| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
BRIEF COMMUNICATION |
Center for Advanced Oral Medicine (T.S.), Hokkaido University Hospital, Sapporo 060-8586, Japan; and Departments of Oral Functional Science (M.M., T.K., W.N., S.O., T.H., H.O.) and Oral Pathobiological Science (M.S.), Graduate School of Dental Medicine, Hokkaido University, Sapporo 060-8586, Japan
Address all correspondence and requests for reprints to: Tetsuo Shirakawa, Ph.D., Center for Advanced Oral Medicine, Hokkaido University Hospital, N13W6 Kita-ku, Sapporo, 060-8586, Japan. E-mail: tshira{at}den.hokudai.ac.jp.
Abstract
Production of nitric oxide (NO) in the hypothalamic paraventricular nucleus (PVN) was examined by microdialysis in rats subjected to immobilization (IMO) stress. A dialysis probe was implanted in the posterior magnocellular subdivision of the PVN and nitrite (NO2–), an oxidized product of NO, was measured continuously. NO2– concentration in dialysate was enhanced to 156% after 30 min of IMO compared with the NO2– level before IMO. Intraperitoneal administration of NG-monomethyl-L-arginine (10 mg/kg), a NO synthase inhibitor, before IMO completely inhibited the increase of NO production that IMO was to induce. Depletion of catecholamines innervating the PVN by an injection of 6-hydroxydopamine into the lateral ventricle before the microdialysis had no suppressive effect on the increase of NO production by IMO. In contrast, NO2– levels in the PVN were lowered by continuous perfusion of the solution containing the ionotropic glutamate receptor antagonists 2-amino-5-phosphonovaleric acid (500 µM) and 6-cyano-7-nitroquinoxaline-2, 3 dione (50 µM) through the dialysis probe, and the IMO-induced increase of NO production was attenuated by the treatment. These results suggest that catecholaminergic drive to the hypothalamus is not necessary for the IMO-induced increase of NO production and that ionotropic glutamate receptors play a role in the basal and IMO-induced NO production.
NITRIC OXIDE (NO) is formed from L-arginine through activation of NO synthase (NOS) in neurons (1). Immunohistochemical staining of neuronal NOS (2) or the reduced nicotinamide adenine dinucleotide phosphate-diaphorase histochemical staining (3) revealed strong NOS labeling of neurons in the hypothalamic paraventricular nucleus (PVN), the well-documented center of stress response in the brain. Localization of NOS-positive neurons in both magnocellular and parvocellular parts of the PVN and up-regulation of neuronal NOS mRNA expression or c-fos expression in the NOS-positive neurons by immobilization (IMO) stress have also been demonstrated (4, 5, 6, 7).
The PVN receives multiple inputs from many brain areas implicated in stress response. Subsets of PVN neurons are predominantly innervated by adrenergic and noradrenergic fibers arising mainly from the nucleus of the solitary tract and the ventrolateral medullary reticular formation (8, 9). These include catecholaminergic neurons, which project to the PVN to activate parvocellular neurosecretory neurons that produce CRH and magnocellular neurons that produce vasopressin or oxytocin. The excitatory amino acids (EAAs), such as L-glutamate, may also be released by some PVN afferents and cause responses in PVN neurons through postsynaptic mechanism. Electrical stimulation of the perifornical region evokes excitatory postsynaptic potentials and currents in both magnocellular and parvocellular neurons, and the response is blocked by EAA antagonists (10, 11). By an injection of the retrograde tracer [3H]D-aspartate into the PVN, it was shown that all intrahypothalamic and telencephalic afferents to this nucleus contained glutamatergic/aspartatergic fibers (12). However, the role of EAAs in the stress-induced NO synthesis in the PVN is yet unclear (13). In the present study, we aimed to investigate the effects of IMO on an acute production of NO in the PVN and to clarify whether glutamatergic and/or catecholaminergic inputs are involved in the response of PVN neurons that produce NO.
Materials and Methods
Animals
Male Sprague Dawley rats weighing 400–450 g were used. They were housed in a constant environment (lights on from 0900 to 2100 h; temperature, 23 ± 1 C; humidity, 60 ± 5%) and given free access to standard laboratory food and water. All procedures were approved by the Animal Care and Use Committees of Hokkaido University Graduate School of Dentistry, and all efforts were made to minimize animal suffering. A dummy probe was inserted stereotaxically into the brain with a stainless steel guide cannula under pentobarbital anesthesia (pentobarbital sodium, 50 mg/kg, ip). The guide cannula was placed unilaterally about 2.0 mm above the right PVN, and the tip of the dummy probe was protruded 1.0 mm from the lower end of the guide cannula. The stereotaxic coordinates for implantation of dialysis probe were decided by reference to the brain atlas by Paxinos and Watson (14); the coordinates were 2.5 mm posterior to the bregma, 0.5 mm lateral to the midsagittal sinus, and 6.0 mm ventral to the surface of the brain, which enabled positioning of the membrane portion of the dialysis probe at the posterior magnocellular subdivision of the PVN. The guide cannula was fixed to the skull with epoxy adhesive, and a swivel was attached to the cannula to avoid entanglement of the sampling tubes. After the surgery, the rats were moved into an individual cage (36 x 30 x 34 cm) and adapted to the experimental conditions for 4 d.
