Introduction

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The response of the organism to stress includes both physical and behavioral adaptations, the familiar "fight or flight" response (Selye, 1990). Usually, cessation of the stress terminates this response, and the organism returns to its original equilibrium. However, very intense or long-lasting stress results in a new biological equilibrium that can be either beneficial (e.g., exercise-induced conditioning of the cardiovascular system) or detrimental (hypertension or stroke) to the body.

Adrenal hormones (catecholamines and glucocorticoids) play roles in noxious effects of stress in the CNS and cardiovascular system (Stratakis and Chorusos, 1995). The hippocampus is a main neural target site for these hormones in the CNS (Sapolsky et al., 1990; Watanabe et al., 1992). Indeed, it has been reported that exposure to chronic stress in humans leads to hippocampal atrophy (Sheline et al., 1996). Some of the noxious actions of adrenal hormones are mediated through the release of EAA, stimulating N-methyl-D-aspartate receptors (Gilad et al., 1990; Moghaddam, 1993; Magariños and McEwen, 1995).

The response of the cardiovascular system to stress has been ascribed mainly to catecholamine hyperstimulation and involves increased cardiac output and vascular resistance, lipid mobilization and stimulation of platelet aggregation. When exaggerated, these effects can contribute to the pathogenesis of hypertension, atherosclerosis and ischemic heart disease (Bassett and Cairncross, 1977; Hjemdahl et al., 1991).

Some detrimental actions of stress hormones may be mediated via the release of secondary mediators. We have, therefore, investigated a possible role of NO as a mediator of stress-induced injury.

Nitric oxide is synthesized from L-arginine by the enzymes called NOS: the eNOS, nNOS and an isoform expressed during inflammatory reactions (iNOS) (Knowles and Moncada, 1994).

Evidence has been presented for the role of NO in some pathological processes in CNS. Indeed, an excessive generation of NO has been demonstrated in epilepsy, hypoxic-ischemic damage and neurodegenerative disorders including Alzheimer's and Parkinson's diseases and Huntington's chorea (reviewed in Moncada et al., 1991).

In the cardiovascular system, the changes in NO generation and/or action have been implicated in the pathogenesis of vascular disorders including diabetes, atherosclerosis, thrombosis and hypertension (reviewed in Radomski and Salas, 1995).

To study the relationship between changes induced by stress and NO, we have used JCR:LA-cp rats. These rats exhibit a corpulent phenotype when homozygous for the cp gene. Male cp/cp rats are obese, hyperphagic, very low density lipoprotein hyperlipemic, hyperinsulinemic and markedly insulin resistant. Furthermore, male cp/cp rats spontaneously develop atherosclerosis and ischaemic myocardial lesions as they age (Russell et al., 1990). In contrast to cp/cp, heterozygotes (cp/+) or rats that are homozygous normal (+/+) are phenotypically lean and metabolically normal. Interestingly, stress induces fat mobilization in these animals (McArthur D, personal communication). These characteristics are similar to the hypermetabolic syndrome induced by stress in humans (Mizock, 1995). Thus, JCR:LA-cp rats provide a good model of the common triad in humans obesity-mild type II diabetes-hyperlipemia that leads to a high risk of cardiovascular disease and, in principle, to a major susceptibility to stress.

	Methods

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Animals. The 12- to 14-wk-old male JCR:LA rats, both lean (+/+?) and obese (cp/cp) phenotypes were used. All experimental protocols adhered to the guidelines of the Canadian Council on Animal Care and were approved by the Health Sciences Animal Welfare Committee of the University of Alberta. The rats were housed individually in polycarbonate cages under standard conditions of temperature and humidity and a 12-hr light/dark cycle (lights on at 08:00 A.M.) with free access to food and water. All animals were maintained under constant conditions for 4 to 7 days before stress. Animals, food and water were weighed daily to monitor body weight and food and water intake.

Immobilization stress. Rats were exposed to stress between 09:00 and 11:00 A.M. in a room adjacent to the animals home room. The immobilization was performed using a plastic film rodent restrainer (Decapi-cone, Braintree, MA) that allowed for a close fit to both lean and obese rats. The following restraint protocols were used: a single 15-min session, and 1 hr every day for 4, 9 or 14 days. Control animals were not subjected to stress, but were accustomed to frequent handling. Animals were killed immediately after the last session of immobilization (still in the restrainer) using halothane.

Treatment. Some animals were treated with oral L-NAME for 15 days (starting 1 day before the 14 days of stress). The dose selected (150 µg/ml in drinking water) had no significant effect on blood pressure in both control and stressed animals.

