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Keywords:

  • hypoxia-ischemia;
  • inflammation;
  • microglia;
  • neurogenesis;
  • radiotherapy;
  • stroke

Abstract

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. References

Cranial radiotherapy is common in pediatric oncology. Our purpose was to investigate if irradiation (IR) to the immature brain would increase the susceptibility to hypoxic-ischemic injury in adulthood. The left hemisphere of postnatal day 10 (P10) mice was irradiated with 8 Gy and subjected to hypoxia-ischemia (HI) on P60. Brain injury, neurogenesis and inflammation were evaluated 30 days after HI. IR alone caused significant hemispheric tissue loss, or lack of growth (2.8 ± 0.42 mm3, p < 0.001). Tissue loss after HI (18.2 ± 5.8 mm3, p < 0.05) was synergistically increased if preceded by IR (32.0 ± 3.5 mm3, p < 0.05). Infarct volume (5.1 ± 1.6 mm3) nearly doubled if HI was preceded by IR (9.8 ± 1.2 mm3, p < 0.05). Pathological scoring revealed that IR aggravated hippocampal, cortical and striatal, but not thalamic, injury. Hippocampal neurogenesis decreased > 50% after IR but was unchanged by HI alone. The number of newly formed microglia was three times higher after IR + HI than after HI alone. In summary, IR to the immature brain produced long-lasting changes, including decreased hippocampal neurogenesis, subsequently rendering the adult brain more susceptible to HI, resulting in larger infarcts, increased hemispheric tissue loss and more inflammation than in non-irradiated brains.

Abbreviations used:
DG

dentate gyrus

GCL

granule cell layer

HI

hypoxia-ischemia

IR

irradiation

MAP-2

microtubule-associated protein-2

P10

postnatal day 10

P-HH3

phospho-histone H3

RT

radiation therapy

SGZ

subgranular zone

SVZ

subventricular zone

Radiotherapy (RT) is one of the most effective tools in the treatment of malignant tumors, and is applied not only to adult patients but also to children who suffer from primary or metastatic brain tumors and central nervous system involvement of leukemia or lymphoma. Irradiation (IR) to the whole body, including the brain, is included in some protocols prior to hematopoietic stem cell transplantation. Damage to normal, surrounding brain tissue constitutes a major problem and radiotherapy is associated with adverse side effects, particularly in pediatric patients (Mulhern and Palmer 2003). Intellectual impairment as well as perturbed growth and puberty are some of the side effects seen after RT (Chin and Maruyama 1984; Packer et al. 1987; Lannering et al. 1990, 1995; Spiegler et al. 2004).

Ionizing radiation can produce free radicals and DNA damage, causing proliferative cells to undergo apoptosis or mitotic catastrophe. Mature neurons are considered to be in a permanent state of growth arrest, whereas stem and progenitor cells have a prominent proliferative capacity and are therefore highly vulnerable to irradiation. Neurogenic areas, the subventricular zone (SVZ) and the subgranular zone (SGZ) of the dentate gyrus (DG) in the hippocampus, are highly susceptible to IR-induced injury. This has been demonstrated in rodents (Tada et al. 2000; Fukuda et al. 2004; Raber et al. 2004b; Fukuda et al. 2005) and appears to be true also for humans (Monje et al. 2007). In addition to acute cell loss, IR can influence the survival of stem and progenitor cells, leading to a limited potential in terms of repopulation or regeneration (Tada et al. 2000). It has been shown that IR induces vascular abnormalities, demyelination and alterations in the microenvironment of the brain, shifting the proliferative response of progenitors from neurogenesis to gliogenesis (Monje et al. 2002). One study showed that a single dose of 1–15 Gy to the heads of adult mice or rats permanently abolished adult neurogenesis in the DG without incapacitating the animals (Monje et al. 2003). In contrast, a recent study using a single dose of 4 Gy to young adult mice found a reversible effect on proliferation and neurogenesis in the DG (Ben Abdallah et al. 2007). To the best of our knowledge, there are no studies evaluating the long-lasting effects of irradiation to the immature brain on a subsequent ischemic insult to the adult brain.

