Sprague-Dawley (SD) male rats, 5 weeks of age, and weighing 150-170 g (Japan SLC, Shizuoka, Japan), were used in this study. They were housed in groups of six rats per cage in a room maintained under standardized 12 hours light/dark cycle (lights on at 07:00) and temperature (22±3℃). Rats had free access to food pellets and tap water. Body weights were recorded every 2 days in the morning after being taken out of restraining cages, for the duration of the experimental period.

Restraint stress was employed and involved enclosing each rat in a flexible wire mesh (5×5 mm) shaped to fit its body. Restrained rats were able to move a little and had no access to food or water during the restraint; control rats also had no access to food or water during this time. Because rats are nocturnal, they were restrained from 19:00 to 07:00 (12 hours) every 24 hours for the 22-day experimental period. After release from the restrainer, rats were returned to their home cages each morning and had access to food and water ad libitum, as did the controls.30

The major salivary glands were removed bilaterally (sialoadenectomy), or a sham operation was performed, in 3-week-old male rats under sodium pentobarbital anesthesia (65 mg/kg, i.p.).9 A 2-cm skin incision in the neck was made, the major salivary glands were isolated from the surrounding subcutaneous fat tissue, a ligature was placed around the supplying vessels, and the salivary glands were then removed bilaterally. Two weeks after surgery, the repeated restraint stress and handling procedure began. All experiments were performed with 12 rats per experimental group. At the end of the experiment, all rats were deeply anaesthetized with sodium pentobarbital (65 mg/kg, i.p.) for blood and tissue collection. To minimize diurnal variations in NT-3 expression, all rats were sacrificed between 07:00 and 11:00; the stressed rats were anaesthetized immediately following the last restraint session. The experimental protocol used in this study was reviewed and approved by the Ethics Committee on Animal Experiments of Kanagawa Dental College, and was carried out with adherence to the Guidelines for Animal Experimentation of Kanagawa Dental College.

At the end of the experiment on the 23rd day, all rats were deeply anaesthetized with sodium pentobarbital as above, and blood samples were collected between 07:00 and 11:00 by cardiac puncture into Venoject® tubes containing EDTA (Terumo, Tokyo, Japan). The tubes were immediately placed on ice and then centrifuged (2000 rpm, 15 minutes, 4℃). Plasma was stored at -20℃ prior to radioimmunoassay.

After blood sampling, the anaesthetized intact rats were sacrificed by decapitation, and submandibular glands were removed. Total RNA isolation from the submandibular glands was performed using the ISOGEN reagent (Nippon Gene, Toyama, Japan) in accordance with the manufacturer's instructions. The RNA was resuspended in 20 µL of diethyl pyrocarbonate treated water. RNA quality was judged from the pattern of ribosomal RNA after electrophoresis through a 1.5% agarose gel containing ethidium bromide and visualization by UV illumination. RNA concentrations were determined by absorbance readings at 260 nm with a SmartSpec Plus spectrophotometer (Bio-Rad, Tokyo, Japan). RNA was stored at -80℃ until use. Total RNA was reverse transcribed at 50℃ for 30 minutes, 99℃ for 5 minutes, and 5℃ for 5 minutes using a single-strand cDNA synthesis kit (Roche Diagnostics Ltd., Lewes, UK) according to the manufacturer's instructions. Following the reverse transcription reaction, the cDNA was stored at -20℃ until use.

Real-time polymerase chain reactions (PCR) was performed using a LightCycler (Roche) according to the manufacturer's instructions. Reactions were performed in a 20 µL volume (NT-3: 0.5 µM each primer and 3 mM MgCl₂). Reactions with Taq DNA polymerase, nucleotides, and buffer for NT-3 were performed with LightCycler-DNA Master HybProbe mix (Roche). Oligonucleotide primers were designed to amplify rat NT-3. The NT-3-specific primers were 5'-TGTGGGTAGCCGACAAGTC-3' (forward) and 5'-GAGTTCCAGTGTTTGTCATC-3' (reverse; NT-3 PCR product: 175 bp) as designed and synthesized by Nippon Gene Laboratory.31 Real-time PCR for amplification of the rat β-actin housekeeping gene was performed using a Lightcycler Primer/Probe set, 5'-CCTGTATGCCTCTGGTCGTA-3' (forward) and 5'-CCATCTCTTGCTCGAAGTCT-3' (reverse; β-actin PCR product: 260 bp),22,23 following the manufacturer's instructions (Nihon Gene Research Labs Inc., Sendai, Japan). Denaturation was performed at 95℃ for 10 minutes, after which Segment 1 (95℃ for 10 seconds), Segment 2 (60℃ for 10 seconds), and Segment 3 (72℃ for 10 seconds) were repeated for 40 cycles. We performed melting analysis and agarose gel electrophoresis to confirm the specificity of the PCR products obtained using each primer pair. Gene expression is stated in terms of the copy number ratio of NT-3 mRNA to β-actin mRNA for each sample.

