the CuZnSOD assay, 2.5, 5.0 and 10 μg of each sample were analyzed on the gel; for the MnSOD assay, 15, 30 and 60 μg of each sample were analyzed. The activity stain creates clear bands on a dark purple background and identifies the location of CuZnSOD and MnSOD in the gel. The band intensity, which is proportional to the enzyme activity, was quantified by Image J 1.36b (NIH, USA) and normalized to total protein loading.
The protein levels of catalase, glutathione peroxidase 1 (GPx-1), CuZnSOD, and MnSOD were determined by western blot analysis. Equal amounts of protein (cortex, 30 μg; hippocampus, 50 μg) from each sample were separated by NuPAGE 4–12% Bis-Tris gel (Invitrogen) and transferred to nitrocellulose membranes (Bio-Rad, Hercules, CA). The following primary antibodies were used for the detection of specific proteins: catalase (C 0979, 1:4,000, Sigma, St. Louis, MO), GPx-1 (LF-PA0019, 1:1,000, LabFrontier Co. Korea), CuZnSOD (LF-PA 0013, 1:2,000, LabFrontier), MnSOD (SOD-110, 1:4,000, Stressgen, Ann Arbor, MI). Protein bands were visualized using DuoLuX Chemiluminescent/Fluorescent Substrate Kit (Lumigen, Southfield, MI) after incubation with HRP-conjugated secondary antibody. All blots were stripped with stripping buffer (Tris-HCl, 62.5 mM; β-mercaptoethanol, 90 mM; 1% SDS) and reprobed with an antibody against β-actin (A 3854, 1:50,000, Sigma) as a loading control. Quantification of western blot results was done by normalizing the signal intensity of each sample to that of β-actin.
Assuming that KO mice were under persistent oxidative stress, we made a qualitative appraisal of 4-hydroxynonenal (4HNE) and nitrotyrosine (NT) immunoreactivities using randomly selected sections of the hippocampus. The sections were immunostained as described previously  using anti-4HNE primary antibody (1:100, JaICA, Fukuroi, Shizuoka, Japan) or anti-NT primary antibody (1:100, Zymed, South San Francisco, CA) with subsequent visualization with anti-mouse secondary antibody conjugated with FITC fluorophore (1:200, Jackson ImmunoResearch Labs, West Grove, PA). Nuclei were counterstained with To-Pro 3 (Molecular Probes, Eugene, OR).
To determine if there was a general difference in the pattern of acute cell death (apoptosis) after irradiation of WT and KO mice, tissues were collected from 2–4 animals of each genotype at 0, 8, 12, 24 and 48 hr after irradiation. For tissue collection, animals were anesthetized as described above and perfused with 50 ml of 10% neutral buffered formalin into the ascending aorta using a mechanical pump (Masterflex Model 7014; Cole Parmer, Chicago, IL) over a 5 min period; brains were removed and immersed in 10% neutral buffered formalin for 3 days. Brains were then placed in a Mouse Brain Matrix (Harvard Apparatus, Natick, MA) and a 5 mm-thick transverse section containing the hippocampus was taken for paraffin embedding. Five, 6 μm-thick sections were cut from the rostral face of each block using a rotary microtome, starting at a point 2.5 mm behind the bregma  and were placed onto polylysine coated glass microscopic slides. A second set of 5 sections were obtained 100 μm caudal to the first set of sections, and a third set was taken after another 100 μm. Thus, sections were taken from 3 distinct areas representing the rostral-mid dentate gyrus [11, 13].
In the acute study we were only interested in determining relative changes in numbers of cells dying within the dentate SGZ as a function of time after irradiation. To do this we used TUNEL (terminal deoxynucleotidyl transferase-mediated dUTP-biotin nick end labeling) staining with a commercial kit (Apotag, Serological Corp., Norcross, GA.) as described before [11, 38]. Because not all apoptotic cells necessarily show TUNEL staining, we also used morphologic criteria to estimate apoptosis . The morphological changes used included fragmentation, or the compaction of chromatin into 2 or more dense, lobulated masses, and pyknosis, which was characterized by small, round, darkly staining nuclei. To minimize the impact of including any normal cell profiles in our counts of apoptosis, if any cytoplasm was observed in conjunction with a small, dense nucleus, that cell was not considered as apoptotic.