Recovery after brain injury depends on the balance between mechanisms of injury and protection (Walton et al. 1999). Accumulating evidence demonstrates that ischemic brain injury increases neurogenesis in rodent brains (Liu et al. 1998; Arvidsson et al. 2002; Qiu et al. 2007). Newborn neurons can become functionally integrated into the dentate gyrus (van Praag et al. 2002). Importantly, newly generated neurons may play a significant role in synaptic plasticity, and a reduction in the number of these cells or inhibition of neurogenesis impairs spatial learning and causes cognitive impairment (Shors et al. 2002; Raber et al. 2004a,b; Rola et al. 2004). It remains to be demonstrated conclusively if ischemia-induced neurogenesis contributes to recovery after an insult. Conversely, it is not known if decreased neurogenesis, as for example after radiotherapy, renders the brain more susceptible to hypoxic-ischemic (HI) injury.

In this study, we used a moderate single dose of ionizing radiation (8 Gy) to induce a persistent reduction of neurogenesis in the left hemisphere of young mice (Naylor et al. 2008). Fifty days later, when the mice were adult, we induced a HI insult in the same hemisphere and evaluated neurogenesis, inflammation and brain injury 30 days later.

Materials and methods

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. References

Irradiation procedure

A linear accelerator (Varian Clinac 600CD, Radiation Oncology Systems, San Diego, CA, USA) with 4 MV nominal photon energy and a dose rate of 2.3 Gy/min was used as described previously (Fukuda et al. 2004, 2005; Naylor et al. 2005). Postnatal day 10 (P10) male C57/BL6J mouse pups (Charles River, Germany) were anesthetized with an intraperitoneal (i. p.) injection of 50 mg/kg tribromoethanol (Sigma, Stockholm, Sweden). The animals were placed in prone position (head to gantry) on an expanded polystyrene bed. The left cerebral hemisphere was irradiated with a radiation field of 1 × 2 cm. The IR source to skin distance was about 99.5 cm. The head was covered with a 1 cm tissue equivalent material to obtain an even IR dose throughout the underlying tissue. The dose variation within the target volume was estimated to be ±5%. The contralateral hemisphere did not receive any direct radiation, but indirect, secondary radiation has been shown to cause some injury also in the non-irradiated hemisphere (Fukuda et al. 2004). The entire procedure was completed within 10 min and no pups died during the procedure. After IR, the pups were returned to the dams until weaning. A single absorbed dose of 8 Gy was administered. This dose is equivalent to approximately 18 Gy to human, respectively, when delivered in repeated 2 Gy fractions, according to the linear quadratic model (Fowler 1989), and an α/β-ratio of 3 for late effects in the normal brain tissue. Sham control animals were anaesthetized but not subjected to IR.

Induction of HI brain injury

Mice were subjected to HI injury in the left hemisphere on P60, according to the Rice-Vannucci model (Rice et al. 1981), adapted for adult mice (Zhu et al. 2005). Animals were anesthetized with isoflurane (5% for induction, 1.5–2.0% for maintenance) in a mixture of nitrous oxide and oxygen (1 : 1). The duration of anesthesia was less than 5 min. The left common carotid artery was cut between double ligatures of prolene sutures (6.0). After the surgical procedure, the wounds were filled with lidocaine for local analgesia. The mice were returned to their cages for 1 h and then placed in a chamber perfused with a humidified gas mixture (10% oxygen in nitrogen) for 25 min at 36°C (Zhu et al. 2005). Following hypoxic exposure, the mice were returned to their cages. The thymidine analog BrdU (Roche, Mannheim, Germany, 5 mg/mL, dissolved in 0.9% saline), was injected intraperitoneally at a dose of 50 mg/kg, once daily for 2 days, starting from 24 h after HI. All mice were killed 4 weeks after the last BrdU injection (Fig. 1a). Control mice were subjected neither to IR nor to HI. All animal experimentation was approved by the Gothenburg Committee of the Swedish Animal Welfare Agency (application no. 244-2006).