The rats were sacrificed under deep anesthesia as above, between 07:00 and 11:00; rats in the chronic stress groups were sacrificed immediately after the last restraint session. Resected rat submandibular gland tissue samples were fixed in 10% buffered formaldehyde (pH 7.4) for 24 hours and embedded in paraffin. Next, serial 3-µm sections were cut and stained with hematoxylin and eosin and processed for immunohistochemistry. Immunohistochemical analysis was performed using the simple stain MAX-PO Kit (Nichirei, Tokyo, Japan). Slides were pre-incubated in 3% H₂O₂ for 5 minutes. Sections were then incubated with anti-NT-3 rabbit polyclonal antibody (sc-547, 1 : 500, Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA) for 1 hour at room temperature. After washing with PBS, sections were reacted with a secondary antibody, horseradish peroxidase (HRP)-labeled anti-rabbit IgG with amino acid polymer (Nichirei), for 30 minutes at room temperature. Color was developed using 0.02% 3,3'-diaminobenzidine-tetrahydrochloride containing 0.0003% H₂O₂ in Tris-buffered saline for 5 minutes, and sections were then counterstained with hematoxylin. To provide negative controls, non-immunized rabbit or mouse IgG was used instead of the primary antibody. To determine the binding specificity, a competitive assay was also carried out using recombinant NT-3 (R&D Systems, Inc., Minneapolis, MN, USA).

Adrenocorticotropic hormone (ACTH) was assayed using a radioimmunoassay (RIA) kit [enzyme-linked immunosorbent assay (ELISA)-ACTH, CIS; Atomic Energy Laboratory of Biochemical Products, Gif-sur-Yvette, France], following the manufacturer's instructions. ACTH concentrations are reported in pg/mL. A COAT-A-COUNT Rat Corticosterone kit (Siemens Medical Solutions Diagnostics, New York, NY, USA) was used for measurement of plasma corticosterone concentration (ng/mL).

Submandibular gland tissue samples were homogenized in ice-cold lysis buffer, containing 137 mM NaCl, 20 mM Tris-HCl (pH 8.0), 1% NP40, 10% glycerol, 1 mM phenylmethylsulfonyl fluoride, 10 µg/mL aprotinin, 1 µg/mL leupetin, and 0.5 mM sodium vanadate. The tissue homogenates were centrifuged at 14000 g for 5 minutes at 4℃. The supernatants were collected and used for quantification of total protein and NT levels. Total protein concentrations were determined by the Bradford method using absorbance readings at 595 nm with a SmartSpec Plus spectrophotometer (Bio-Rad).

NT-3 concentrations in the submandibular gland and plasma were assayed using an ELISA kit (Promega, Co., Madison, WI, USA). Briefly, standard 96-well flat-bottom NUNC-immunoMaxisorp ELISA plates were incubated with the corresponding captured antibody, which binds the NT of interest, overnight at 4℃. The plates were blocked by incubation for 1 hour at room temperature with a 1×block and sample buffer. Serial dilutions of a known amount of NT-3, ranging from 0 to 300 pg/mL, were performed in duplicate for standard curve determination. Wells containing the standard curve samples and supernatants of submandibular gland tissue homogenates and plasma were incubated at room temperature for 6 or 2 hours, as specified by the protocol. They were then incubated with a second specific antibody overnight at 4℃ or for 2 hours at room temperature. A species-specific antibody conjugated to horseradish peroxidase was used for a tertiary reaction for 2.5 or 1 hour at room temperature following this incubation step. TMB One Solution was used to develop color in the wells. This reaction was terminated with 1M hydrochloric acid after a specific time (10-15 minutes) at room temperature, and absorbance was then recorded at 450 nm in a plate reader within 30 minutes of stopping the reaction. NT-3 values were determined by comparison with the regression line for NT-3 standard. Using these kits, NT-3 can be quantified over a range of 0-300 pg/mL. For the NT-3 assay kit, cross-reactivity with other neurotrophic proteins was <2-3%.