PVN microdialysis
In vivo microdialysis was performed as described previously (15, 16). On the day of microdialysis, rats were lightly anesthetized with diethylether for the replacement of the dummy probe with a dialysis probe (150 µm in outer diameter) between 0930 and 1000 h. The dialysis probe consisted of cuprophane membrane, a 25-gauge stainless steel needle, and fused-silica tubes. An efficient area of the dialysis membrane was approximately 0.6 mm2, and the cut-off molecular weight was 1000. The probe was perfused with balanced Earle’s salt solution (GIBCO, Tokyo, Japan) continuously throughout the experiment, and the dialysate was collected every 30 min into a sampling tube.
IMO procedure
Perfusion of the dialysis probe was started at 1100 h at a constant flow rate of 1 µl/min. For acute IMO, animals were placed in the prone position from 1300 to 1330 h on a restraint apparatus originally designed in our laboratory. The trunk of the animals was held inside the apparatus, and their movement was minimized with flexible tapes. At the end of IMO, rats were immediately returned to the individual cage, and the sampling of dialysate was continued for at least 3 h. After the sampling, rats were decapitated under pentobarbital anesthesia, and the brains were removed from the skull and fixed with 4% paraformaldehyde for 2–3 d. Cresyl violet staining was carried out on 50-µm-thick brain sections to determine the position of the dialysis probe. Effects of IMO on NO production in the PVN were evaluated in 12 rats, and unstressed rats (n = 6) were used as control (experiment 1).
Drugs
NG-monomethyl-L-arginine (L-NMMA), 2-amino-5-phosphonovaleric acid (AP5), and 6-cyano-7-nitroquinoxaline-2, 3 dione (CNQX) were purchased from Research Biochemicals International (Natik, MA). 6-Hydroxydopamine (6-OHDA) was purchased from Sigma (St. Louis, MO). L-NMMA (10 mg/kg, ip), an inhibitor of NO synthesis from arginine, was dissolved in 100 µl of saline and administered to rats (n = 9) 30 min before the start of IMO (experiment 2). Treatment of rats with 6-OHDA, a neurotoxin that destroys catecholaminergic system in the brain, was carried out by injecting 250 µg of 6-OHDA dissolved in 5 µl of saline into the lateral ventricle under pentobarbital anesthesia 1 wk before the day of microdialysis (n = 10). The rats were subjected to IMO, as described earlier, and the dialysates were divided into two groups, one for an assay of nitrite (NO2–) and NO3–(NOx–, n = 5) and the other (n = 5) for a norepinephrine (NE) assay (experiment 3). NE was also measured in 6-OHDA-uninjected rats subjected to IMO (n = 6). The N-methyl-D-aspartate (NMDA) receptor antagonist AP5 and the non-NMDA receptor antagonist CNQX were dissolved in Earle’s salt solution and perfused through the dialysis probe in the PVN (n = 9), beginning 90 min before IMO until the end of sampling (experiment 4). The final concentrations of AP5 and CNQX mixed in Earle’s solution were 500 µM and 50 µM, respectively.
Determination of NO2–, NO3–, and NE concentrations
Dialysates were stored at 4 C, and the NOx– assay was carried out on the day of microdialysis within 2 h after sampling. NO2– and NO3– concentrations in the dialysates were determined by the use of an automated NOx– detector-HPLC system (ENO-10; Eicom, Kyoto Japan). The principle of the NOx– detection in our system was described elsewhere (16, 17). Briefly, each fraction of dialysate was added to a buffer solution composed of 10% methanol, 0.15 M NaCl-NH4Cl, and 0.5 g/liter 4Na-EDTA, and the mixture was delivered to the separation column (Eicom). Separated NO2– was mixed with the Griess reagent composed of 1.25% HCl, 5 g/liter sulfanilamide, and 0.25 g/liter N-naphthylethylenediamine, which resulted in coloring of the solution into purple in proportion to the NO2– concentration. Absorbance at 540 nm was measured by the use of a flow-through spectrophotometer (Eicom). For the determination of NO3– concentration, the separated NO3– fraction was passed through a copper-cadmium reduction column (Eicom) to reduce NO3– to NO2–, and the NO2– concentration was determined as described earlier. The quantity limit of this assay system to detect NO2– and NO3– was 1.0 pmol. NE in dialysate was determined by the use of HPLC with electrochemical detection as reported previously (15), and the minimum quantity for NE detection was 1.0 pg.