image

Figure 1.  Brain injury evaluation. (a) The experimental design, indicating when the different insults occurred in each of the four groups. Irradiation (IR) was performed on postnatal day 10 (P10) and hypoxia-ischemia (HI) on P60. BrdU injections were performed on P61 and P62, followed evaluation on P90. (b) Representative pictures of coronal sections from the level of the hippocampus (left column) and the striatum (right column) showing MAP2 stainings of control (Cont), irradiated (IR), hypoxic-ischemic (HI) and irradiated and subsequently hypoxic-ischemic (IR + HI) brains. (c) Infarct volumes (MAP2-negative volumes) were seen only after HI, and they were significantly increased after IR. (d) Pathological scores of the cortex (Cx), hippocampus (Hipp), striatum (Str) and thalamus (Tha) after HI alone or in combination with IR. (e) Tissue loss at P90 (loss of MAP2-positive tissue) in the ipsilateral compared with the contralateral hemisphere was observed after IR alone, more so after HI and synergistically after combination of IR and HI. *p < 0.05, **p < 0.01, ***p < 0.001.

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Immunohistochemistry

The animals were deeply anesthetized with phenobarbital and perfusion-fixed with 5% formaldehyde in 0.1 M phosphate-buffered saline (PBS), followed by immersion fixation in the same fixative for 24 h at 4°C. After dehydration with graded ethanol and xylene, the brains were paraffin-embedded, serial cut in 5 μm coronal sections and mounted on glass slides. Every 100th section (typically 12 sections) was stained for MAP2. Every 50th section from the hippocampus level (typically six sections) and every 25th section from the striatum level (for the SVZ, typically six sections) were stained for BrdU or phospho-histone H3. Antigen retrieval was performed by boiling the sections in 10 mM citrate buffer (pH 6.0) for 10 min. Sections were incubated for 30 min in 4% horse or donkey or goat serum in PBS in order to block non-specific binding. Monoclonal mouse anti-MAP2 (1 : 1000, clone HM-2, Sigma, St Louis, MO, USA), monoclonal rat anti-BrdU (1 : 100, 5 μg/mL; clone: BU1/75, Oxford Biotechnology Ltd. Oxfordshire, UK), rabbit anti-phospho-histone H3 (ser10) (1 : 500, 2 μg/mL, Upstate, Temecula, CA, USA), goat anti-IL-18 (sc-6179, 4 μg/mL, Santa Cruz Biotechnology, Santa Cruz, CA, USA) primary antibody was applied and incubated at 20°C for 60 min, followed by biotinylated horse anti-mouse (1 : 200, Vector Laboratories, Burlingame, CA, USA), donkey anti-rat IgG (H + L) (1 : 200, 5.5 μg/mL; Jackson ImmunoResearch Lab. PA, USA), goat anti-rabbit (1 : 200, Vector Laboratories, Burlingame, CA, USA) or biotinylated horse anti-goat IgG (2 μg/mL) secondary antibody for 60 min at 20°C. Endogenous peroxidase activity was blocked with 3% H2O2 in PBS for 10 min. Visualization was performed using Vectastain ABC Elite (Vector Laboratories, Burlingame, CA, USA) with 0.5 mg/mL 3,3′-diaminobenzidine enhanced with 15 mg/mL ammonium nickel sulfate, 2 mg/mL beta-d glucose, 0.4 mg/mL ammonium chloride, and 0.01 mg/mL beta-glucose oxidase (all from Sigma).

