Role of fractalkine–CX3CR1 pathway in seizure-induced microglial activation, neurodegeneration, and neuroblast production in the adult rat brain

Neurobiology of Disease 74 (2015) 194–203

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Role of fractalkine–CX3CR1 pathway in seizure-induced microglial activation, neurodegeneration, and neuroblast production in the adult rat brain Idrish Ali, Deepti Chugh, Christine T. Ekdahl ⁎ a b

Inflammation and Stem Cell Therapy Group, Wallenberg Neuroscience Center, Division of Clinical Neurophysiology, Lund University, Sweden Lund Epilepsy Center, Lund University, SE-221 84 Lund, Sweden

a r t i c l e

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Article history: Received 4 August 2014 Revised 21 October 2014 Accepted 12 November 2014 Available online 21 November 2014 Keywords: CX3CR1 Fractalkine Epilepsy Neurogenesis Inflammation Neurodegeneration

a b s t r a c t Temporal lobe seizures lead to an acute inflammatory response in the brain primarily characterized by activation of parenchymal microglial cells. Simultaneously, degeneration of pyramidal cells and interneurons is evident together with a seizure-induced increase in the production of new neurons within the dentate gyrus of the hippocampus. We have previously shown a negative correlation between the acute seizure-induced inflammation and the survival of newborn hippocampal neurons. Here, we aimed to evaluate the role of the fractalkine–CX3CR1 pathway for these acute events. Fractalkine is a chemokine expressed by both neurons and glia, while its receptor, CX3CR1 is primarily expressed on microglia. Electrically-induced partial status epilepticus (SE) was induced in adult rats through stereotaxically implanted electrodes in the hippocampus. Recombinant rat fractalkine or CX3CR1 antibody was infused intraventricularly during one week post-SE. A significant increase in the expression of CX3CR1, but not fractalkine, was observed in the dentate gyrus at one week. CX3CR1 antibody treatment resulted in a reduction in microglial activation, neurodegeneration, as well as neuroblast production. In contrast, fractalkine treatment had only minor effects. This study provides evidence for a role of the fractalkine–CX3CR1 signaling pathway in seizure-induced microglial activation and suggests that neuroblast production following seizures may partly occur as a result of microglial activation. © 2014 The Authors. Published by Elsevier Inc. This is an open access article under the CC BY-NC-SA license (http://creativecommons.org/licenses/by-nc-sa/3.0/).

Introduction Mesial temporal lobe epilepsy (MTLE), one of the most common forms of focal epilepsies, is associated with a pathology in the hippocampus characterized by an acute inflammatory response and activated parenchymal microglial cells, neurodegeneration, aberrant synaptic reorganizations and an increase in adult neurogenesis (Bengzon et al., 1997; Parent et al., 1997; De Simoni et al., 2000; Crespel et al., 2002; Pitkanen and Sutula, 2002; Scharfman and Pedley, 2007; Ravizza et al., 2008; Lu et al., 2009). A growing body of literature from both clinical and experimental epilepsy research has during the last decade focused in particular on how the inflammatory processes may contribute to the development and severity of epilepsy (Vezzani et al., 2011). Brain inflammation can affect both excitotoxicity-mediated neurodegeneration (Vezzani, 2005; Ulmann et al., 2013) as well as modulate the seizure-induced neurogenesis (Jakubs et al., 2006; Ekdahl et al.,

⁎ Corresponding author at: Inflammation and Stem Cell Therapy Group, Division of Clinical Neurophysiology, Lund Epilepsy Center, BMC A11, Sölvegatan 17, Lund University, SE-221 84 Lund, Sweden. Fax: +46 46 2220560. E-mail address: [email protected] (C.T. Ekdahl). Available online on ScienceDirect (www.sciencedirect.com).

2009). Acute microglial activation following epileptic seizures is detrimental to the survival of newly formed neurons (Ekdahl et al., 2003). Both seizure-induced pathology and lipopolysaccharide-induced brain inflammation alter the functional properties and the expression of synaptic proteins, including adhesion molecules and scaffolding proteins, of adult born hippocampal neurons (Jakubs et al., 2006; Wood et al., 2011; Jackson et al., 2012; Chugh et al., 2013). In addition, it has been speculated that microglia may regulate synaptic pruning and transmission in adult born neurons (Ekdahl, 2012). Since newborn neurons are thought to be involved in memory formation during normal physiological conditions (Jessberger et al., 2009; Deng et al., 2010), the additional number of neurons born after seizures, may be an important restorative capacity of the brain. The acute inflammatory response following seizures, which compromises neurogenesis, may therefore be detrimental for brain recovery. The function of new neurons in a seizure-environment may, however, be different from physiological conditions, though previous observations from new seizure-induced neurons remaining within the granule cell layer (GCL) of the dentate gyrus have suggested that they may counteract hyperexcitability (Jakubs et al., 2006; Jackson et al., 2012). Conversely, following more severe seizures, some new neurons migrate aberrantly into the dentate hilus and there they may instead propagate excitability (Parent et al., 1997; Scharfman et al., 2000;

http://dx.doi.org/10.1016/j.nbd.2014.11.009 0969-9961/© 2014 The Authors. Published by Elsevier Inc. This is an open access article under the CC BY-NC-SA license (http://creativecommons.org/licenses/by-nc-sa/3.0/).