Statistical analysis
To evaluate the difference of the NOx– levels before and after IMO in each group, we employed the Dunnett’s method as a planned test. Significance of time-dependent changes in NOx– in the course of IMO experiments (from 30 min before the onset of IMO to 120 min after the onset of IMO) was determined by one-way ANOVA for repeated measures, and two-way ANOVA was used to compare the difference between the experimental groups. The Tukey-Kramer test was used to compare means within and between groups after the analyses done by ANOVA. All data were expressed as means ± SEM, and differences were considered significant at P < 0.05.
Results
IMO increased NO production in the PVN
Levels of NO2– and NO3– in the PVN dialysate measured before IMO were 11.45 ± 2.46 and 48.55 ± 8.02 pmol/30 min, respectively. Both NO2– and NO3– concentrations reached their peaks (17.89 ± 2.78 and 75.11 ± 12.27 pmol/30 min, respectively) 60 min after the onset of IMO (experiment 1; Fig. 1
, A and B). We found a significant effect of IMO on both NO2– and NO3– levels by comparing the peak values with the levels before IMO (Dunnett test, P < 0.05). The time-dependent variations of both NO2– and NO3– induced by IMO were significant (repeated measures ANOVA: F(5,55) = 2.69, P = 0.031 for NO2– and F(5,55) = 3.29, P = 0.011 for NO3–). On the other hand, at overall level, neither NO2– nor NO3– in the stressed group was significantly different from the counterpart in control group (two-way ANOVA: F(1,96) = 2.95, P = 0.089 for NO2– and F(1,96) = 2.14, P = 0.147 for NO3–), indicating that the 30-min IMO elevated NOx– levels in the PVN for only a short period. Administration of L-NMMA before IMO (experiment 2; Fig. 1C) completely inhibited the increase of NO2– caused by IMO (repeated measures ANOVA: F(5,40) = 1.56, P = 0.195), and a significant difference was found in NO2– levels between the L-NMMA-injected and uninjected groups (two-way ANOVA: F(1,124) = 3.97, P = 0.048). Because NO in an aqueous solution is oxidized primarily to NO2– (18), we regarded the increase of NO2– in dialysate after IMO as an indication of acute NO production around the dialysis membrane implanted in the PVN (19).
|
NO2–) from the basal level was calculated in 6-OHDA-injected rats and uninjected rats (rats subjected to IMO in experiment 1; Fig. 2A). There was no statistical difference in NO2– between the two groups (F(1,90) = 0.17, P = 0.676). We confirmed that 250 µg of 6-OHDA injected into the lateral ventricle suppressed IMO-induced release of NE from the terminals in the PVN (20). IMO rapidly elevated NE levels in uninjected rats, whereas the effect of IMO on NE release was small in 6-OHDA-injected rats. NE reached its peak 30 min after the onset of IMO, and the level was 8.03 ± 1.05 pg/30 min in uninjected rats and 4.59 ± 1.27 pg/30 min in 6-OHDA- injected rats. The time-dependent changes in NE were significant in uninjected rats (F(5,25) = 8.60, P < 0.001) but not in 6-OHDA-injected rats (F(5,20) = 0.73, P = 0.605). Difference of the NE concentration (NE) from the basal level was calculated in 6-OHDA-injected rats and uninjected rats (Fig. 2B). A significant difference was found in NE values between the two groups (F(1,54) = 17.60, P = 0.0001).
|
NO2– from the basal level was calculated in the AP5/CNQX-treated rats and untreated rats (rats subjected to IMO in experiment 1; Fig. 2C
). A significant difference was found in NO2– values between the two groups (F(1,114) = 26.96, P < 0.0001). In AP5/CNQX-treated rats, NO2– in the PVN was increased moderately by IMO, whereas the time-dependent change in NO2– was significant (F(5,40 = 3.93, P = 0.005). Discussion
IMO rapidly increased production of NO in the PVN. This effect of IMO was inhibited by ip administration of the NOS inhibitor L-NMMA. These results indicate that the enhanced production of NO after IMO was mediated by an activation of NOS. A number of studies demonstrate the presence of NO-producing neurons in the PVN (2, 3, 4, 5, 6, 7); they are localized in both the magnocellular and parvocellular parts of the PVN. Among the NO-producing neurons identified by the histochemical reaction for nicotinamide adenine dinucleotide phosphate diaphorase, medium- to large-sized neurons are located in the posterior magnocellular subdivision (5, 7), and about one half and one third of them show oxytocin- and vasopressin-immunoreactivity, respectively (5). In the present study, we found enhancement of extracellular NOx concentration by IMO in the PVN in situ. It is likely that IMO not only activates NO production but also influences the levels of oxytocin and/or vasopressin in the PVN. In fact, a significant rise in the release of oxytocin and vasopressin caused by a forced swimming stress (21) was reported in the rat PVN.
