Role of the 5-HT4 receptor in chronic fluoxetine treatment-induced neurogenic activity and granule cell dematuration in the dentate gyrus

Animals

All mice were housed under standard illumination parameters with a 12-h light/dark
cycle and ad libitum access to water and food. Male 5-HT₄ receptor heterozygous mutant mice that had been backcrossed to the C57BL/6 J background
for 10 generations were purchased from the Jackson Laboratory (Bar Harbor, ME, USA).
Male (6–12 weeks old) wild-type (WT) and homozygous mutant (5HT4R KO) mice prepared
by heterozygous mating were used as previously described 10],34]. For the cranial irradiation experiments, C57BL/6 N mice (5 weeks old) were purchased
from the Japan SLC (Hamamatsu, Japan) and habituated for over one week before experimental
procedures. All experimental procedures were approved by the Committee of Animal Research
of Kyoto University Faculty of Pharmaceutical Sciences.

Cranial irradiation

Mice (6 weeks of age) were anesthetized by pentobarbital (50 mg/kg; Kyoritsu Pharma),
and exposed to cranial irradiation using a Rigaku Radiofrex 350 X-ray generator operated
at 250 kV and 15 mA with a 1-mm-thick aluminum filter. X-rays at a dose of 10 Gy were
delivered at a dose rate of 0.74 Gy/min. A lead shield was placed over the body of
the mice except the head. Non-irradiated controls received anesthesia only. The fluoxetine
treatment was started 14 days after irradiation.

SSRI treatment

Fluoxetine hydrochloride (LKT laboratories, Inc., St. Paul, MN, USA) was intraperitoneally
injected for 21 days at a dose of 22 mg/kg. The fluoxetine solution was prepared every
day. Control mice were given saline. For the comparison analysis of gene expression
in the DG (Figure 4D), 11 out of 21 mice were orally given fluoxetine hydrochloride dissolved in 0.2%
of saccharin solution, which was applied for 28 days at a dose of 22 mg/kg. The concentration
of fluoxetine was determined for individual mice based on the amount of liquid consumption
quantified during the preceding 24 h period and the body weight 10]. Control mice were given water (orally).

Measurement of tissue monoamine contents and in vivo microdialysis

Measurement of tissue 5-HT and 5-HIAA contents were performed as previously described
with some modifications 35]. Briefly, 1-mm-thick coronal slices were prepared using a brain matrix on ice. Then,
tissue blocks containing the dorsal raphe nucleus and median raphe nucleus or hippocampus
were dissected, homogenized, and sonicated in 300 ?L of ice-cold 0.1 M HClO₄ containing 10 mM Na₂S₂O₅ and 1 mM EDTA. Protein concentrations were measured by using the Bradford protein
assay (Bio-Rad, Hercules, CA, USA). Homogenates were centrifuged at 16,000?×?g for 15 min at 4°C, and supernatants were stored at ?80°C until use. Supernatants
were thawed on ice and analyzed by high-performance liquid chromatography with an
electrochemical detector (HPLC-ECD) (Eicom, Kyoto, Japan). The measured 5-HT concentration
was normalized against the total protein concentration. The detection limits for both
5-HT and 5-HIAA were estimated to be approximately 0.5–0.6 fmol per 25 ?L of sample.

For in vivo microdialysis, mice were stereotactically implanted with a guide cannula (Eicom)
in the ventral hippocampus according to the atlas of Franklin and Paxinos 36] with the following coordinates: anterior-posterior, ?2.8 mm from the bregma; lateral,
3.0 mm from the mid line; dorsal-ventral, ?2.5 mm from the skull surface. Experiments
were performed 1 day after surgery in awake and freely moving mice. On the day of
the experiment, a dialysis probe with a length of 1 mm (Eicom) was inserted into the
guide cannula and perfused with Ringer’s solution (147 mM NaCl, 4 mM KCl, 3 mM CaCl₂) at a flow rate of 1 ?L/min for 2–3 h. After the initial perfusion period, dialysate
samples were collected every 20 min. On-line quantification of 5-HT in the dialysate
was accomplished by HPLC-ECD, and seven basal samples were collected before administration.
Fluoxetine (22 mg/kg) or saline was then administered IP and samples were collected
for an additional 120 min. The average values of the seven basal samples for each
animal were defined as 100% and used for normalization. Probe placement in each mouse
was histologically verified by examining the coronal brain sections after completion
of the experiment.

Immunohistochemistry

On the day following the last treatment of fluoxetine, mice intraperitoneally received
BrdU (150 mg/kg, Sigma B5002). Two hours after BrdU injection, the mice were anesthetized
by chloral hydrate (Nacalai Tesque, Kyoto, Japan) and transcardially perfused with
cold saline followed by 4% paraformaldehyde (PFA) in 0.1 M phosphate buffer, pH 7.4.
Brains were postfixed with 4% PFA overnight at 4°C, cryoprotected in 20% sucrose overnight,
and stored at ?80°C. Serial sections were then cut through the entire hippocampus
36] at a thickness of 30 ?m with a cryostat (Leica CM3050) and stored in a non-freezing
solution at ?20°C until stained.