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Parent and Lowenstein, 2002; Bielefeld et al., 2014). The role of inflammation for these aberrant neurons remains unclear. One way to further understand how inflammation and other seizure-induced brain pathologies are related is to study single potential signaling pathways between microglia and neurons. Various inflammatory mediators which are known to regulate microglial activation (Cardona et al., 2006; D'Haese et al., 2012), have also been implicated in adult neurogenesis and synaptic transmission (Butovsky et al., 2006; Lauro et al., 2010; Scianni et al., 2013). Fractalkine is an inflammatory chemokine secreted by neurons and astrocytes. It acts on G-protein coupled CX3CR1 receptors, present primarily on microglial cells, and regulates its activation, migration and phagocytic activity (Harrison et al., 1998; Fuhrmann et al., 2010; Noda et al., 2011). In addition, the fractalkine–CX3CR1 signaling pathway has also been implicated in tuning neurogenesis and synaptic transmission (Bachstetter et al., 2011; Paolicelli et al., 2011). Furthermore, recent studies have depicted a role of the fractalkine–CX3CR1 pathway in the pathogenesis of epilepsy and associated cell death (Yeo et al., 2011; Xu et al., 2012). An increased expression of fractalkine and CX3CR1 receptors is reported in the surgically resected brain samples from MTLE patients and in experimental animal models of MTLE (Xu et al., 2012; Roseti et al., 2013). In the present study, we aimed to investigate the role of the fractalkine–CX3CR1 signaling pathway for acute seizure-induced microglial activation, neurodegeneration, and neuroblast production following partial status epilepticus (SE). To achieve this, we performed continuous intraventricular infusion of anti-CX3CR1 antibody or recombinant fractalkine for seven days immediately following electricallyinduced SE in the temporal lobe of adult rats.

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according to the protocol described in Jackson et al. (2012). Electrodeimplanted rats with no stimulations served as controls for the SE rats. Initially, afterdischarge thresholds were determined for each rat by applying a square wave biphasic pulse (50 Hz) of 1 s train duration at a starting intensity of 10 μA, then increasing by 10 μA every 1 min until a 10 s afterdischarge was evoked. For inducing SE, suprathreshold stimulation was applied for 1 h, with interruptions in the stimulation every 9th minute to record electroencephalographic (EEG) activity for 1 min. Subsequently, the ictal EEG activity was recorded for another 2 h and the behavioral seizures were classified according to Racine's (1972) scale. Only rats that displayed self-sustained ictal EEG activity for 2 h in the temporal lobe as well as a partial seizure semiology were included in this study. The ictal EEG activity was then completely interrupted by administering pentobarbital (65 mg/kg, intraperitoneal injection). CX3CR1GFP/+ heterozygous mice underwent electrode implantation surgery (coordinates: 2.9 mm posterior and 3.0 mm lateral from bregma; and 3.0 mm ventral from dura) (Franklin and Paxinos, 1997) and SE induction, similar to the protocol described for rats. Intracerebroventricular drug infusion

Materials and methods

Once rats were anesthetized following SE, the brain infusion cannula was connected with osmotic pumps (1007D, Alzet) carrying either of the following drugs: rabbit anti-CX3CR1 antibody (20 μg/ml (low dose) or 60 μg/ml (high dose) (Abcam, UK)), recombinant rat fractalkine (2 μg/ml, R&D Systems, USA) or vehicle (phosphate buffer saline). Infusion rate was 0.5 μl/h. Non-stimulated rats were also connected with osmotic pumps carrying vehicle to serve as controls. The osmotic pumps were placed in the subcutaneous pocket in the dorsal region of the neck.

Animals

Tissue processing

Adult male Sprague–Dawley rats (n = 77) weighing between 200 and 250 g were procured from Charles River (Germany). Rats with generalized-status epilepticus semiology, absence of sustained seizures after electrical stimulations, incorrect electrode placements or hydrocephalous due to hippocampal injury were excluded from the study. After exclusions, 33 animals served as non-stimulated or SEvehicle treated controls, whereas 28 animals were used for either of the treatment groups (SE-CX3CR1 and SE-fractalkine) included in this study. The animals were either used for immunohistochemical analysis or ELISA/Western blots studies. The number of animals used for each assessment is described in the figure legends as well as in the relevant text in the Results section. Additionally, we used CX3CR1GFP/+ heterozygous mice (n = 4), 2 non-stimulated controls and 2 SE mice (originally bred by Dr. Steffen Jung, Rehovot, Israel, and kindly provided by Dr. William Agace, Lund, Sweden). The animals were housed in a 12 h light/dark cycle with ad libitum food and water. All experimental procedures were performed in accordance with the guidelines set by the MalmöLund Ethical Committee for the use and care of laboratory animals.

Seven days after SE induction, rats and mice were either used for immunohistochemical studies or ELISA/Western blots. An additional cohort of rats was sacrificed at 6 h after the induction of SE to assess the levels of inflammatory cytokines using ELISA. For immunohistochemistry, rats and mice were trans-cardially perfused with 0.9% saline and 4% paraformaldehyde. The brains were collected and post-fixed in paraformaldehyde overnight, after which they were kept in 20% sucrose solution at 4 °C until sectioning. The brains were cut in 30 μm sections on a freezing microtome. The sections were collected in a glycerol based anti-freeze solution distributed into ten series, and stored in − 20 °C until used for immunohistochemistry. The cohort of rats for ELISA and Western blot analysis was transcardially perfused with 0.9% saline. The brains were collected, and contralateral hippocampal tissues were extracted and immediately frozen in dry ice. The tissues were then stored in −80 °C until protein isolation.