The phenotype of BrdU-labeled cells was determined using antibodies against NeuN and Iba1 to detect mature neurons and microglia, respectively. Antigen recovery, was performed as above, followed by incubation with rat anti-BrdU (1 : 100, 5 μg/mL; clone: BU1/75, Oxford Biotechnology Ltd. Oxfordshire, UK) together with either mouse anti-NeuN monoclonal antibody (1 : 200, 5 μg/mL; clone: MAB377, Chemicon, Temecula, CA, USA) or rabbit anti-Iba1 antibody (1 : 1000, 0.5 μg/mL; Wako, Osaka, Japan) in PBS at 20°C for 60 min. After washing, the sections were incubated with secondary antibodies, Alexa Fluor 488 donkey anti-rat IgG (H + L), combined with either Alexa Fluor 555 donkey anti-mouse IgG (H + L) or Alexa Fluor 555 donkey anti-rabbit IgG (H + L) at 20°C for 60 min. All secondary antibodies were from Jackson ImmunoResearch Lab, and were diluted 1 : 1000. After washing, the sections were mounted using Vectashield mounting medium.

Injury evaluation

Brain injury was evaluated by infarct volume, neuropathological scoring and the volume of total hemispheric tissue loss. The MAP2-positive and -negative areas in each section were measured using Micro Image (Olympus, Japan). The infarct volume was calculated from the MAP2-negative areas according to the Cavalieri principle using the following formula: V = ΣA × P × T, where V = total volume, ΣA = the sum of area measurements, P = the inverse of the sampling fraction, and T = the section thickness. The total hemispheric tissue loss was calculated as the MAP2-positive volume in the contralateral hemisphere minus the MAP2-positive volume in the ipsilateral hemisphere. The neuropathological score for the cortex, hippocampus, striatum and thalamus was assessed as described previously (Zhu et al. 2005). Briefly, serial sections from each brain were stained for MAP2 and scored by an observer blinded to the treatment of the animals. The cortical injury was graded from 0 to 4, where 0 indicates no observable injury, 1 indicates one to three small, isolated infarctions, 2 indicates one to several slightly larger, isolated infarctions, 3 indicates a moderate, confluent infarction and 4 confluent infarction encompassing most of the cerebral cortex. The damage in the hippocampus, striatum and thalamus was assessed both with respect to hypotrophy (0–3, where 0 indicates no atrophy, 1–3 indicate mild, moderate and extensive atrophy, respectively) and observable cell injury/infarction (0–3, where 1–3 indicate mild, moderate and severe infarction, respectively) resulting in a neuropathological score for each of these three brain regions (0–6). The hippocampal granule cell layer (GCL) volumes and SVZ volumes were measured in sections stained with thionin/acid fuchsin. The GCL and SVZ areas in both hemispheres were traced and the volume was calculated from a series of sections with a 250 μm interval for the GCL or 125 μm for the SVZ, using the Stereo Investigator software (MicroBrightField, Magdeburg, Germany).

Cell counting

In every 50th section, area contours were drawn and measured. BrdU-positive cells were counted in the hippocampus, including the GCL in the dentate gyrus and the whole cornu ammonis (CA1-3), as well as in the cortex and striatum, using the Stereo Investigator software (MicroBrightField, Magdeburg, Germany). Phospho-histone H3-positive (P-HH3) cells were counted only in the GCL of the hippocampus. The number of positive cells was expressed as the total number per GCL. The BrdU-positive cells in the cortex and striatum were expressed as the total number per mm3. For phenotypic determination, at least 50 BrdU-positive cells were counted using a confocal laser scanning microscope (Leica TCS SP2, Wetzlar, Germany) in each brain region and the ratio of double-labeled cells was calculated separately for each brain and region. The total number of BrdU-positive cells of each phenotypic lineage was calculated by applying the specific ratios to the total number of BrdU-positive cells for each brain region, thereby generating the total number of newborn neurons and microglia surviving until P90. The numbers of IL-18-positive cells in different brain regions were counted in three sections, 250 μm apart, from each brain and expressed as the average number per mm2.

Statistics

All data are expressed as mean ± SEM. An unpaired t-test was used when comparing two groups. anova with Fisher’s post-hoc test was used when comparing three or more groups.

