FGF-2 Overexpression Increases Excitability and Seizure Susceptibility but Decreases Seizure-Induced Cell Loss

Animals.

TgFGF2 mice were used for all experiments. These mice harbor a transgene that includes the full-length human FGF-2 cDNA under the control of phosphoglycerokinase (pgk) 5′ regulatory elements (Coffin et al., 1995). These animals were created in the FVB/N strain, because of its suitability for transgenic manipulation (Taketo et al., 1991). The TgFGF2 mice express all four isoforms of human FGF-2 that arise from alternative initiation sites for translation (Florkiewicz and Sommer, 1989). TgFGF2 mice have been previously characterized from both molecular and phenotypic perspectives. Western blot analysis has shown that the murine and human FGF-2 isoforms are tissue-specifically expressed and translationally regulated (Coffin et al., 1995). By gross examination, the majority of mice seem to be indistinguishable from their WT littermates, but some of them are affected by skeletal malformations, such as shortening and flattening of long bones and moderate macrocephaly (Coffin et al., 1995).

Mating of TgFGF2 males with nontransgenic females generated 1:1 nontransgenic (or wildtype, WT) to TgFGF2 littermates that were used to generate experimental groups at 6–8 weeks of age. Presence of the FGF-2 transgene was confirmed by genomic PCR as described previously (Coffin et al., 1995). The genotype of each animal was assessed twice using PCR of genomic DNA isolated from tail (before experiment and after kill). The mice that were used for all experiments were 6–8 months old WT and TgFGF2 littermates. All animals were housed under standard conditions: constant temperature (22−24°C) and humidity (55–65%), 12 h dark-light cycle, free access to food and water. Procedures involving animals were conducted in accordance with European Community and national (Ministry of Health) laws and policies (authorizations: D.M. 82/2001-B and 93/2004-B).

In situ hybridization.

Mice were killed by decapitation under light diethyl-ether anesthesia, and their brains rapidly removed, frozen in isopentane cooled in a dry ice/methanol bath, and then stored at −70°C until use. Twenty μm coronal sections at the level of the dorsal hippocampus (plate 45; Paxinos and Franklin, 2001) were thaw-mounted onto polylisine coated slides, fixed in paraformaldehyde 4%, soaked in phosphate saline buffer (PBS 1×: PO₄²⁻ 0.01 m; NaCl 0.15 m) for 10 min at room temperature, then rinsed in a graded ethanol series and dried. Sections were stored at −70°C until use. Immediately before in situ hybridization, they were pretreated with proteinase K (10 μg/ml in Tris-EDTA buffer) for 10 min at 37°C and with acetic anhydride 0.25% v/v for 10 min at room temperature.

The FGF-2 riboprobe was obtained using the plasmid pBluescript-mFGF-2 (kindly provided by Dr. G.R. Martin, Department of Anatomy, University of California, San Francisco, CA, USA) containing a 475 bp portion of mouse FGF-2 cDNA. The human and mouse FGF-2 mRNAs in this region are 89.3% homologous (Huang and Miller, 1991). The plasmid was linearized with XbaI and transcribed with T3 RNA polymerase to obtain antisense riboprobes, or linearized with EcoRI and transcribed with T7 RNA polymerase to obtain sense riboprobes. Riboprobes were obtained by running the respective transcription assays in the presence of α[³³P]rUTP (PerkinElmer Life Sciences).