To determine the overall early impact of irradiation on specific cellular components of the SGZ we quantified proliferating cells and immature neurons 48 hr after irradiation, when apoptosis was complete . We also quantified these cells from tissues obtained 1 month after irradiation, at the time when our neurogenesis study was done (see below). Proliferating cells were labeled with an antibody against Ki-67 (DakoCytomation, Caprinteria, CA, diluted 1:100 in PBS with 2% normal rabbit serum), a nuclear antigen expressed during all stages of the cell cycle except Go [40, 41]. Immature neurons were labeled using an antibody against Doublecortin (DCx, Santa Cruz Biotechnology, Santa Cruz, CA; diluted 1:500 in PBS with 5% normal horse serum), a tubulin-associated protein expressed in migrating neuroblasts [42–46] as described previously . Briefly, sections were incubated overnight at 4°C with primary antibodies and subsequently for 30 min at room temperature with biotinylated secondary antibodies (Vector Lab, Burlingame, CA; rabbit anti-rat IgG, diluted 1:200 in PBS with 2% normal rabbit serum for Ki67 and anti-goat IgG, diluted 1:500 in 5% normal horse serum for DCx). Binding of secondary antibodies was detected using an avidin-biotinylated peroxidase complex system (ABC; Vector Lab) and developed with 0.025% 3,3′-diaminobenzidine (DAB, Zymed) dissolved in double distilled water containing 0.005% H2O2. Sections were then counterstained with Gill’s hematoxylin, dehydrated and mounted.
Cell numbers positive for TUNEL/morphologic changes, Ki-67, and DCx were scored using a histomorphometric approach with the genotype and treatment blinded to the researcher. The methods used are well standardized and have been used effectively to determine responses of cells in the dentate SGZ in both mice and rats after radiation and other lesions [11, 47–49]. For the analysis of cell death, we counted all apoptotic cells in the suprapyramidal and infrapyramidal blades of the dentate SGZ and granular cell layer (GCL). For the analyses of Ki-67 and DCx, we focused our counting on all positively labeled cells within the SGZ, defining it as the area immediately adjacent to the hilus and extending roughly 3 cells deep into the dentate granule cell layer. For each animal, 3 slides were counted, one from each of the 3 regions of the dentate. For each marker and each mouse, the total number of positively labeled cells was determined by summing the values of all 3 sections; this approach of counting cells in 3 evenly-spaced, non-contiguous tissue sections is referred to as our standardized counting area.
To determine the effects of irradiation on the survival and fate of newly generated cells in the SGZ as a function of genotype, groups of WT (n = 4) and KO mice (n = 4) received a single i.p. injection (50 mg/kg) of 5-bromo-2′ deoxyuridine (BrdU, Sigma, St. Louis, MO) daily for 7 days starting at 30 days after irradiation. Three weeks after the last BrdU injection, mice were anesthetized as described and perfused with ice-cold saline followed by freshly prepared, ice-cold 4% paraformaldehyde. The brain was removed, processed, and sectioned using a sliding microtome . Fifty micrometer sections were stored at 4°C in cryoprotectant solution until needed. Free floating sections were immunostained as described [11, 13] using the following primary antibodies and working concentrations: rat anti-BrdU (1:10; Oxford Biotechnology, Kidlington, Oxford, UK); mouse anti-NeuN (1:200; Chemicon, Temecula, CA); rabbit anti-NG2 (1:200; Chemicon); goat anti-GFAP (1:100, Santa Cruz Biotechnology); rat anti-CD68 (1:20; Serotec, Inc. Raleigh, NC).