Surgery and induction of status epilepticus (SE) Animals were anesthetized with 2% isoflurane and implanted with a bipolar insulated stainless steel electrode (Plastics One, Roanoke, VA) into the right ventral CA1/CA3 region of the hippocampus (coordinates: 4.8 mm posterior and 5.2 mm lateral from bregma; and 6.3 mm ventral from dura, toothbar set at −3.3 mm) (Paxinos and Watson, 1998) for stimulation and recording. Another electrode (unipolar) was placed between the skull and adjacent muscle to serve as ground electrode. Additionally, a brain infusion cannula (Brain Infusion Kit 1, Alzet) was implanted in the lateral ventricle (coordinates: 1.0 mm posterior and 1.5 mm lateral to bregma; and 3.5 mm ventral to the flat skull position with bregma as reference) in the ipsilateral hemisphere. Following a week of recovery, rats were subjected to electrically-induced SE

Immunohistochemistry For immunohistochemistry, the following primary antibodies were used: rabbit anti-Iba1 (1:1000, Wako, Japan), mouse anti-CD68/ED1 (1:200, AbD Serotec, Germany), goat anti-doublecortin (DCX) (1:200, Santa Cruz Biotechnology, Germany), rabbit anti-Ki67 (1:400, Leica Biosystems, Novocastra, Sweden), rabbit anti-CX3CR1 (1:100, Abcam, UK), mouse anti-NeuN (1:100, Millipore, Bioscience Research Reagents, CA, USA), rabbit anti-fractalkine (1:200, Abcam), and mouse anti-glial fibrillary acidic protein (GFAP) (Sigma, Germany). For all stainings (except for Iba1 and ED1), free-floating sections were subjected to an initial antigen retrieval step with incubation of sections in sodium citrate buffer (10 mM sodium citrate, 0.05% Tween-20, pH 6). Subsequently, sections were incubated with an appropriate primary antibody overnight at 4 °C in blocking serum, except during CX3CR1 staining when sections were incubated in primary antibody overnight at room temperature. This was followed by 2 h of incubation in secondary antibody at

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room temperature. For each immunohistochemical assessment, some brain sections went through the entire protocol without primary antibody incubation, to serve as the negative controls. The following secondary antibodies were used: Cy3-conjugated donkey anti-mouse/ rabbit (Jackson Immunoresearch, UK), biotinylated goat anti-rabbit (1:200, Vector Laboratories, UK), biotinylated horse anti-goat (1:200, Vector Laboratories), or Alexa-488 conjugated streptavidin (1:200, Invitrogen, Sweden). The sections were mounted on gelatin-coated slides and coverslipped using a glycerol-based mounting medium (DABCO, Sigma).

Fluoro-Jade staining Free-floating sections were washed with potassium phosphate buffer saline mounted on gelatin-coated slides, and dried overnight. The sections were then hydrated, pretreated with 0.06% potassium permanganate for 15 min, rinsed with distilled water and treated with 0.001% Fluoro-Jade (Histo-Chem, Jefferson, AR, USA) for 30 min, washed with distilled water, dehydrated by treatment with ethanol and xylene, and then coverslipped with Pertex mounting medium. Cell counts and image analyses

ELISA Protein samples from contralateral hippocampi were isolated as described before (Chugh et al., 2013). Total protein concentration was determined by BCA protein assay (BCA, Pierce, Rockford, USA) as per the manufacturer's instructions. Interleukin-1β (IL-1β), IL-6, IL-4, and IL-10 concentrations were determined by ELISA (Duoset, R & D Systems, MN, USA) according to the manufacturer's protocol. Briefly, 50–100 μg of a protein sample was loaded per well and incubated on a shaker for 2 h at room temperature. For each assay, samples were analyzed in duplicates, and were compared with protein standards. Plates were read on a Perkin Elmer Victor™ X multilabel plate reader. For the assessment of inflammatory mediators at 6 h after SE, cytokine levels were measured in hippocampal supernatants by sandwich immunoassay methods using a commercially available electrochemiluminescent detection system, and commercially available plates and reagents (V-PLex Pro-inflammmatory Panel 1 (rat) kit, Meso-scale Discovery (MSD), Gaithersburg, Maryland) as per the manufacturer's instructions with minor modifications. Briefly, 50 μl of the protein sample was loaded per well of the MSD plate (~100 μg of protein). The samples were incubated overnight at 4 °C with shaking. For each assay, samples were analyzed in duplicates, and compared with known concentrations of protein standard. Plates were analyzed using the SECTOR Imager 2400.

Western blot analysis Protein samples were denatured at 99 °C for 5 min in 2× Laemmli sample buffer (Bio-Rad, Germany). Total protein (10 μg) was resolved on precast 4 –15% mini-PROTEAN® TGX™ (Bio-Rad) sodium dodecyl sulfate polyacrylamide gels and transferred using Trans-Blot® Turbo™ mini nitrocellulose transfer packs (Bio-Rad). The membranes were blocked for 2 h at room temperature in Tris-buffered saline (pH 7.4) with 0.2% (w/v) Tween 20 (TBS-T) containing 5% nonfat dried milk for β-actin, gephyrin and post-synaptic density (PSD)-95 and with 3% bovine serum albumin (BSA, Sigma) for IL-1 receptor-1 (IL-1R1) and tumor necrosis factor receptor 2 (TNFR2). Membranes were then incubated for overnight at 4 °C with primary antibodies diluted in TBS-T containing 0.5% BSA. The following primary antibodies were used: mouse monoclonal anti-β actin (1:10,000, Sigma), mouse monoclonal anti-gephyrin (1:3000, Synaptic Systems, Germany), mouse monoclonal anti-PSD-95 (1:200, Abcam), rabbit polyclonal anti-TNFR2 (1:250, Santa Cruz Biotechnology), and rabbit polyclonal anti-IL1R1 (1:500, Santa Cruz Biotechnology). After washing off the primary antibody, membranes were incubated with secondary antibody diluted in 0.5% BSA for 2 h at room temperature. Secondary antibodies were either horseradish peroxidase-conjugated anti-mouse (1:5000, Sigma), or anti-rabbit (1:5000, Sigma) diluted in TBS-T containing 0.5% BSA. The membranes were then washed again for three times in TBS-T. Immunoreactive bands were visualized by enhanced chemiluminescence (Bio-Rad), and images were acquired using the Chemidoc™ XRS+ system (Bio-Rad). Band intensities were quantified using ImageJ software (NIH, USA) and β-actin was used as a loading control.