In situ hybridization was performed as previously described (Simonato et al., 1996, 1998). The in situ hybridization mixture contained 50% deionized formamide, 0.6 m NaCl, 2 mm EDTA, 20 mm Tris (2× NTE), 5× Denhart's solution, 100 μg/ml ssDNA, 100 μg/ml tRNA, 0.05% sodium pyrophosphate and 60 ng/ml ³³P-riboprobe. Sections were incubated overnight at 52°C with 40 μl of hybridization mixture. After hybridization, they were rinsed twice for 10 min in 4× SSC (0.6 m NaCl, 0.06 m citric acid) and treated with RNase A (20 μg/ml) for 30 min at 37°C. Sections were then rinsed in 1× SSC for 10 min, 0.1× SSC at 52°C for 30 min, 0.1× SSC at room temperature for 10 min, and dehydrated through an ethanol series. Autoradiograms were generated by apposing these dried sections alongside ³³P-riboprobe standards to Hyperfilm β-max (GE Healthcare Bio-Sciences) at −70°C for 4 weeks. Films were developed in Kodak D-19 (5 min) and fixed in Kodak fixer (5 min).

After development of the film, a digital analysis system (RBR Altair, Firenze, Italy) was used to obtain optical density measurements over structures of interest in the autoradiogram. The mean total optical density within a particular area was calculated by multiple sampling of that area in 4 sections taken from each animal. Background (nonspecific) hybridization was estimated using other sections incubated with the sense probe, and subtracted from the calculated values of total optical density. These specific optical density measurements were then compared with those obtained from the ³³P-riboprobe standards. By using this method of standardization, small differences in OD resulting from differences in film development were corrected, allowing values obtained from homologous regions to be compared across different films.

FGF-2 immunohistochemistry.

Mice were killed and their brains were rapidly removed, immersed in 10% formaline and then paraffin-embedded. Coronal sections (6 μm thick) were cut at the level of the dorsal hippocampus and mounted onto poly-lysine-coated slides.

The immunohistochemistry protocol was based on a previously described one (Marconi et al., 1999, 2005). Briefly, sections were deparaffinized (2 washes in xylol 10 min, 5 min in ethanol 100%, 5 min in ethanol 95%) and then rehydratated in distilled water for 5 min and in PBS 1× for 10 min. The FGF-2 antigen was unmasked using a commercially available kit (Unmasker Diapath), according to the manufacturer's instructions. After incubation in H₂O₂ 3V for 15 min at room temperature, sections were rapidly rinsed in distilled water and washed again in PBS. They were then incubated with Ultra V Block (Ultra Vision Detection System; Lab Vision Corporation) for 5 min at room temperature, to block nonspecific background staining. After washing in PBS 1× for 5 min, sections were incubated overnight at 4°C in humid atmosphere with the primary antibody for FGF-2 (mouse monoclonal; 5 μg/ml; BD Transduction Laboratories). After rinsing in PBS 1×, they were incubated with biotynilated goat anti-polyvalent serum (Ultra Vision Detection System; Lab Vision Corporation) at room temperature for 10 min, washed in PBS 1× for 5 min and then incubated in Streptavidine Peroxidase (Ultra Vision Detection System; Lab Vision Corporation) at room temperature for 10 min. The reaction product was detected as a brown substrate using a 10 mg tablet of 3,3-diaminobenzidinetetrahydrochloride (DAB; Sigma) in a solution containing 16.6 ml of PBS 1× and 130 μl of H₂O₂ 24V. Finally, sections were washed three times in PBS 1× (5 min each), counterstained with hematoxylin for 2 min and washed again in PBS 1× (5 min). Coverslips were mounted using Gel/mount (Biomeda). The specificity of immunolabeling was verified in all experiments by controls in which the primary antibody was omitted.

FGFR1 immunohistochemistry.

FGFR1 immunohistochemistry was performed using the method described for FGF-2. The primary antibody directed against FGFR1 (rabbit polyclonal; Santa Cruz Biotecnology) was diluted 1:100 in PBS and applied for 1 h at 37°C.

FGF-2 and GFAP double immunofluorescence.

WT and TgFGF2 mice were killed and their brains were rapidly removed, immersed in 10% formalin and then paraffin-embedded. Coronal sections (6 μm thick) were cut at the level of the dorsal hippocampus (plate 45, Paxinos and Franklin, 2001) and mounted onto polarized slides (Superfrost slides, Diapath).