To calculate the numbers of BrdU-positive (BrdU+) cells in the dentate gyrus, at least 12 sections of a one-in-six series were scored per animal. All counts were limited to the dentate granule cell layer and a 50 μm border along the hilar margin that included the dentate SGZ. Total numbers were obtained by multiplying the measured value by 6; overestimation was corrected using the Abercrombie method . Total numbers of BrdU+ cells displaying the various lineage-specific phenotypes were determined using confocal microscopy to score the colocalization of BrdU and phenotypic markers in representative sections from each animal [11, 13]. Confocal microscopy was performed using a Nikon C-1 confocal microscope (Melville, New York), using techniques previously described [10, 11, 13]. Appropriate gain and black-level settings were obtained on control tissues stained with secondary antibodies alone. Upper and lower thresholds were always set using a range indicator function to minimize data loss due to saturation. Each cell was manually examined in its full ‘z’ dimension with use of split panel analysis, and only those cells for which the BrdU+ nucleus was unambiguously associated with the lineage-specific marker were scored as positive. For each lineage-specific marker, the percentage of BrdU+ cells expressing that marker was determined. Total numbers of lineage-specific BrdU+ cells were then calculated by multiplying this percentage by the total number of BrdU+ cells in the dentate gyrus.
The impact of irradiation on total numbers of activated microglia in the SGZ was determined using free floating brain sections from groups of WT (n = 4) and KO mice (n = 4). After 3 washes with TBS and quenching in TBS with 3% H2O2 for 15 min, sections were incubated in TSA blocking buffer (PerkinElmer Life Sciences, Emeryville, CA) for 30 min, followed by application of the polyclonal rabbit anti-CD68 antibody (1:20; Serotec, Inc. Raleigh, NC). After 12hr at 4ºC, sections were incubated for 1 hr at room temperature with a secondary anti-rabbit biotinylated antibody (Vector, Burlingame, CA), followed by incubation with an Avidin+Biotin amplification system (Vector) for 45 minutes. Cell staining was visualized using the TSA fluorescence system CY3 (PerkinElmer Life Sciences, Emeryville, CA); nuclear counterstaining was with Sytox-Green (Molecular Probes, Eugene, OR). No staining was detected in the absence of the primary or secondary antibodies.
Total numbers of activated microglia were counted in each of 4 coronal sections of the dentate gyrus and hilar region at the level of the dorsal hippocampus (~−3.6 mm posterior to the Bregma). Images were reconstructed as described previously  and shown in Fig. 5. The mosaics were collected with a Zeiss AxioImager Apotome microscope using a 20X objective. The parameters were kept constant across sections. Regions of interest containing the dentate gyrus and hilar region was selected using AxioImager imaging software (Zeiss) and the numbers of positive cells were counted within the selected area. To avoid classification errors, we carefully verified that the staining belonged to the cell of interest and not to a dendrite or the cell body of an adjacent cell. The final numbers of activated microglia were expressed as number of cells/per mm2.
For each endpoint, values for all animals of a given treatment group were averaged and standard errors of mean (SEM) were calculated. Student’s t test was used to compare the results from western blotting and IEF gel analyses. For the studies of cell numbers (when applicable), a one-way ANOVA with post-hoc Duncan “D” or Fisher tests with Bonferroni correction were used to determine if there were significant differences between groups as a function of radiation treatment and genotype. To directly compare the 2 genotypes (EC-SOD KO and WT), if applicable, a two-way ANOVA was used, with treatment, genotype and treatment-genotype interaction as factors.
No differences in catalase, GPx-1, MnSOD, and CuZnSOD protein levels were detected between WT and KO mice with or without irradiation (Fig. 1A and 1B). There was also no difference in CuZnSOD and MnSOD activities (Fig. 1C and 1D). This suggested that at least for the enzymes analyzed here, there were no apparent compensatory changes in the major antioxidant defense systems when EC-SOD was knocked out.