Results

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. References

Irradiation aggravated subsequent HI injury

The brain injury was evaluated using infarct volume, pathological score and volume of total hemispheric tissue loss. Infarcts were only seen after HI (Fig. 1b). The infarct volume after HI alone was 5.1 ± 1.6 mm3 (n = 18) and if the mice received 8 Gy IR 50 days prior to HI, the infarct volume was 9.8 ± 1.8 mm3 (n = 14, p < 0.05) (Fig. 1c). The injury in different brain regions was evaluated using pathological scoring, and the score was significantly increased in the cortex (p < 0.01), hippocampus (p < 0.001) and striatum (p < 0.05), but not the thalamus, if mice were irradiated 50 days prior to HI (Fig. 1d). The total hemispheric tissue loss after HI was 18.2 ± 5.8 mm3 and if the mice had been irradiated 50 days prior to HI the tissue loss was 32.0 ± 3.5 mm3 (n = 14, p < 0.05) (Fig. 1e). The total tissue loss reflects infarct volume plus subsequent degeneration and/or lack of growth of brain tissue in the ipsilateral compared with the contralateral hemisphere. For example, the tissue loss after HI alone was more than three times larger than the infarct volume. The GCL, where neurogenesis occurs throughout life, is sensitive to IR and the volume was significantly decreased after IR (18.5%, n = 8, p < 0.001) (Fig. 2b). After HI, the decrease was 24.8% (p < 0.05) and pre-treatment with IR resulted in a synergistic decrease of the GCL (67.4%, p < 0.001) (Fig. 2b). The SVZ, another neurogenic area, is also sensitive to IR, but unlike the GCL HI did not affect the volume of the SVZ and there was no observable additive or synergistic effect after IR + HI (Fig. 2c and d).

image

Figure 2.  Irradiation aggravated injury in the DG but not the SVZ after HI. (a) Representative pictures of the hippocampus showing HE staining of control (Cont), irradiated (IR), hypoxic-ischemic (HI) and irradiated and subsequently hypoxic-ischemic (IR + HI) brains. The white arrows point to the HI-injured CA1 area and black arrows point to the granule cell layer (GCL). (b) The average GCL volume in the different groups in the contralateral (Contral) and ipsilateral (Ipsilat) hemispheres. (c) Representative pictures of the SVZ in the different groups. Black arrows point to the ipsilateral SVZ. (d) The average SVZ volume in the different groups in the contralateral and ipsilateral hemispheres. *p < 0.05, **p < 0.01, ***p < 0.001.

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IR transiently enhanced post-ischemic proliferation

BrdU injections on day 1 and 2 after HI (P61 and 62) enabled us to label a population of cells that were proliferating at that time (Fig. 3a, d and e), followed by counting and phenotyping of the cells that survived until P90. The total number of BrdU-positive cells in the ipsilateral GCL was significantly decreased after IR (−41.2%, p < 0.05) (Fig. 3b), but not in the CA (Fig. 3c), striatum (Fig. 3f) or cortex (Fig. 3g), as expected from earlier studies (Tada et al. 2000; Fukuda et al. 2004; Naylor et al. 2008; Hellström et al. 2009). After HI alone, however, the number of BrdU-labeled cells in the GCL increased (+74%, p < 0.01) and this increase was further enhanced if the brains had been irradiated (+213%, p < 0.05) (Fig. 3b). A similar tendency but even more pronounced was seen also in non-neurogenic regions, including the CA (HI: +1000%, IR + HI: +5991%, Fig. 3c), striatum (HI: 3617%, IR + HI: +5750%, Fig. 3f) and cortex (HI: +4033%, IR + HI: +4650%, Fig. 3g). One month after the insult, at P90, proliferation in the GCL was assessed by phospho-histone H3 (P-HH3) staining. P-HH3-positive cells were not significantly decreased after IR alone in these mice. This indicates that the IR-induced decrease in proliferation had recovered in the DG 80 days after IR (Fig. 4), unlike the findings in rats after irradiation at P9 (Hellström et al. 2009). HI alone did not reduce proliferation significantly either, but if HI was preceded by IR, proliferation was reduced (Fig. 4b), approximately corresponding to the extent of reduction of the GCL volume (Fig. 2b). Together, the BrdU incorporation and P-HH3 stainings indicate that overall proliferation was increased early after HI (as judged by BrdU-incorporated cells surviving until P90) (Fig. 3b) and that this was further increased if the brains had previously been irradiated, despite the fact that IR alone decreased proliferation.