Sections were deparaffinized, rehydratated, and the FGF-2 and GFAP antigens were unmasked as described for FGF-2 immunohistochemistry. Then, after a 1× PBS wash, they were preincubated with PBS containing 0.3% Triton X-100 for 10 min. After two rinses in PBS, they were blocked for 30 min in a solution of PBS containing 5% bovine serum albumin (BSA) and 5% normal goat serum. Finally, sections were incubated overnight at 4°C with the primary antibodies, anti-FGF-2 (mouse monoclonal; 10 μg/ml; BD Transduction Laboratories) and anti-GFAP (rabbit polyclonal; 1:100; Sigma).

The next day, sections were rinsed twice with PBS and preincubated with PBS containing 0.3% Triton X-100 for 30 min. Next, they were rinsed with PBS again and then incubated for 3 h at room temperature with a secondary antibody mixture containing goat anti-mouse, fluorescein isothiocyanate (FITC)-conjugate (1:100 dilution; Jackson ImmunoResearch) and a goat anti-rabbit, TX red-conjugate (1:100; Jackson ImmunoResearch). Finally, samples were washed three times with PBS, the last wash containing 100 ng/ml 4′, 6-diamidino-2-phenylindole (DAPI, Sigma). DAPI was used as a fluorescent counterstain for cell nuclei. Coverslips were mounted using anti-fading Gel/Mount water based media (Biomeda). The specificity of immunolabeling was verified in all experiments by controls in which the primary antibody was omitted.

VGLUT1 and VGAT immunohistochemistry.

On coronal, adjacent sections to those used for FGF-2 immunostaining, we performed VGLUT1 and VGAT immunohistochemistry, similar to the procedures described above for FGF-2. After pretreatment, tissue sections were incubated overnight in a humidity chamber at room temperature in PBS containing a primary antibody against either VGLUT1 (rabbit polyclonal; 1:500 dilution; Synaptic Systems) or VGAT (mouse monoclonal; 1:200 dilution; Synaptic Systems). DAB staining was performed as described above and coverslips were mounted using Gel/Mount (Biomeda). The specificity of immunolabeling was verified in all experiments by controls in which the primary antibody was omitted.

Extracellular recordings.

WT and TgFGF2 mice were anesthetized with ether and decapitated. Brains were quickly removed and transferred in cold oxygenated artificial CSF (aCSF; composition in mm: NaCl 124, KCl 2, KH₂PO₄ 1.25, NaHCO₃ 26, MgSO₄ 2, CaCl₂ 2, glucose 10; pH 7.4). Transverse dorsal hippocampal slices of 450 μm were cut with a vibroslicer (Campden Instruments Ltd.). After at least 1 h of incubation at room temperature for recovery, slices were placed in a recording chamber and continuously perfused with oxygenated aCSF at a flow rate of 2.5 ml/min. Experiments were performed at 30°C.

To probe for differential excitability in the two experimental groups, input-output (I/O) curves were generated. CA1 Schaffer collateral fibers were stimulated electrically (trains of 0.05–0.1 ms rectangular pulses at 0.033 Hz) through a twisted tungsten bipolar electrode, using a computer-driven stimulator unit (Digitimer DS2A, from AutoMate Scientific Inc.), and field excitatory postsynaptic potentials (fEPSPs) were recorded from CA1 stratum radiatum by means of an extracellular microelectrode filled with aCSF. The stimulating electrode was positioned 0.5–1 mm away from the recording electrode. Acquisition was done using an AC amplifier (A-M System 3000, A-M Systems) controlled by the LTP 2.30D software (Anderson and Collingridge, 2001). After several minutes of settling, evoked responses for a given stimulus were stable throughout the recording.

A series of test stimuli of increasing intensity were initially applied to identify the response threshold. The threshold stimulus intensity was always in the range of 3–10 V, and was not different in slices taken from WT or TgFGF2 mice (average threshold intensity in WT: 4.5 ± 0.9 V; in TgFGF2 5.0 ± 0.9 V). Intensity of input stimuli was increased by steps, starting at threshold (the maximal stimulus intensity that did not produce any detectable response) up to a maximal saturating value.