Statistical analyses were carried out using the SPSS (Version 17.0; SPSS Inc., Chicago, IL, USA) statistics program. The Student's t-test and one-way ANOVA followed by Tukey's post-test were used for statistical analysis. A probability level of 0.05 or less was accepted as significant.

Prior to the experimental period, all rats had similar body weights. Because the rats were in their active growth phase, their body weight increased during the experiment. No significant differences in the mean body weight between the control and chronic stress groups were observed throughout the duration of the experimental period (data not shown).

Plasma ACTH levels were significantly elevated in the chronic stress group, with values of 78.24±3.1 pg/mL and 374.67±61.11 pg/mL observed in the control and chronic stress groups, respectively. Similarly, there were also significant differences in ACTH levels between the sham-control group and the sham-chronic stress group. Chronic stress also significantly elevated plasma corticosterone levels from 39.22±5.72 ng/mL to 530.97±49.8 ng/mL in the control and chronic stress groups, respectively. Similarly, significantly elevated corticosterone levels were observed in the sham-chronic stress group compared to the sham-control group.

Melting curve analysis demonstrated a single fluorescent peak representing the Tm of NT-3 mRNA in all samples, except for the negative sample (data not shown). In addition, a single band was observed following agarose gel electrophoresis (data not shown). These findings confirmed that the PCR product was NT-3 mRNA. Expression ratios (NT-3 mRNA/β-actin mRNA) were calculated in submandibular gland of the chronic stress and control groups. The expression ratios in the submandibular gland were 0.0028±0.0017 and 0.0084±0.00467 in the control and chronic stress groups, respectively (Fig. 1). Chronic stress resulted in a significant increase in NT-3 mRNA expression in the submandibular gland (Fig. 1). Similarly, submandibular gland NT-3 mRNA expression was significantly elevated in the sham-chronic stress group as compared to the sham-control group.

Exposure to chronic stress significantly increased submandibular gland NT-3 levels from 369.22±49.64 pg/mL to 557.60±244.17 pg/mL (Fig. 2), with a similar, significant increase observed in the sham groups.

Plasma NT-3 levels were significantly elevated as the result of chronic stress, increasing from 11.67±6.98 pg/mL to 406.00±322.25 pg/mL (Fig. 3). Similarly, there was also a significant increase in plasma NT-3 levels between the sham-chronic stress and sham-control group.

Rat pancreas sections were used as a positive control for NT-3 immunohistochemistry,19 and showed high levels of NT-3 expression in the islets of Langerhans (data not shown). Non-stressed submandibular gland tissue exhibited a weak positive signal for NT-3 expression in various duct-type cells (Fig. 4A). In comparison, intense NT-3 expression was observed in various duct-type cells of the chronic-stress group; however, NT-3 expression was not consistently observed in acinar cells or myoepithelial cells in the chronic stress group (Fig. 4B). NT-3 expression was not detected in the competitive assay (data not shown).

In sialoadenectomized rats, ACTH levels were 80.02±4.26 pg/mL and 430.00±44.71 pg/mL in the control and chronic stress groups, respectively, representing a significant increase in ACTH levels following chronic stress (Table 1). Similarly, there were also significant differences in ACTH levels between the sham-control group and sham-chronic stress group. Chronic stress also significantly increased corticosterone levels, from 35.76±4.91 ng/mL to 466.42±43.14 ng/mL, in sialoadenectomized rats (Table 1). Likewise, significantly elevated corticosterone levels were observed in the sham-chronic stress group compared to the sham-control group.

The plasma NT-3 levels were 406.00±332.25 pg/mL for non-sialoadenectomized group, 217.50±187.66 pg/mL for sialoadenectomized group, and 292.60±158.30 pg/mL for sham-chronic stress group under the chronic stress condition. There were no significant differences in plasma NT-3 levels among the groups under the chronic stress condition (Table 2).