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Figure 3.  BrdU incorporation after IR and/or HI. (a) Representative microphotographs of cells in the hippocampus labeled with BrdU at P61–62 that survived to P90 in controls (Cont), IR, HI and IR + HI. (b and c) The average number of BrdU-positive cells in the DG (GCL + SGZ) and CA of both ipsilateral (Ipsilat) and contralateral (Contral) hemispheres in the different groups. (d and e) Representative microphotographs of BrdU labeling in the striatum and cortex in different groups. (f and g) Quantification of BrdU-labeled cells in both ipsilateral and contralateral of different groups. *p < 0.05, **p < 0.01, ***p < 0.001.

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image

Figure 4.  Cell proliferation in DG after IR and/or HI. (a) Representative microphotographs of phospho-histone H3 (P-HH3) staining in the DG at P90 in controls (Cont), IR, HI and IR + HI. (b) Quantification of the number of P-HH3-positive cells in the DG (GCL + SGZ) in both the ipsilateral (Ipsilat) and contralateral (Contral) hemispheres from the different groups. ***p < 0.001.

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IR decreased neurogenesis whereas HI increased microglia proliferation and caused persistent inflammation

The phenotype of the BrdU-labeled cells was investigated by immunofluorescent labeling (Fig. 5a and b). The number of BrdU/NeuN double-positive cells, reflecting neurons born on P61 and P62 surviving to P90, was significantly decreased in irradiated mice when compared with controls (−48%, p < 0.05) (Fig. 5c). HI alone had no significant effect on the number of BrdU/NeuN double-positive cells compared with controls. Interestingly, if the brains had been irradiated 50 days before HI, the number of BrdU/NeuN double-labeled cells was no longer significantly decreased, compared with controls, as after IR alone (Fig. 5c). The number of BrdU/Iba1 double-positive cells, reflecting microglia born on P61 and P62 surviving to P90, increased +1080% (p < 0.01) in the GCL after HI alone, and if HI was preceded by IR, this number was further increased almost three-fold (p < 0.05) (Fig. 5d). Similarly, increased microglia proliferation was seen also in the CA (+5740% after HI alone, p < 0.01, and a further four-fold increase after IR + HI, p < 0.01) (Fig. 5e), in the striatum (+1904% after HI alone, p < 0.05, and a further more than two-fold increase after IR + HI, p < 0.05) (Fig. 5f) and in the cortex (+3330% after HI alone, p < 0.01 and a further +38% after IR + HI) (Fig. 5g). The increase after IR + HI compared with HI alone was clearly synergistic, because the increase after IR alone was not significant in any of the regions studied. To investigate if this dramatic increase in the number of newborn microglia (approximately 30- to 40-fold compared with controls) in mice subjected to IR + HI resulted in more pronounced inflammation, the expression of the proinflammatory cytokine IL-18 was investigated in tissue sections (Fig. 6a). The expression of IL-18, as judged by the number of cells immunopositive for IL-18, was highest in the IR + HI group in all the examined brain regions (Fig. 6b–e), although this was most pronounced in the GCL and CA of the hippocampus (Fig. 6b–c). Notably, this increase was persistent, being detectable still 80 days after IR and 30 days after HI (Fig. 6b–e).

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Figure 5.  Neurogenesis and microglia proliferation after IR and/or HI. (a) Representative microphotographs of BrdU/NeuN immunofluorescence staining. (b) Representative microphotographs of BrdU/Iba1 immunofluorescence staining. (c) The bar graph shows the average number of BrdU/NeuN double-positive cells in the DG, i.e. neurons born on P61-62 and surviving to P90. (d and e): The bar graphs show the average number of BrdU/Iba1 double-positive cells in the DG and CA, respectively, i.e. microglia born on P61-62 and surviving to P90. (f and g): The bar graphs show the density of BrdU/Iba1 double-positive cells in the infarct border zone area of striatum and cortex, respectively. *p < 0.05, **p < 0.01, ***p < 0.001.