Immunostaining of WT and KO mouse tissues for 4HNE showed a qualitative increase in immunoreactivity in KO mice compared to their WT littermates (Fig. 2A and 2B); this was consistent with a higher baseline level of oxidative stress in the KO mice. Based strictly on morphology, the localization of 4HNE appeared to be vascular and, perhaps, microglial (Fig. 2B, inset). Given the relationship between excess superoxide and the formation of peroxynitrites , we also used immunohistochemistry to qualitatively assess nitrotyrosine, and again, there was increased immunoreactivity in non-irradiated KO mice (Fig. 2C and 2D).
In non-irradiated animals, the total number of TUNEL-positive nuclei in our standardized counting area ranged from about 15 to 20 in both WT and KO mice; these values are slightly lower than reported by us previously  and there was no significant difference between WT and KO mice. Dying cells were seen in the SGZ of both blades of the dentate and only occasionally in the hilus and GCL. After irradiation, however, the number of apoptotic nuclei increased by 15–20 times in both WT and KO at the peak of apoptosis (8 hr after irradiation; data not shown). There was no apparent difference in the pattern or magnitude of apoptosis between WT and KO mice, and in both genotypes, the numbers of TUNEL-positive cells returned to control levels by 24 hr after exposure (data not shown).
In our standardized counting area, the number of Ki-67-positive cells averaged 49.8 ± 5.4 (n = 5) in non-irradiated WT mice and 49.5 ± 14.9 (n = 4) in non-irradiated KO mice (Fig. 3A). The numbers of DCx-positive immature neurons averaged 298.0 ± 19.2 (n = 5) and 321.7 ± 51.8 (n = 4) in non-irradiated WT and KO animals, respectively (Fig. 3B). Forty-eight hours after irradiation there were highly significant reductions in the numbers of proliferating cells (p=0.003 and p=0.002 for WT and KO, respectively) and immature neurons (p=0.001 and p=0.002 for WT and KO, respectively) in both genotypes (Fig. 3A and 3B).
While our acute studies showed that there were substantial decreases in the numbers of proliferating cells and their progeny shortly after irradiation (Fig. 3A and 3B), some precursor cells survived the x-ray treatment. To determine if the initial loss of cells translated into a longer-term impairment of neurogenic populations in the SGZ and if the condition of EC-SOD deficiency affected those populations, we assessed the numbers of proliferating cells and immature neurons 1 month after irradiation. At that time there were still apparent reductions in the numbers of proliferating cells and immature neurons after irradiation (Fig. 3C and 3D). However, technical problems resulted in the loss of some tissues and reduced the numbers of animals in some of the treatment groups; this compromised our ability to do complete statistical analyses. Given that caveat, the data in Fig. 3 suggest that while the radiation-induced loss of proliferating cells and immature neurons appeared to be persistent, there might have been some recovery in both genotypes 1 month after irradiation compared to that at 48 hr post irradiation.
BrdU+ cells observed 3 weeks after the last BrdU injection represent the long-term survival of newly generated cells (Fig. 4A, B). In the non-irradiated mice, the total number of BrdU+ cells in the dentate gyrus of WT mice averaged 4187 ± 614, while in KO mice the average value was 3197 ± 325; the difference was not significant (p = 0.2). In WT mice, an average of 2920 ± 301 newly generated cells differentiated into neurons (BrdU+/NeuN+; Fig. 4C, D). In non-irradiated KO mice, the number of newly generated neurons averaged 1793 ± 226 (Fig. 4C and D); compared to WT the difference was highly significant (p = 0.007). In non-irradiated WT mice there was an average of 164.4 ± 43.1 BrdU+ cells that co-expressed GFAP (newly generated astrocytes) while in non-irradiated KO mice the average number of GFAP-positive cells was 292.2 ± 98.3; this difference was not significant (p = 0.3; Fig. 4E and F). The numbers of newly generated oligodendrocytes (BrdU+/NG2+) were not different between WT and KO mice before irradiation (Fig. 4G and H).