All quantifications were performed in 3–4 sections (from −3.3 mm to −4.6 mm posterior to bregma) from brain hemisphere contralateral to the stimulated region by an experimenter blind to the treatment conditions as previously described in Jakubs et al. (2006). The Iba1/ED1, DCX and Fluoro-Jade+ cell counts were performed using an Olympus BX61 epifluorescence microscope in the hippocampal GCL, dentate hilus and molecular layer (ML), while Ki67+ cells were additionally counted in the subgranular zone (SGZ), the zone comprising the innermost row of cells in the GCL along with two cell diameter thick regions at the GCL/hilar border. The data is expressed as mean number of cells/ section, based on average number of cells in 3–4 sections. Moreover, a total of 100 Iba1+ cells were analyzed for all of the three microglial morphological subtypes including ramified, intermediate and round/ amoeboid in each region of interest. The relative occurrence of each morphological subtype is expressed as the percentage of the fixed number of microglial Iba1 counts (100 cells/rat) for each region of interest. For estimating the co-labeling of GFP+ with Iba1+, GFAP+ and NeuN+ cells, respectively, 50–60 cells were analyzed per section from a total of 4 sections per mouse in the GCL/SGZ, ML, and dentate hilar region in the naïve and SE-induced CX3CR1GFP/+ heterozygous mice. For CX3CR1 expression, four representative images were taken at a 20× objective (from − 3.3 to − 4.6 mm posterior to bregma) with a BX61 epifluorescence microscope and the fluorescent intensity on the microglial cells were qualitatively graded from 1 to 5, with 1 being the lowest intensity and 5 being the highest. The values were then expressed as median ± range for each experimental group. To quantify fractalkine and GFAP expression, four representative images from the hippocampal GCL and dentate hilus (from − 3.3 to − 4.6 mm posterior to bregma) were acquired using a BX61 epifluorescence microscope with 20× objective. The images were then analyzed as described previously in Chugh et al. (2013) using the ImageJ software. The fluorescence intensities are reported after subtracting the intensity measurements from that of negative control samples. Statistical analyses Statistical analyses (Graphpad Prism software) were performed using the unpaired Student's t-test when comparing 2 groups. Statistical analyses for CX3CR1 expression was performed using the Mann–Whitney test. Microglial morphology profile data was analyzed by using two-way ANOVA. All data are expressed as mean ± SEM, except for CX3CR1 expression, which is expressed as median ± range. Differences were considered statistically significant at p ≤ 0.05. Results Acute pathophysiological changes in the dentate gyrus of the hippocampus one week after status epilepticus At one week following SE, we observed profound microglial activation in the hippocampus with increased microglial cell numbers as well as the presence of activated microglial phenotypes. The number of Iba1+ cells was increased in the GCL and dentate hilus, but not in the ML in vehicle-treated SE (SE) compared to vehicle-treated non-stimulated

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control rats (NSC) (Fig. 1A). Activated/phagocytic microglial cells, quantified as the percentage of Iba1+ cells expressing ED1, were increased in the GCL, ML, and dentate hilus (Fig. 1B). In addition, we assessed the morphological phenotypes of microglia; ramified/surveying (Fig. 1C), intermediate (Fig. 1D) and round/ameboid (Fig. 1E) morphology. In SE rats, the percentage of surveying/ramified microglia was decreased by about 85% along with an increased intermediate and round morphology, which suggests a more activated phenotype, in the GCL (Fig. 1F), ML (Fig. 1G) and dentate hilus (Fig. 1H). However, despite a strong increase in the microglial activation one week following SE induction, we could not detect any changes in the protein expression levels of pro-inflammatory cytokines in the hippocampus including IL-1β (373.4 ± 53.06 pg/mg in NSC vs 365.8 ± 15.48 pg/mg in SE rats; n = 5 for each group) and IL-6 (233.4 ± 54.29 pg/mg NSC vs 255.9 ± 85.83 pg/mg SE), or anti-inflammatory cytokines

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including IL-4 (18.93 ± 3.30 pg/mg NSC vs 16.66 ± 2.24 pg/mg SE) and IL-10 (12.08 ± 1.01 pg/mg NSC vs 16.18 ± 3.31 pg/mg SE). Similarly, no changes were detected with Western blots in mean protein levels of two cytokine receptors; IL-1R1 (100.0 ± 3.02% NSC vs 94.8 ± 11.31% SE; n = 5 for each group) or TNF R2 (100.0 ± 17.29% NSC vs 124.5 ± 21.79% SE) normalized to the expression of β-actin, at this time point. However, we did find a significant acute increase in the expression of IL-6 in the hippocampus 6 h after SE compared to non-stimulated control rats (Supplementary Fig. 1). There was a moderate increase in ongoing cell death within the hippocampus (Figs. 1I and J) in terms of the increased number of Fluoro-Jade+ cells in the GCL and dentate hilus of SE compared to NSC rats (Fig. 1K). In order to detect a possible imbalance in synaptic excitation and inhibition one week post-SE, we quantified the hippocampal protein