The slope of dendritic fEPSPs was calculated using the LTP 2.30D software and plotted versus increasing stimulus intensity. For each animal, the fold-increase in input stimulus intensity necessary for evoking a half-maximal response was calculated (based on the individual input-output curve) using the GraphPad Prism 4.0 software. These individual values were used for statistical analysis of the data that was conducted using the Student's t test for unpaired data.

Patch-clamp procedures.

Adult mice were deeply anesthetized (i.p. injection of 60 mg/kg sodium pentobarbital) and decapitated. The brain was exposed, chilled with aCSF, and the hippocampus was dissected. Transverse hippocampal slices (300–350 μm) were cut with a vibratome and stored at room temperature in a holding bath containing aCSF. After a recovery period of at least 1 h, slices were transferred to the recording chamber (1 cm³ volume) and mounted on an Olympus BX50WI microscope. Slices were constantly superfused using a gravity flow system (2 ml/min), with a aCSF having the following composition (mm): 125 NaCl, 2.5 KCl, 26 NaHCO₃, 1.25 NaH₂PO₄, 2 CaCl₂, 1 MgCl₂, and 15 glucose, continuously bubbled with 95% O₂/5% CO₂; the osmolarity was adjusted at 305 mOsm with glucose.

The pipette-filling solution contained the following (mm): 120 KCl, 10 NaCl, 2 MgCl₂, 0.5 CaCl₂, 5 EGTA, 10 HEPES, 2 Na-ATP, 10 glucose. The osmolarity was adjusted to 295 mOsm with glucose and the pH was set to 7.2 with NaOH. Membrane currents were recorded and acquired with an Axopatch 200A amplifier (Molecular Devices), and a 12 bit A/D-D/A converter (Digidata 1200B; Molecular Devices); off-line analysis was performed using version 10 of pClamp (Molecular Devices). A 70–80% compensation of the series resistance was routinely used.

Excitatory synaptic transmission was examined in CA1 pyramidal cells in both WT and TgFGF2 mice using the whole-cell patch-clamp technique in thin slices continuously perfused with bicuculline (10 μm). Schaffer collateral fibers were cut between CA3 and CA1 to avoid the onset of status epilepticus after perfusion with bicuculline. Bipolar twisted NiCr-insulated electrodes localized into the Schaffer collaterals bundle were used to evoke synaptic responses. We used minimal stimulation at the frequency of 0.05 Hz to activate only one presynaptic fiber connected to the neuron under recording. According to the technique described by Jonas et al. (1993) and Allen and Stevens (1994), the stimulation intensity was decreased until only a single axon connected to the recorded CA1 neuron was activated. This was achieved when the mean amplitude of the postsynaptic currents and the failure probability remained constant over a range of stimulus intensities near threshold for detecting a response (in the range of 2.5–6.5 mV). The latency and the shape of individual synaptic responses remained constant for repeated stimuli; in all recordings paired stimuli were applied at 50 ms intervals. Failures of transmission were judged subjectively for each trace. Data were statistically analyzed using the Origin 7.5 software (OriginLab), with the Student's t test.

Ca²⁺-dependent inactivation of NMDA currents.

Mice were decapitated under deep anesthesia and their brains were quickly removed and placed in a cold carboxygenated solution with the following composition (in mm): KCl 3, NaH₂PO₄ 1.25, NaHCO₃ 21, MgCl₂ 2, CaCl₂ 1.6, glucose10; sucrose 200, α-tocopherol 0.1; pH 7.4, osmolarity 300–305 mOsm. The Na⁺ concentration was lowered to prevent hypoxia-induced Na⁺ influx into neurons and α-tocopherol was added as a scavenger of free radicals (Gabriel et al., 2004). Transverse slices (300–350 μm) were cut by using a tissue vibratome and then transferred to an interface tissue chamber and maintained in a bicarbonate buffer solution (BBS) bubbled with 95% O₂ and 5% CO₂ at room temperature at least 1 h before their use. The BBS composition was (in mm): NaCl 125, KCl 2.5, NaHCO₃ 26, NaH₂PO₄ 1.25, MgCl₂ 1, CaCl₂ 2, glucose 15.