There is a growing body of evidence indicating that the PVN receives glutamatergic innervations from large brain areas involving PVN itself and several other nuclei in and outside the hypothalamus (12, 22). Among the neuroanatomical origins of glutamatergic afferents to the PVN, dorsomedial hypothalamic nucleus is the candidate locus for glutamatergic neurons that could be activated by IMO. Microinjection of NMDA into the dorsomedial hypothalamic nucleus caused an increase in glutamate release in the PVN (23) and resulted in a cardiovascular response very similar to the one evoked by an acute emotional stress (23, 24). An in situ hybridization study revealed that mRNAs of NMDA and 
-amino-3-hydroxy-5-methylisoxazole-propionate receptors, the major ionotropic glutamate receptors mediating the EAA signals, exist in the PVN (25). Application of NMDA or glutamate into the PVN induced increases in NO production (13, 26) and elicited a change in heart rate (27). As we have demonstrated in Fig. 2C, continuous perfusion with ionotropic glutamate receptor antagonists through the dialysis probe lowered basal NO levels and attenuated IMO-induced NO production in the PVN. These observations indicate that PVN neurons produce NO, at least in part, through the activation of ionotropic glutamate receptors, and this NO production is enhanced by IMO. The mechanism by which IMO raised NO levels moderately in the PVN in the presence of AP5 and CNQX remains unknown. Reports showed that a forced swimming stress induced the release of oxytocin and vasopressin within the PVN and the supraoptic nucleus (21, 28) but failed to alter the NO release in the supraoptic nucleus (28). Given these results, we suggest that PVN neurons respond to the stressors and release neuropeptides and NO; this response is mediated probably by both glutamatergic and nonglutamatergic pathways innervating the PVN.
We found that depletion of catecholaminergic afferents by the 6-OHDA treatment affected neither basal NO2– levels nor IMO-induced increase of NO2– in the PVN. It has been shown that both the magnocellular and parvocellular subdivisions of the PVN are innervated by catecholaminergic fibers (8, 9), and we verified that the 6-OHDA treatment was effective in suppressing IMO-induced release of NE in this nucleus. These results can be interpreted to mean that the catecholaminergic inputs arising from the brain stem do not play crucial roles in the production of NO during stressful events. NE was recently shown to act as a modulator of glutamate release in the PVN via actions on presynaptic terminals (29). Bath application of NE caused an increase in the frequency of spontaneous glutamatergic excitatory postsynaptic currents in magnocellular neurons (29). The presynaptically released NE at glutamatergic synapses should have some modulatory effects on neuronal activity but may not be related to the NO production caused by IMO. In contrast, basal and IMO-induced release of NO was significantly lowered by the AP5/CNQX treatment in our study; hence, the glutamatergic transmission may be essential for the production of NO in the PVN.
In conclusion, our study first demonstrates that catecholaminergic drive to the hypothalamus is not necessary for the IMO-induced increase of NO production and that ionotropic glutamate receptors play a role in the basal and IMO-induced NO production in the PVN. A recent report indicated that NO in the PVN, which was released by activation of the NMDA receptor, also inhibited NMDA-mediated increases in sympathetic nerve activity (26). This negative feedback of NO on the glutamate receptor activation within the PVN may ensure adequate physiological responses to stressors.
Footnotes
This work was supported in part by Grants-in-Aid for Scientific Research from the Japan Society for the Promotion of Science (No. 15390631).
Abbreviations: AP5, 2-Amino-5-phosphonovaleric acid; CNQX, 6-cyano-7-nitroquinoxaline-2, 3 dione; EAA, excitatory amino acid; IMO, immobilization; L-NMMA, NG-monomethyl-L-arginine; NE, difference of the norepinephrine concentration; NE, norepinephrine; NMDA, N-methyl-D-aspartate; NO, nitric oxide; NO2–, difference of the nitrite concentration; NO2–, nitrite; NOS, nitric oxide synthase; 6-OHDA, 6-hydroxydopamine; PVN, paraventricular nucleus.
Received January 21, 2004.
Accepted for publication April 29, 2004.
References
This article has been cited by other articles:
 |
|
 |
|
  S. Gingerich and T. L. Krukoff Estrogen in the Paraventricular Nucleus Attenuates L-Glutamate-Induced Increases in Mean Arterial Pressure Through Estrogen Receptor {beta} and NO Hypertension, December 1, 2006; 48(6): 1130 - 1136. [Abstract] [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Endocrinology | Endocrine Reviews | J. Clin. End. & Metab. |
| Molecular Endocrinology | Recent Prog. Horm. Res. | All Endocrine Journals |