Fig. 1. Status epilepticus (SE)-induced increase in microglial activation, neuronal degeneration and neuroblast production in the dentate gyrus of the hippocampus in adult rats. Quantification of Iba1+ (A) and percentage of Iba1+ED1+ cells (B) in the granule cell layer (GCL), molecular layer (ML) and hilus of the dentate gyrus in vehicle-treated, non-stimulated controls (NSC) and SE. C–E, Representative photomicrographs of different microglial morphological phenotypes including ramified (Ram, C), intermediate (Inter, D) and round/ameboid (R/A, E) Iba1+ cells. F–H, Quantification of relative percentage of microglia with different morphologies in the GCL (F), ML (G) and dentate hilus (H). I–J, Representative images of FluoroJade+ (FJ+) cells (arrow heads) in NSC (I) and SE (J). K, Quantification of FJ+ cells in the GCL, ML and dentate hilus. L–M, Representative images of doublecortin+ (DCX+) cells (arrow heads) in NSC (L) and SE (M). N–O, Quantitation of DCX+ cells in the GCL/sub-granule zone (SGZ) (N) and Ki67+ cells in the SGZ of the dentate gyrus (O). Means ± SEM, n = 5 NSC and n = 4 SE group. *, p ≤ 0.05 un-paired t-test compared to controls in A, B, K, N and O and two-way ANOVA in F–H. Scale bars are 5 μm for C–E and 40 μm in I–M.

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expression of gephyrin and PSD-95, synaptic scaffolding proteins at inhibitory and excitatory synapses, respectively. However, we did not detect any significant changes (gephyrin; 100.0 ± 6.7% NSC vs 109.3 ± 7.9% SE, PSD-95; 100.0 ± 9.9% NSC vs 117.7 ± 32.5% SE, normalized to the expression of β-actin and presented as percentage change in expression relative to non-stimulated controls). Neuroblast production, estimated by the number of DCX+ cells in the GCL/SGZ of the dentate gyrus (Figs. 1L and M), was increased in SE compared to NSC, thus corroborating previous findings (Fig. 1N). Also, the number of Ki67+ cells in the SGZ was increased, which reflects the extent of cell proliferation (Fig. 1O).

Fractalkine and CX3CR1 expression in the dentate gyrus of the hippocampus one week after status epilepticus We did not detect a change in the expression of fractalkine in the GCL (mean intensities: SE 28.88 ± 1.97; n = 8 vs NSC 31.50 ± 3.28; n = 7) or the dentate hilus (SE 23.45 ± 1.56 vs NSC 30.48 ± 3.22) of SE compared to NSC rats (Figs. 2A–B). However, there was a significant increase in the CX3CR1 expression in the dentate gyrus of SE rats, when compared to NSC rats (Figs. 2C–D) (median of 4 with a range between 3.75 and 5 in intensity scores for SE; n = 4 vs median of 1 with a range of 0.25 to 1.25 for NSC; n = 5). In order to assess the microglial specificity of CX3CR1, we used CX3CR1GFP/+ heterozygous mice to confirm a 100% overlap between CX3CR1 expression and Iba1+ cells in control mice as well as one week after electrically-induced SE (Figs. 2E–F). No overlap was observed with the astrocytic and neural progenitor marker GFAP (Figs. 2G–H) or the neuronal marker NeuN (Figs. 2I–J).

Effect of fractalkine infusion on status epilepticus-induced pathology We next explored if modulating the fractalkine–CX3CR1 pathway by fractalkine infusion affects hippocampal pathology one week after SE. First, we assessed the microglial cell numbers and activation profiles in the vehicle-treated SE versus fractalkine-treated SE rats (Figs. 3A–F). Fractalkine infusion following SE, did not affect Iba1+ cell counts in the GCL, ML or dentate hilus of the hippocampus (Fig. 3A). Similarly, no changes were detected in the percentage of Iba1+ microglial cells expressing ED1 (Fig. 3B). In addition, there were no alterations in the number of proliferating Ki67+ cells (Fig. 3C) or microglial morphological phenotypes in the GCL, ML or dentate hilus (Figs. 3D–F).

Consistent with the immunohistochemical findings, we did not detect any changes in the protein expression levels of IL-1β (365.8 ± 15.48 pg/mg SE-Veh, n = 5 vs 328.8 ± 22.34 pg/mg SE-fractalkine, n = 4), IL-6 (255.9 ± 85.83 pg/mg SE-Veh vs 66.9 ± 26.67 pg/mg SE-fractalkine), IL-4 (16.66 ± 2.24 pg/mg SE-Veh vs 11.83 ± 2.3 pg/mg SE-fractalkine), or IL-10 (16.18 ± 3.31 pg/mg SE-Veh vs 12.3 ± 2.58 pg/mg SE-fractalkine) in the hippocampus. Similarly, no changes were detected in mean protein levels of IL-1R1 (94.8 ± 11.31% SE-Veh vs 101.9 ± 9.4% SE-fractalkine) or TNF R2 (124.5 ± 21.79% SEVeh vs 92.8 ± 16.7% SE-fractalkine normalized to the expression of β-actin and presented as percentage change in expression relative to non-stimulated controls rats). In addition, we observed no changes in the protein expression of gephyrin (109.3 ± 7.9% SE-Veh vs 112.0 ± 14.4% SE-fractalkine) or PSD-95 (117.7 ± 32.5 SE-Veh vs 142.6 ± 23.7% SE-fractalkine) in the hippocampus. Furthermore, fractalkine treatment did not affect the numbers of Fluoro-Jade+ cells in GCL, ML, or dentate hilus (Fig. 3G) or DCX+ cells in the GCL/SGZ (Fig. 3H). However, it did lead to an increase in the numbers of Ki67+ cells in the SGZ (Figs. 3I–K).