In conventional whole-cell patch-clamp recordings, some of the endogenous Ca²⁺ buffer systems and of other intracellular components that regulate NMDA receptor function may be washed out. To avoid this problem, we used perforated patch recordings to study the calcium-dependent inactivation of NMDA receptors. This configuration, maintaining a relatively intact intracellular environment, has been shown to ensure optimal results in the study of calcium-dependent inactivation (Kyrozis et al., 1996). The antibiotic amphotericin B was used as perforating agent (Horn and Marty, 1988). Before each experiment, 2 mg of amphotericin B were dissolved in 50 μl of dimethylsulphoxide (DMSO). This solution was sonicated for 2 min and then added to the pipette solution.

NMDA (50 μm) was focally applied to the cell body of neurons via a glass tube with an internal diameter of 0.8 mm, positioned at ∼2 mm distance from the cell (Biologic RSC-160, Claix, France). Membrane currents were recorded and acquired with an Axopatch 200A amplifier (Axon Instruments) and a 12 bit A/D-D/A converter (Digidata 1440A; Axon Instruments). Off line analysis was performed using pClamp version 10.1 (Axon Instruments).

Kainate.

Kainate was administered i.p. (20 mg/kg), and the mouse behavior was observed for 2 h thereafter. The occurrence of seizures and their severity were blind-scored by two investigators, using the following classification (Janumpalli et al., 1998; Bregola et al., 2002): 1, chewing and drooling; 2, head nodding; 3, unilateral forelimb clonus; 4, bilateral forelimb clonus; 5, bilateral forelimb and/or hindlimb clonus with falling; 6, running or jumping seizure; 7, tonic hindlimb extension; 8, death.

Bromodeoxyuridine in vivo labeling and immunohistochemistry.

For bromodeoxyuridine (BrdU) immunohistochemistry, WT and TgFGF2 mice were divided in two groups after kainate administration. The first group was used to study proliferation of neural stem cells and progenitors. Three days after kainate administration, mice were given a series of four BrdU injections (50 mg/kg i.p., dissolved in propilenic glycol and water, Boehringer Mannheim), every 2 h over a period of 6 h. These animals were killed 24 h after the last BrdU injection. The second group was used to study neurogenesis. For 5 d, beginning 3 d after kainate, WT and TgFGF2 mice in this group received a single BrdU injection per day (50 mg/kg i.p.). These animals were killed 9 d after kainate.

BrdU immunostaining was performed as previously described (Marconi et al., 2005). Briefly, DNA was denatured by incubating tissue sections in 2 N HCl for 30 min at 37°C. Sections were rinsed in PBS 1× and treated with 3% H₂O₂ 3V to block endogenous peroxidase. Slices were then washed in PBS 1× and incubated with Ultra V Block. Incubation with a primary antibody for BrdU (mouse monoclonal; 1:100 dilution in PBS 1×; Boehringer Mannheim) was conducted overnight at room temperature in a humidity chamber. After 3× 5 min rinses in PBS, they were incubated with biotinylated secondary antibody and then in Streptavidine Peroxidase. The reaction product was detected using DAB, as described above. Finally, sections were counterstained with hematoxylin and the coverslips mounted using Gel/mount. The specificity of immunolabeling was verified in all experiments by controls in which the primary antibody was omitted.