For BrdU immunostaining, every sixth section was mounted on glass slides and incubated
in 0.01 M citric acid at 90°C for 5 min, denatured with 2 M hydrogen chloride at 37°C
for 30 min, neutralized with 0.1 M boric acid (pH 8.5) at room temperature for 10 min,
blocked in 10% equine serum in PBS containing 0.3% Triton X-100 at room temperature
for 60 min, and incubated with monoclonal rat anti-BrdU (1:200; Serotec OBT0030) at
room temperature overnight. For DCX and calbindin immunostaining, sections were blocked
in 10% equine serum and incubated with polyclonal goat anti-DCX (1:500; Santa Cruz
SC8066) or monoclonal mouse anti-calbindin D-28 K (1:3000; Sigma Aldrich, C9848).
Sections were then incubated for 60 min with biotinylated goat anti-rat IgG (1:1000;
Vector BA9400) as the anti-BrdU secondary antibody or biotinylated horse anti-goat
IgG (1:1000; Vector BA9500) as the anti-DCX secondary antibody, or biotinylated horse
anti-mouse IgG (1:1000; Vector BA2000) as the calbindin D-28 K secondary antibody,
followed by incubation with ABC Vectastain Kit (Vector). Antigen detection was performed
with 0.06% 3,3?-diaminobenzidine (DAB) staining and counterstaining with Nuclear Fast
Red (Vector).

For immunofluorescence staining, sections were blocked in 10% equine serum and incubated
overnight at 4°C with polyclonal rabbit anti-?-galactosidase (1:2000, MP, 55976) and
either polyclonal goat anti-DCX (1:500; Santa Cruz SC8066), monoclonal mouse anti-calretinin
(1:3000; Millipore, MAB 1568), monoclonal mouse anti-NeuN (1:500; Millipore, MAB377),
or monoclonal mouse anti-calbindin D-28 K (1:3000; Sigma Aldrich, C9848). Sections
were then incubated for 60 min with a secondary antibody conjugated with AlexaFluor
488 or AlexaFluor 555 (Molecular Probes).

Quantification of BrdU-labeled cells, DCX-positive cells, and calbindin-IR

For BrdU-labeled cell quantification, a modified unbiased stereological procedure
was used as previously described 37]. Sections were coded to ensure that the analysis was performed by a blind observer
using a light microscope (Nikon Ecripse E200, Tokyo, Japan), and BrdU-labeled cells
in the SGZ of the hippocampus were counted. Cells were included in SGZ counts if they
were touching the border between the GCL and the hilus, or in the deepest layer of
the GCL. If a cell was more than two cell diameters from the GCL, it was excluded.
Every sixth 30-?m-thick section was evaluated throughout the hippocampus, and the
sum of the cell counts was multiplied by six to provide an estimate of the total number
of BrdU-labeled cells in the entire region. For DCX-positive cell quantification,
3–4 sections of the DG were photographed using a Biozero BZ-8000 microscope (Keyence
Corporation, Osaka, Japan) that was fitted with a 20X objective. The boundaries of
the DG were set as regions of interest (ROI), and each ROI was measured; then, DCX-positive
cells were manually counted within the ROI and their number was expressed as the number
of cells per square millimeter. For quantification of calbindin-IR, 2 sections of
the DG were photographed in the manner described above. The pictures were converted
into 16-bit gray scale pictures and the average signal intensity of calbindin-IR in
the GCL region or ML region was quantified by computer-assisted image analysis (ImageJ,
NIH, Bethesda, Maryland).

In situ hybridization

In situ hybridization was performed using a digoxigenin (DIG)-labeled riboprobe as previously
described 38]. A Ht4r cDNA template probe was cloned by PCR with gene-specific primers (Table 1), verified by sequencing, and used to produce digoxigenin (DIG)-labeled riboprobes
with the DIG RNA Labeling Kit (Roche, Mannheim, Germany). Coronal brain sections (thickness,
10 ?m) were cut on a cryostat, mounted onto slides, fixed in 4% PFA, acetylated, and
dehydrated prior to hybridization. Sections were hybridized with the DIG-labeled riboprobe.
DIG was visualized using an alkaline phosphatase-conjugated anti-DIG antibody and
BM purple substrate (Roche).

Table 1. List of primers used for quantitative real-time PCR analysis and for the5ht4rcDNA probe

Quantitative RT-PCR

Total RNA was extracted from the DG using the ReliaPrep RNA Miniprep System (Promega),
and subjected to reverse transcription with Superscript VILO (Invitrogen), and followed
by real-time PCR on a LightCycler (Roche) using the Fast Start DNA Master SYBR Green
I (Roche). Expression levels of target genes were normalized to the levels of 18S
rRNA. Primer sequences are shown in Table 1. The specificity of each primer set was confirmed by checking the product size by
gel electrophoresis.

Statistical analyses

All data are presented as the mean?±?S.E.M., and experiments with 2 groups were compared
using unpaired Student’s t-test, whereas experiments with 4 groups were subjected to two-way ANOVA, followed
by the Bonferroni post hoc test. A two-way repeated measure ANOVA was used for the microdialysis assay. Significance
marks in figures are based on results from the t-test, two-way ANOVA test, and Bonferroni post hoc tests. The threshold for statistical significance was P??0.05. All analyses were
performed using PRISM 5 software (GraphPad, San Diego, CA).