CX3CR1 antibody infusion decreases status epilepticus-induced microglial activation We evaluated the infusion of two different doses of CX3CR1 antibody based on Yeo et al. (2011). The lower dose of CX3CR1 antibody infusion during one week post-SE led to a significant reduction in the number of Iba1+ cells in the GCL of the hippocampus as well as a reduced percentage of Iba1+ cells expressing ED1 in the GCL, ML and dentate hilus compared to the vehicle-treated SE group (Figs. 4A–D). Also, the number of Ki67+ cells was significantly reduced in the dentate hilus, while no differences were detected in the GCL or ML (Fig. 4E). Moreover, when comparing the morphological profiles for different microglial activation states, we observed a significant interaction between the microglial morphology and treatment groups in the GCL (Fig. 4F) and ML (Fig. 4G), which was primarily due to an increase in the percentage of microglial cells with ramified morphology in the CX3CR1 antibody-treated rats. However, microglial morphology was not affected in the dentate hilus (Fig. 4H). The change in microglial activation profile induced by CX3CR1 antibody treatment was not accompanied by a change in the protein expression levels of IL-1β (365.8 ± 15.48 pg/mg SE-Veh vs 354.8 ± 22.43 pg/mg SE-CX3CR1; n = 5 in each group), IL-6 (255.9 ± 85.83 pg/mg SE-Veh vs 133.0 ± 42.17 pg/mg SE-CX3CR1), IL-4 (16.66 ± 2.24 pg/mg SE-Veh vs

Fig. 2. Expression of fractalkine and CX3CR1 in the dentate gyrus one week after SE. A–D, Representative photomicrographs of fractalkine expression in NSC (A) and SE (B) and CX3CR1 expression (arrow heads) in NSC (C) and SE (D). E–F, Representative orthogonal projections with confocal microscopy of Iba1+ cells (red) co-localized with GFP+ cells (green) in CX3CR1GFP/+ mice, NSC (E) and SE (F). G–H, Images of GFAP+ cells (red) not co-localized with GFP+ cells (green) in CX3CR1GFP/+ mice, NSC (G) and SE (H). I–J, Images of NeuN+ cells (red) not colocalized with GFP+ cells (green) in CX3CR1GFP/+ mice, NSC (I) and SE (J). Scale bar is 40 μm in A for A–D and 10 μm in E for E–J.

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Fig. 3. Effect of fractalkine treatment on SE-induced pathology. Quantification of the number of Iba1+ (A), percentage of Iba1+ED1+ cells (B), and Ki67+ (C) cells, relative percentage of microglial morphologies in the GCL (D), ML (E) and dentate hilus (F), and FJ+ (G), and DCX+ cells (H) of vehicle-treated SE (SE-Veh) and fractalkine-treated SE rats (SE-fractalkine). I–J, Representative images of Ki67+ cells (arrow heads) in the dentate gyrus of the SE-Veh (I) and SE-fractalkine groups (J). K, Quantification of Ki67+ cells in the SGZ of the dentate gyrus of the SE-Veh and SE-fractalkine groups. Means ± SEM, n = 8 and 7 for SE-Veh and SE-fractalkine respectively for A–F and K and n = 4 in each group for G and H. *, p ≤ 0.05 un-paired t-test compared to controls in A–C & G–K and two-way ANOVA in D–E. Scale bar is 40 μm.

18.28 ± 2.16 pg/mg SE-CX3CR1), or IL-10 (16.18 ± 3.31 pg/mg SE-Veh vs 14.68 ± 0.72 pg/mg SE-CX3CR1) at one week post-SE. Similarly, no changes were detected in mean protein levels of IL-1R1 (94.8 ± 11.31% SE-Veh vs 97.4 ± 8.5% SE-CX3CR1) or TNF R2 (124.5 ± 21.79% SE-Veh vs 110.2 ± 13.84% SE-CX3CR1). In contrast to changes in microglial activation, the lower dose of the CX3CR1 antibody treatment did not induce changes in fluorescent intensity of the astrocytic marker GFAP in the GCL, ML or the dentate hilus of the hippocampus (Fig. 4I). The higher dose of CX3CR1 antibody did not reduce microglial activation and numbers, or altered their morphologies in the dentate gyrus after SE (n = 4 in each group; data not shown). In fact, we even observed a trend towards an increase in the number of Iba1+ cells (p = 0.07) in the dentate hilus (data not shown). Since there were no significant changes, we opted to use the lower dose for further evaluations. CX3CR1 antibody infusion decreases status epilepticus-induced neuronal degeneration The CX3CR1 antibody infusion reduced the number of Fluoro-Jade+ cells in the dentate hilus of the hippocampus one week post-SE (Figs. 5A–C), suggesting a neuro-protective effect in the sub-region of the dentate gyrus with the highest extent of SE-induced cell death. No differences were seen in the GCL or ML (Fig. 5C). In terms of imbalance in excitatory/inhibitory scaffolding proteins, the CX3CR1 infusion did not alter the expression of gephyrin (109.3 ± 7.9%

SE-Veh vs 108.6 ± 1.4% SE-CX3CR1) or PSD-95 (117.7 ± 32.5 SE-Veh vs 110.8 ± 6.96% SE-CX3CR1) in the hippocampus. CX3CR1 antibody infusion decreases status epilepticus-induced neuroblast production Finally, we quantified the initial stages of seizure-induced neurogenesis within the hippocampus by measuring neuroblast production one week following SE. We found a 38% reduction in the number of DCX+ cells in the GCL/SGZ of CX3CR1 antibody-treated SE as compared to vehicle-treated SE rats (Figs. 6A–C). The decrease in DCX+ cells was further supported by a 53% decrease in the number of proliferating Ki67+ cells within the SGZ of the dentate gyrus (Figs. 6D–F). Discussion In the current study, we describe a role of the fractalkine–CX3CR1 pathway for the acute pathological changes in the brain following an epileptic insult. Blocking this pathway by administering the anti-CX3CR1 antibody diminished electrical SE-induced microglial activation, neurodegeneration and neuroblast formation in the adult rat hippocampus. We observed an increased expression of CX3CR1 in the hippocampus one week following SE. Increased expression of fractalkine and CX3CR1 has been reported in resected hippocampal sections from epileptic patients as well as in the animal model of MTLE (Yeo et al., 2011; Xu et al., 2012; Roseti et al., 2013). An increased level of