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Figure 6.  Immunostaining of IL-18 after IR and /or HI. (a) Representative microphotographs of IL-18 immunostaining in the DG, CA, striatum and cortex on P90. (b and c) The bar graphs show quantification of IL-18-positive cells in the DG and CA, respectively, in the different groups. (d and e) The bar graphs show quantification of IL-18-positive cells in the striatum and cortex, respectively, in the different groups. *p < 0.05, **p < 0.01, ***p < 0.001.

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Discussion

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. References

This is the first report, to the best of our knowledge, demonstrating that irradiation to the young brain aggravates a subsequent ischemic injury in adulthood. Irradiation has been shown to have multiple effects on the brain. Areas with neurogenesis, the dentate gyrus (DG) in the hippocampus and the subventricular zone (SVZ), are particularly susceptible to IR-induced DNA damage resulting in apoptosis or mitotic catastrophe (Peissner et al. 1999; Tada et al. 2000; Gudkov and Komarova 2003; Fukuda et al. 2004, 2005; Naylor et al. 2008). These effects are age-dependent and the immature brain is more vulnerable to IR (Fukuda et al. 2005). Previously we have shown that moderate, clinically relevant doses of IR to developing rodent brains caused extensive damage to stem and progenitor cells and significant reduction in the size of the proliferative areas of the brain, the GCL of the DG and the SVZ, as well as reduced myelination and even lower final brain weight (Fukuda et al. 2004; Zhu et al. 2007). These deficits can occur at low doses even in the absence of detectable histopathology, such as neuronal loss or white matter damage (Crossen et al. 1994). In this study, IR alone to the immature (P10) brain caused significant reduction of DG volume, due to inhibited growth rather than loss of tissue, as well as reduction of total hemispheric volume, when the mice were evaluated at the age of P90. If the brain is still growing at the time of IR, the reduction of final size will be more pronounced (Fukuda et al. 2005), whereas a single dose of IR up to 10 Gy to the adult rat brain showed no apparent morphological effects (Hodges et al. 1998). The side effects of IR, including cognitive deficits, are dose-dependent (Armstrong et al. 2004). It is, however, not clear from the literature if the effect on neurogenic regions and white matter formation are sufficient to explain the total tissue loss. Presumably, other mechanisms, including vascular and inflammatory changes, also contribute to these developmental aberrations (Monje et al. 2002). Importantly, the total hemispheric volume was decreased much more than would be expected from only reducing the size of the neurogenic regions and the lack of white matter development. Large non-neurogenic regions like the cortex and striatum displayed increased pathological scores in the brains that had been irradiated 80 days earlier. It is possible that the regenerative mechanisms activated after cerebral ischemia (Liu et al. 1998; Arvidsson et al. 2002; Raber et al. 2004a; Qiu et al. 2007; Naylor et al. 2008; Zhu et al. 2009) are more important for the final outcome than previously recognized. Recently, we demonstrated that IR to the brains of P9 mice reduced the levels of stem cells, neural precursors and neurogenesis in the DG, perturbed the integration of newborn neurons at P90 and even caused hyperactive behavior (Naylor et al. 2008). All these changes could be significantly ameliorated by 4 weeks of voluntary running (Naylor et al. 2008) indicating that the brain harbors a considerable regenerative capacity. The study on the effects of voluntary running, together with the present study, indicate that powerful dynamic changes can occur both before and after an insult and that these can greatly influence the final outcome. An increasing number of patients treated for childhood malignancies survive to adulthood and the present study indicates that the brains of irradiated children are more susceptible to subsequent ischemic insults, such as stroke. It is therefore imperative to investigate the nature of this ‘memory’ imprinted in the brains after irradiation. If this ‘memory’ can be erased or modified it is conceivable that the resistance of the brain to insults can be restored, and perhaps even the progressive decline in cognitive function seen after radiotherapy could be ameliorated.