Quantification of BrdU immunohistochemistry experiments was performed in a blinded manner by two investigators, counting the number of BrdU-positive cells in the granular layer and in the hilus of the dorsal hippocampus dentate gyrus. Paraffin-embedded brains were cut in successive 6 μm sections across the entire dorsal hippocampus (i.e., 141 sections, corresponding to 846 μm; plates 42–49, Paxinos and Franklin, 2001). One section every 28 (that is, every 168 μm) was used for BrdU immunohistochemistry. Five sections have been examined per each animal (progressive section numbers: 1, 29, 57, 85, 113). An estimate of the total BrdU-positive cells in the area under examination was obtained by multiplying the number of BrdU-positive cells in each section by 28 and summing the five resulting counts. Statistical analysis has been performed using ANOVA and post hoc the Newman–Keuls test.

Activin A immunohistochemistry.

Activin A (ActA) immunohistochemistry was performed using the method described for FGF-2. The primary antibody directed against ActA (rabbit polyclonal; dilution 1:50; R&D Systems) was applied overnight in a humidity chamber at 4°C. The specificity of immunolabeling was verified by controls in which the primary antibody was omitted.

BrdU and doublecortin double immunofluorescence.

BrdU and doublecortin (DCX) double immunofluorescence was performed as described above for FGF-2 and GFAP immunofluorescence. Sections were incubated overnight at 4°C with the primary antibodies, anti-BrdU (rat polyclonal; dilution 1:50; AbD Serotec) and anti-DCX (goat polyclonal; dilution 1:50; Santa Cruz Biotechnology).

The next day, they were incubated for 3 h at room temperature with a secondary antibody mixture containing donkey anti-goat, FITC-conjugate (dilution 1:25; Jackson ImmunoResearch) and a donkey anti-rat, TX red-conjugate (1:50; Jackson ImmunoResearch). DAPI was used as a fluorescent counterstain for nuclei. The specificity of immunolabeling was verified by controls in which the primary antibody was omitted. The quantification of the number of DCX-positive cells in the dentate gyrus was performed as described above for BrdU. It must be noted that the use of very thin (6 μm), paraffin-embedded sections offers the advantages of very high resolution, of clear identification of stained cells (in that no overlap of multiple cell layers can occur), and of precise anatomical localization (thanks to the possibility of counterstaining the sections with standard procedures, such as DAPI).

Analysis of cell damage.

Analysis of seizure-induced cell damage was performed in WT and TgFGF2 mice killed 3 d after kainate administration, by using Fluoro-Jade B (FJB) staining and silver impregnation. As described above for BrdU, paraffin-embedded brains were cut in successive 6 μm sections across the entire dorsal hippocampus (142 sections, corresponding to 852 μm; plates 42–49, Paxinos and Franklin, 2001). Two adjacent sections every 35 were used for FJB staining and silver impregnation. FJB staining was performed as described (Schmued and Hopkins, 2000; Zucchini et al., 2002). Briefly, sections were mounted onto poly-lysine coated slides, deparaffinized (as described above) and rehydrated in a solution containing 1% NaOH in 80% ethanol (5 min), then 70% ethanol (2 min), and then distilled water (2 min). Subsequently, they were incubated for 10 min in 0.06% potassium permanganate, washed for 2 min in distilled water and transferred to a 0.001% FJB staining solution. After staining, sections were washed 3× 1 min in distilled water, dried, immersed for 1 min in xylene and then coverslipped.

Silver staining impregnation has been performed according to the method of Gallyas et al. (1980). Sections were deparaffinized and immersed in a pretreating solution (0.5% ammonium nitrate in 4.5% NaOH) to prepare the tissue for reaction with silver nitrate solution (0.75% silver nitrate in 8% ammonium nitrate and 4.5% NaOH). Neurodegenerated cells were identified by inclusion of silver in the nuclei. To avoid aspecific precipitation of silver nuclei, washes of 3× 5 min were performed for eliminating the excess of silver nitrate accumulated on sections. Slices were then dipped in developing solution (0.6% formaldehyde, 0.05% citric acid, 10% ethanol and 0.001% ammonium nitrate) for 1 min, to increase the size of the silver nuclei, which become visible at light microscopy. Finally, sections were soaked in a 0.5% acetic acid solution 3× 10 min.