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Fig. 4. CX3CR1 antibody treatment decreases microglial activation one week after SE. A–B, Representative images of Iba1+ (green) and Iba1+/ED1+ (yellow) cells in vehicle-treated SE (SE-Veh) (A) and CX3CR1 antibody-treated SE rats (SE-CX3CR1) (B). Quantification of the number of Iba1+ (C), and percentage of Iba1+ ED1+ cells (D), Ki67+ cells (E), and relative percentage of microglial morphologies in the GCL (F), ML (G), and dentate hilus (H). I, Densitometric analysis of GFAP expression in the GCL, ML, and dentate hilus of the SE-Veh and SE-CX3CR1 groups. Means ± SEM, n = 8 in each group for A–H and n = 4 in each group for I. *, p ≤ 0.05 un-paired t-test compared to controls in C–E & I and two-way ANOVA in F–H. Scale bar is 40 μm.

fractalkine has also been reported in the cerebrospinal fluid from epileptic patients (Xu et al., 2012). However, we could not detect an increase in the fractalkine expression one week post-SE in our animal model. This finding is in line with a previous study (Yeo et al., 2011), which even reported a reduced fractalkine expression at 3 days after SE, despite showing an increase at earlier time points. We found no evidence for the presence of CX3CR1 in cells expressing the astrocytic and neural progenitor cell marker GFAP (Jessberger et al., 2005) or the

mature neuronal marker NeuN, but 100% co-localization was observed with the microglial cell marker Iba1, in the dentate gyrus of both naïve and SE-induced CX3CR1GFP/+ heterozygous mice. This suggests that modulation of the CX3CR1 pathway primarily affects the microglia population. Administration of the anti-CX3CR1 antibody reduced SE-induced microglial activation, in terms of increased numbers of ramified/ surveying microglia together with a reduced percentage of microglial

Fig. 5. CX3CR1 antibody treatment decreases neuronal degeneration one week after SE. A–B, Representative photomicrographs of Fluoro-jade+ (FJ+) cells (arrow heads) in the dentate gyrus of the SE-Veh (A) and SE-CX3CR1 groups (B). C, Quantification of FJ+ cells. Means ± SEM, n = 4 in each group. *, p ≤ 0.05 un-paired t-test compared to controls. Scale bar is 40 μm.

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Fig. 6. CX3CR1 antibody treatment decreases neuroblast production one week after SE. A–B, Representative images of DCX+ cells (green) in the SE-Veh (A) and SE-CX3CR1 groups (B). C, Quantification of DCX+ cells (C). D–E, Representative images of Ki67+ cells (red) in the SE-veh (D) and SE-CX3CR1 groups (E). F, Quantification of Ki67+ cells. Means ± SEM, n = 4 in each group. *, p ≤ 0.05 un-paired t-test compared to controls. Scale bar is 40 μm.

cells that were ED1+ phagocytic cells, in the hippocampal dentate gyrus. This decrease is in line with the findings of Yeo et al. (2011), where they showed reduced microglial numbers in the hippocampus, 3 days after pilocarpine-induced SE in rats treated with the same low dose (20 μg/ml) of anti-CX3CR1 antibody used in this study, from 4 days pre- and 3 days post-SE. Conversely, in an attempt to further activate the pathway, we administered recombinant fractalkine. However, the present treatment protocol did not change microglial numbers, morphology, or ED1 expression in the dentate gyrus, possibly due to the high percentage of already activated microglia in the SEvehicle rats. Ablation of CX3CR1 receptors in mice leads to reduced microglial numbers during the first month of age during normal physiological condition (Paolicelli et al., 2011). However, blocking CX3CR1 signaling in the physiological environment with administration of an antibody (10 μg/day) has also been shown to enhance microglial numbers (Bachstetter et al., 2011). Similarly, in other brain pathological conditions, such as neurodegenerative disorders, ischemic brain injury or spinal nerve injury, modulating this pathway has given discordant outcomes with respect to microglial activation (Lee et al., 2010; Sheridan and Murphy, 2013; Wu et al., 2013; Tang et al., 2014). The mixed reports including both increased and decreased activation of microglia in response to inhibition of the fractalkine–CX3CR1 pathway in a pathological brain environment has already been suggested to depend on the severity and chronicity of the insult (Cardona et al., 2006). It has also been suggested that during physiological conditions, tonic activation of CX3CR1 by fractalkine helps to keep the microglia in a quiescent state. However, during an inflammatory condition, levels of fractalkine/CX3CR1 expression may increase and promote microglial migration and activation to the site of inflammation (Rogers et al., 2011; Sheridan and Murphy, 2013). Hence, another possibility for the discrepancy among different reports could be differences in the degree of inhibition, timing of treatment and dose–responses of treatment. Our results may support this argument since the low dose CX3CR1 antibody