Neurogenesis in the DG of the mammalian brain persists throughout life but decreases with age (Kuhn et al. 1996). It can be stimulated by both physiological conditions, such as enriched environment and physical exercise (Brown et al. 2003; Naylor et al. 2008), and pathological conditions, such as ischemic brain injury (Liu et al. 1998; Qiu et al. 2007; Zhu et al. 2009). In this study, HI alone at P60 increased BrdU incorporation during the two days following the insult. IR alone at P10, however, resulted in decreased BrdU incorporation at P61–62, but if the irradiated brains were subjected to HI, BrdU incorporation was synergistically increased (Fig. 3). The large increase in BrdU incorporation could not be explained by neurogenesis, though, because BrdU/NeuN labeling was not significantly altered (Fig. 5c). Interestingly, while neurogenesis was not increased by HI alone, the ischemic insult did appear to increase neurogenesis in the irradiated brains, but only back to control level. This observation is consistent with the finding that voluntary running increased neurogenesis more in the DG of irradiated than control brains (Naylor et al. 2008), indicating that the down-regulation of neurogenesis observed in irradiated brains can be at least partly overcome by both physiological (e.g. exercise) and pathological (e.g. ischemia) stimuli.

The majority of the BrdU-labeled cells were identified as microglia (Iba1-positive) (Fig. 5b and d–g), the endogenous phagocytes and antigen-presenting cells in the brain. We have previously demonstrated increased formation of microglia after HI in neonatal and juvenile mice, both in the hippocampus (Qiu et al. 2007) as well as in the cortex and striatum (Zhu et al. 2009), and this increase of microglia in the adult brain was concurrent with increased expression of the proinflammatory markers CCL-2 and IL-18 three days after the insult (Qiu et al. 2007; Naylor et al. 2008; Zhu et al. 2009). In the present study, we evaluated the expression of IL-18 much later, 30 days after the insult, demonstrating persistent, increased expression (Fig. 6). Emerging evidence suggests that neuroinflammation after IR is a negative regulator of neurogenesis (Monje et al. 2003). Furthermore, chronic treatment with minocycline after cerebral ischemia was shown to reduce microglia activation and enhance neurogenesis in the DG (Liu et al. 2007). However, microglia are also thought to have an instructive role in brain restoration, contributing to the maintenance of neurogenesis (Aarum et al. 2003). Specific inflammatory mediators have been shown to play pivotal regulatory roles in neurogenic and other regenerative responses (Yan et al. 2007). Further work is needed to elucidate the roles of microglia and consequences of inflammation after IR and HI, but injury-induced inflammation may be an important target for therapeutic interventions.

In conclusion, this study demonstrates that a moderate dose of ionizing radiation to the immature brain induces persistent changes not only to the neurogenic regions. When the animals became adult, ischemic brain injury was synergistically exacerbated not only in the hippocampus, but also in the cortex and striatum. This raises a cautionary note for the increasing number of children who are treated with radiotherapy and survive their disease. Our results indicate that the therapy may render their brains more susceptible to future insults and that this should be taken into consideration in future studies of late effects in adult survivors. From a neurobiological point of view, it remains to be elucidated if the increased susceptibility to ischemic injury in the irradiated brain is due to increased degeneration or decreased regeneration, or both.

Acknowledgments

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. References

This work was supported by the Swedish Research Council, the Swedish Childhood Cancer Foundation (Barncancerfonden), Swedish governmental grants to scientists working in health care (ALF), Torsten and Ragnar Söderbergs stiftelse, the National Natural Science Foundation of China (to CZ:30470598), the King Gustav V Jubilee Clinic Research Foundation (JK-fonden), the Wilhelm and Martina Lundgren Foundation, the Frimurare Barnhus Foundation, the Gothenburg Medical Society, the Swedish Society of Medicine, the Swedish Research Links Program through the Swedish International Development Cooperation Agency (SIDA).

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  5. Discussion
  6. Acknowledgments
  7. References
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