Total numbers and numbers of newly generated (BrdU+/CD68) activated microglia were detected using the anti-CD68 antibody, and were present in the dentate SGZ of both non-irradiated WT and KO mice. In non-irradiated mice there were no differences in total numbers of activated microglia between WT and KO mice (Fig. 5) and while there appeared to be a difference in newly generated activated microglia (WT: 318.7±77.9, KO: 526.5±154.7), the values were not significantly different (p = 0.5). Finally, peripheral monocytes were detected by scoring BrdU+ cells that expressed both NG2 and CD68 [53, 54]. The average numbers of peripheral monocytes were relatively low, ranging from 14.1 ± 14.1 in WT to 34.1 ± 26.4 in KO mice; given inter-animal variability, the difference was not significant (p = 0.5).
In WT mice after a single dose of 5 Gy, the average number of BrdU+ cells was reduced by 76%, to 1003 ± 503 (p = 0.002; Fig. 4B), and this was translated into a 86% reduction in the BrdU+/NeuN+ population (p = 0.002; Fig. 4D). In contrast, the average number of BrdU+ cells seen after irradiation of KO mice increased from 3197 ± 325 to 4140 ± 595 (p = 0.2; Fig. 4B). Consequently, irradiation resulted in no significant reduction (p = 0.4) in the number of BrdU+/NeuN+ cells in EC-SOD KO mice (Fig. 4D). The differences in the total number of BrdU+ and BrdU+/NeuN+ cells between irradiated WT and KO mice were highly significant (p = 0.002 and p = 0.007, respectively). A 2-way ANOVA showed that for new neuron production there was a highly significant (p = 0.001) interaction between irradiation and genotype In terms of newly generated astrocytes, 5 Gy resulted in about a 12% reduction in the number of BrdU+/GFAP+ cells in the WT group and in an apparent increase in KO mice (Fig. 4F) although these changes were not significantly different from non-irradiated controls. However there was a significant difference in the number of BrdU+/GFAP+ cells between irradiated WT and irradiated KO mice (p=0.03). While there appeared to be slightly fewer BrdU+/NG2+ cells in irradiated WT animals (p = 0.7) (Fig. 4H), there was a trend toward increasing numbers in KO mice (p = 0.1). Consequently, there was a significant difference in the number of BrdU+/NG2+ cells between irradiated WT and irradiated KO mice (p=0.03).
After irradiation there were significant increases in the total numbers of activated microglia in both the WT (p = 0. 056) and KO mice (p = 0.01) when compared to non-irradiated controls (Fig. 5). Furthermore, there was a significant difference (P = 0.03) in the total numbers of activated microglia between irradiated WT and irradiated KO mice. In terms of newly generated activated microglia (BrdU+/CD68+), there were non-significant increases in both genotypes after irradiation; however there was a significant difference between irradiated WT and irradiated KO mice (p=0.03; not shown). Finally, there was a trend toward increasing numbers of BrdU+ peripheral monocytes in the SGZ of both WT and KO mice, but the differences were not significant compared to non-irradiated controls, and the numbers of peripheral monocytes were almost an order of magnitude lower than the numbers of BrdU+ activated microglia (data not shown).
The main findings of the present study were: 1) EC-SOD KO mice had a lower baseline hippocampal neurogenesis compared to WT mice; 2) proliferating precursor cells and immature neurons of the dentate SGZ in EC-SOD KO mice showed acute (6–48 hr) sensitivity to irradiation and the level was comparable to that of WT mice; 3) survival of newly generated cells in the dentate SGZ after 5 Gy irradiation was substantially higher in EC-SOD KO mice compared to WT controls; and 4) contrary to what was seen in WT mice, irradiation did not reduce the survival of newly generated neurons and glia in EC-SOD KO mice. These findings showed that after irradiation, an environment lacking EC-SOD was much more permissive in the context of hippocampal neurogenesis. The mechanism(s) behind this observation is not yet known, but this effect might constitute a basis for future interventions aimed at rescue or at least amelioration of the risks for cognitive dysfunction in individuals subjected to CNS irradiation.