treatment reduced microglial activation, while we observed a trend towards an increase in microglial activation with the higher dose (60 μg/ml). Despite a pronounced SE-induced increase in microglial numbers and morphological alterations, we did not observe a change in the hippocampal expression of some pro- and anti-inflammatory cytokines at one week post-SE, with or without the CX3CR1 antibody treatment. The initial increase in IL-6, within hours after SE, was only transient. This does not exclude alterations in cell-type or sub-regional specific expression within the hippocampus at any of the investigated time points after the insult (De Simoni et al., 2000; Ravizza et al., 2008). However, our data suggests that there is no prominent continuous cytokine release at one week post-SE. The anti-CX3CR1 antibody treatment diminished the SE-induced neurodegeneration in the dentate hilus. In agreement, the fractalkine– CX3CR1 pathway has been shown to regulate both microglial activation and microglia-induced neurotoxicity by modulating microglial mobilization, their binding to dying neurons and release of cytokines (Harrison et al., 1998; Fuhrmann et al., 2010; Fumagalli et al., 2013; Sheridan and Murphy, 2013; Brown and Neher, 2014). Accordingly, we observed a reduced percentage of activated microglial cells in the dentate hilus of CX3CR1 antibody-infused rats. This, along with an overall increased population of ramified and less intermediate forms of microglia in the dentate gyrus could mitigate the SE-induced excitotoxicity and cell death (Li et al., 2012). However, previous reports are rather complex and suggest even a protective effect of the fractalkine–CX3CR1 pathway in microglia/neuronal cell cultures in response to LPS and glutamate-mediated neurotoxicity (Zujovic et al., 2000; Deiva et al., 2004; Limatola et al., 2005). In an attempt to further evaluate the neuroprotective effect of inhibiting the CX3CR1 pathway, we studied the expression of excitatory and inhibitory synaptic scaffolding proteins, PSD-95 and gephyrin, which may suggest an imbalance in neuronal excitability as a contributor to neurodegeneration. PSD-95 and gephyrin are known to modulate excitatory and inhibitory synaptic

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transmission (Beique and Andrade, 2003; Prange et al., 2004; Varley et al., 2011) and the fractalkine–CX3CR1 pathway has been shown to regulate both glutamatergic excitatory and GABAergic inhibitory synaptic transmission (Lauro et al., 2008; Lauro et al., 2010; Piccinin et al., 2010; Scianni et al., 2013). In addition, in cortical brain tissues from TLE patients, modulating this pathway by adding recombinant fractalkine, decreased GABA current reduction on repeated GABA application, though, the recovery of currents to basal values was also affected, suggesting a possible desensitization of GABA receptors (Roseti et al., 2013). However, we did not find changes in the synaptic protein expression at one week post-SE, with or without CX3CR1 antibody treatment. Further studies are needed to evaluate whether more sub-regional changes in network excitability may explain the neuroprotection induced by the current anti-CX3CR1 treatment. Given that neuroblast production is normally increased acutely following seizures and the survival is compromised by microglial activation (Ekdahl et al., 2003), the reduced neuroblast production one week post-SE following CX3CR1 antibody treatment was unexpected. However, since both the number of proliferating cells and neuroblasts were reduced, it is likely that the CX3CR1 antibody treatment was primarily affecting the proliferation phase. When the pathological environment surrounding the neurogenic niche was reduced due to CX3CR1 antibody treatment, this could have resulted in less proliferation. Alternatively, the downstream effect of the CX3CR1 pathway on cell proliferation could be independent of its role in microglial activation. Partly supporting this later speculation is the fact that CX3CR1 receptor knockout mice show a reduced basal neurogenesis without a change in microglial cell numbers (Paolicelli et al., 2011; Vukovic et al., 2012; Fumagalli et al., 2013). Furthermore, a recent study has shown that CX3CR1 deficient mice do not display increased neurogenesis despite environmental enrichment (Reshef et al., 2014). However, young mice treated with anti-CX3CR1 antibody during normal physiology show reduced neurogenesis along with increased microglial cell numbers (Bachstetter et al., 2011). In our study, we also observed increased cell proliferation in the SGZ following fractalkine treatment post-SE. Because neuroblast production was not altered at this time point, it is not clear what cell type the increased proliferation pool represents. It may still result in neuroblast production at later time points and thereby be consistent with the finding from Bachstetter et al. (2011). They showed increased cell proliferation, without a change in neuroblast production or microglial activation during normal physiological conditions after one week of fractalkine treatment in aged rats, but an increase in both proliferation and neuroblast production after 2 weeks of administration. In summary, to further understand the diverse results presented so far following CX3CR1 modulation, manipulation of single downstream pathways of the CX3CR1 may help in defining its importance for microglial activation and a dependent or independent role in cell proliferation and neuroblast production. Whether the fractalkine–CX3CR1 pathway is also modulating later stages of neurogenesis, including differentiation and synaptic integration, remains to be investigated. In addition, future studies will show whether the alterations in seizure-induced pathology following CX3CR1 antibody treatment during early periods of epileptogenesis may eventually decrease the development of spontaneous seizures.

Conclusions This study shows an important role of the fractalkine–CX3CR1 signaling pathway in limbic seizure-induced hippocampal pathologies including microglial activation, neurodegeneration and neuroblast production. It also suggests that neuroblast production following seizures may partly occur as a result of microglial activation. Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.nbd.2014.11.009.

Acknowledgment We thank biomedical analyst Susanne Jonsson for technical support. This work was supported by the Swedish Research Council ( C0229701, C0229702), ALF Grant for funding for medical training and research, Zoega's Foundation, O.E. and Edla Johansson's Scientific Foundation, Tore Nilson's Foundation, and Åhlens' Foundation. The research leading to these results has also received funding from the European Union's Seventh Framework Programme (FP7/2007–2013) under grant agreement no. 602102 (EPITARGET).

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