Analysis was conducted using a Leica microscope (DMRA2, Leica). The degree of cell damage was quantified blinded by two investigators, based on two distinct approaches. The first was a modification of a previously described scoring system named “neurodegeneration score” (Zucchini et al., 2002; Franceschetti et al., 2007). The score range spanned from 0 (absence of FJB- or silver impregnation-positive neurons) to 5 (maximal neurodegeneration pattern in the hippocampus). A score of 1 was given for observation of positive cells either in the hilus, in the CA3 pyramidal layer, or in the CA1 pyramidal layer. Whenever a very intense neurodegeneration was observed in the CA3 or CA1 pyramidal layer (many positive cells with confluent fluorescence), the score related to that subarea was increased by 0.5; another 0.5 was added when fluorescent cells were detectable in the strata oriens and/or radiatum (therefore, the score related to CA3 and to CA1 could reach a maximal value of 2). The second quantification method was based on the thresholding of FJB digital images. Images of the hippocampus were captured using a Leica DFC300FX camera and transformed into gray levels. Using Adobe Systems Photoshop CS2 (version 9.0.2), the hippocampus was cut out and the mean ± SD gray level was calculated. FJB-positive pixels were identified by thresholding at the gray level corresponding to the mean plus three SDs. Using this approach, only those pixels that were significantly above background (that is, positive to FJB) were selected. Data were then expressed as percentage of FJB-positive pixels over total hippocampal pixels.

As stated above, two every 35 of the 142 6 μm sections cut across the dorsal hippocampus were stained (one with FJB, the other with silver impregnation). Thus, five FJB (progressive section numbers: 1, 36, 71, 106, 141) and five silver impregnation sections (progressive numbers: 2, 37, 72, 107, 142) have been examined for each animal. The mean neurodegeneration scores in the five sections examined were obtained for each staining approach (FJB and silver impregnation). Finally, these two neurodegeneration scores were averaged, obtaining a single value per animal that was used for statistical analysis. Alternatively, the mean percentage of FJB-positive pixels in the five sections examined was calculated for each animal and used for statistical analysis. Statistical analysis was conducted using the Mann–Whitney two-tailed U test.

Cleaved caspase-3 immunohistochemistry.

Cleaved caspase-3 immunohistochemistry was performed using a method very similar to the one described above, for FGF-2 immunostaining. The essential difference was that the primary antibody was directed against active caspase-3 (rabbit polyclonal; Cell Signaling Inc.) instead of FGF-2. Anti-capsase-3 was diluted 1:100 in PBS and applied overnight at 4°C. Subsequent application of secondary antibody, processing for microscopy and analysis of the samples was identical to procedures described above.

Timm staining.

Thirty days after kainate treatment, a group of WT and TgFGF2 mice was trans-cardially perfused with 4% paraformaldehyde and 1.25% glutaraldehyde in phosphate buffer. After postfixation (0.4% Na₂S for 30 min, 1% paraformaldehyde and 1.25% glutaraldehyde overnight), 20 μm coronal brain sections were cut at the level of the dorsal hippocampus and Timm staining was performed as described by Wenzel et al. (1997), to reveal axonal sprouting of the mossy fibers. Briefly, free-floating sections were put in a developer [30 ml of 25% arabic gum, 5 ml of 2 m citrate buffer, pH 3.7, 15 ml of 5.67% hydroquinone and 0.25 ml of 17% AgNO₃] for at least 60 min in the dark, under constant agitation. Sections were then washed in distilled water for 5 min and dehydrated with alcohol and xylene.

Analysis was performed using a digital analysis system (RBR Altair, Firenze, Italy). The areas of interest were divided in circular areas according to procedures described by Adams et al. (2002) and a score (from 0 to 5, according to Cavazos et al., 1991) was given to every circle. The “Timm index” was the sum of these scores.