The effects of irradiation on hippocampal structure and function have been extensively studied (reviewed in [5, 55]) but only recently has it been suggested that radiation-induced injury of the neurogenic cell populations within dentate gyrus [10, 11, 39, 48] may play a role in cognitive sequelae of cranial irradiation. Studies from our lab [10, 11, 13, 14, 48] and others [12, 56] provide a quantitative description of the impact that ionizing radiation exerts on hippocampal neurogenesis. Those studies show that the stem/precursor cell populations within the neurogenic areas are very vulnerable to injury and that changes are associated with radiation-related impairments of hippocampal-dependent cognitive tasks [12, 14, 57, 58]. Furthermore, radiation-induced changes in neurogenesis have been shown to be associated with microenvironmental factors including neuroinflammation [56, 59], vascular changes [10, 60] and oxidative stress [28, 61].
In this study we were interested in determining if and how the absence of EC-SOD would impact neurogenesis. EC-SOD immunoreactivity is found throughout the brain, and is particularly prominent in the hippocampus . It is of particular interest that alterations in EC-SOD expression in mice is associated with impaired learning , and that over-expression of EC-SOD protects synaptic plasticity and learning and memory against oxidative damage . Here we found that the absence of EC-SOD was not associated with compensatory changes in other SODs or other anti-oxidant enzymes (Fig. 1), although given the intracellular localization of these molecules it might be unlikely to expect them to have compensatory functions in terms of affecting extracellular superoxide free radicals. It could be possible that circulating antioxidants, including ascorbic acid, tocopherol, uric acid, bilirubin, proteins and other compounds, could be increased as a compensatory mechanism, but we did not address this in the current study. While total antioxidant capacity in the serum can be assessed quantitatively , without knowing the amount of circulating EC-SOD and its contribution to total antioxidant capacity in the serum, how these compounds may or could affect regional effects in the SGZ may be difficult to interpret. However, a consideration of such ideas is worthy of further study. Regardless, our data show that at least qualitatively, 2-month-old EC-SOD KO mice showed indications of a persistent oxidative stress (Fig. 2). As a result, and based on data relating EC-SOD with cognitive function [34, 63], we expected neurogenesis in the KO mice to be lower than that of WT mice. This was, in fact, what we observed (Fig. 4). Whether these changes are responsible for the cognitive impairment observed in EC-SOD KO mice is not clear. However, given the extensive data available associating changes in neurogenesis with cognitive performance [11, 13, 14, 65–68], it certainly seems possible, and at the very least may play a contributory role. The baseline reduction in new neuron production in KO mice was different from what was seen in glia (Fig. 4) and inflammatory cells (Fig. 5), where there were apparent increases in cell number in KO mice relative to WT. While the significance of these latter findings may be affected by the variability in the data, increased numbers of astrocytes and activated microglia would be consistent with an elevated level of oxidative stress in the KO animals.
After irradiation with a relatively low dose of x-rays, the early responses, including apoptosis, altered cell proliferation, and changes in numbers of immature neurons, were similar between WT and KO mice. This suggested that the absence of EC-SOD and the increased oxidative stress status of KO animals did not influence mechanisms responsible for acute cell death after irradiation. Furthermore, there were no major differences between WT and KO mice in cell proliferation and numbers of immature neurons at 1 month after irradiation (Fig. 3). It was of interest, therefore, to see substantial and significant differences in the survival of newly generated cells after irradiation as a function of genotype, with KO mice showing almost 4 times the number of surviving newly generated cells than WT (Fig. 4). This surprising result also translated into a very significant and nearly 4-fold difference in the number of surviving newly generated neurons in irradiated KO as compared to irradiated WT mice. In fact, while radiation caused an 85% reduction in newly generated neurons in WT mice, the same dose resulted in virtually no difference in KO mice. To the best of our knowledge, this is the first report to show that an EC-SOD deficient environment provides a ‘protective’ effect in hippocampal neurogenesis after irradiation. In a general sense, this resembles the preconditioning seen in stroke , heart attack  or acute lung injury  where oxidative mechanisms plays a crucial role in the pathogenesis of the disease. While the precise mechanism(s) responsible for this effect has not yet been clarified, our study rules out simple compensatory changes in expression and activities of other antioxidant enzymes (Fig. 1). Our findings clearly suggest that EC-SOD deficient mice have developed a resistance to radiation-induced inhibition of neurogenesis that may involve some type of adaptation within the microenvironment, without compensatory changes in other major intracellular antioxidants. These ideas are being tested further using inducible in vitro and in vivo models of EC-SOD expression. Furthermore, it is possible that a specific set of trophic factors and signaling molecules that favor differentiation and long-term survival of newly-generated neurons in the SGZ is up-regulated or activated in the irradiated EC-SOD KO brains to counter the chronic inflammatory environment created from irradiation. These trophic factors and signaling molecules may include brain derived trophic factor (BDNF), vascular endothelial growth factor (VEGF), and nitric oxide (NO), all of which have been shown to favor differentiation and survival of neurons [72–75]. At the same time, the phenotype of the activated microglia [76, 77] in the irradiated EC-SOD null environment may also play a role in enhanced neurogenesis. Whatever the mechanism(s) involved, our data clearly show that the production of new neurons is more sensitive to irradiation in WT animals than in the EC-SOD KO mice.
Due to the relatively lower numbers of newly generated astrocytes and oligodendrocytes, there is more variability in the data for these cells, but clearly there appears to be a trend toward increased numbers in KO vs. WT mice after 5 Gy of x-rays (Fig. 4). This suggests that perhaps the same mechanism responsible for the relative protective effect of neurons in EC-SOD KO mice is operative in glial cells as well. Given recent data showing that neural stem cells in the SGZ express GFAP , it is possible that the increased numbers of GFAP+ cells seen in KO mice represent, in part, increased stem cell or regenerative responses. On the other hand, the observed changes in BrdU+/GFAP+ cells may very well be a sign of the well-described astrogliosis seen in the oxidative stress conditions  as well as after irradiation .
Neurogenesis depends upon a complex microenvironment that involves signaling between multiple cell types , and changes in the oxidative status along with irradiation could affect any or all of these cells or their interactions. While the precise nature of such effects has not yet been clarified, previous studies have suggested chronic inflammatory changes as one of the important factors [10, 11, 13, 56]. We found significant increases in the total numbers of activated microglia in both WT and KO mice after irradiation as compared to non-irradiated mice (Fig. 5). Furthermore, there appeared to be increases in numbers of newly generated activated microglia (BrdU+/CD68+) in both genotypes although the magnitude of the differences was not statistically significant. That increasing numbers of activated microglia after irradiation had no apparent association with neurogenesis in EC-SOD KO mice is somewhat surprising, given recent studies showing that increased neuroinflammation was linked to an inhibition of hippocampal neurogenesis [13, 56, 59, 82]. It is particularly interesting, therefore, that a recent study has suggested that microglial phenotype critically influences their ability of these cells to support or impair cell renewal processes in the adult brain . It may be that in a microenvironment characterized by persistent oxidative stress (i.e., EC-SOD KO), the subsequent activation of microglia may, in fact have a beneficial effect, at least in terms of neurogenesis. This would suggest that microglia may respond differently to a given stimulus (irradiation) depending upon the presence of another and preceding stimulus (oxidative stress); such ideas have recently been reviewed . While intriguing, such a hypothesis is speculative at this time, but further studies seem warranted.
While the mechanistic relationship between EC-SOD and forebrain neurogenesis is not yet known, the data shown here clearly suggest that the lack of EC-SOD provides some sort of survival advantage to neurogenic cell populations after irradiation. This could have major implications with respect to protecting or ameliorating the adverse effects of radiation on specific brain functions if the underlying changes in the molecular network can be identified.
The authors want to thank Dr. Kathleen Lamborn of the Brain Tumor Research Center, Dept. Neurological Surgery, UCSF, for assistance in the statistical analyses. This work was supported in part by NIH grant R01 NS46051 (JRF) NASCOR grant NNJ04HC90G (JRF), NIH grant AG24400 (TTH) and ACS grant RSG-00-036-04-CNE (CLL)
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