Blood pressure and platelet preparation. After induction of anesthesia with halothane, blood pressure was recorded through a catheter inserted into a femoral artery. Blood was collected by cardiac puncture and anticoagulated in the presence of tri-sodium citrate (3.15% w:v, 1 vol citrate per 9 vol blood). Platelet-rich plasma was obtained by centrifugation at 220 × g for 15 min at room temperature. Platelets were collected from plasma after centrifugation at 800 × g for 10 min at room temperature.

Tissue collection. Samples of aorta, myocardium, brain (cortex, hippocampus, hypothalamus and cerebellum) and adrenal glands were snap-frozen in liquid nitrogen and stored until assayed at 80°C.

Glutamate uptake in forebrain synaptosomes. After decapitation, a part of the forebrain (300 mg) was dissected on ice. All subsequent steps were performed at 4°C. The tissue was immediately homogenized in 25 vol (w/v) of 0.32 M sucrose in a glass homogenizer fitted with a Teflon pestle. The homogenate was centrifuged at 200 × g for 10 min and the supernatant was centrifuged at 20,000 × g for 20 min. The pellet was resuspended in 0.32 M sucrose and centrifuged at 20,000 × g for 20 min. The crude synaptosomal pellet was finally resuspended in 3 ml of 0.32 M sucrose and used for the assays. Sodium-dependent glutamate uptake was measured according to the procedure described by Robinson et al. (1991) with some modifications. Briefly, 25-µl aliquots of synaptosomes were added to 250 µl of incubation buffer (Tris 5 mM, HEPES 10 mM, KCl 2.5 mM, NaCl 1.4 M, CaCl₂ 1.2 mM, MgCl₂ 1.2 mM, K₂HPO₄ 1.2 mM, dextrose 10 mM) containing L-[³H] glutamic acid (Amersham, Buckinghamshire, UK) 0.125 µM and incubated for 3 min at 37°C in a shaking bath. The reaction was terminated using 1 ml ice cold choline buffer (incubation buffer in which equimolar concentration of choline chloride was substituted for NaCl), and the samples were centrifuged at 10,000 × g for 4 min to recover synaptosomes. The [³H]-bound radioactivity was measured in a liquid scintillation counter (Beckman LS-6500).

NO synthase activity. Frozen tissues were homogenized by sonication (VibraCell) in an ice-cold buffer (pH 7.4) containing Tris HCl (50 mM), sucrose (320 mM), dithiothreitol (1 mM), leupeptin (10 µg/ml) soybean trypsin inhibitor (10 µg/ml) and aprotinin (2 µg/ml), followed by centrifugation at 10,000 × g for 20 min 4°C. The samples (40 µl) of supernatant were incubated at 37°C for 20 min in a buffer: KH₂PO₄ (50 mM), MgCl₂ (1 mM), CaCl₂ (0.2 mM), L-valine (50 mM), L-citrulline (1 mM), L-arginine (20 µM) and dithiothreitol (1.5 mM) containing L-[¹⁴C]-arginine (0.5 µCi/ml, Amersham, Oakville, Ontario, Canada). The reaction was terminated by removing the substrate by the addition of 1 ml of 1:1 H₂O Dowex AF 500W-8 resin (Bio-Rad). The activity of the calcium-dependent NOS was calculated from the difference between L-[¹⁴C]-citrulline produced from control samples and samples containing ethylene glycol-bis(-aminoethyl ether) N,N,N'-tetraacetic acid (EGTA, 1 mM); the activity of the calcium-independent isoform was determined from the difference between samples with EGTA and samples containing 1 mM N^G-monomethyl-L-arginine. The [¹⁴C]-bound radioactivity was counted using a liquid scintillation counter (Salter et al., 1991).

Western blot. Proteins in tissue homogenates were subjected to 7% sodium dodecyl sulphate-polyacrylamide gel electrophoresis. Electrophoresis was carried out in reducing conditions according to Laemmli (1970). After electrophoresis, samples were electroblotted onto polyvinylidene fluoride membranes and proteins identified and detected using monoclonal antibodies against eNOS and nNOS (at concentration 0.1 µg/ml, Transduction Laboratories, Lexington, KY) or polyclonal antibodies against iNOS (at concentration 0.1 µg/ml; Santa Cruz Biotechnology, Santa Cruz, CA) and ECL kit (Amersham).

cGMP levels. Tissue samples were homogenized as described above but the homogenizing buffer contained 100 µM 3-isobutyl-1-methylxanthine and the homogenate assayed for cGMP content using enzyme immunoassay system (Amersham).

Excitatory amino acids in brain tissue. Concentrations of aspartate, glutamate, asparagine, serine, glutamine, glycine and -aminobutyrate were measured using HPLC. The samples were derivatized with o-phtalaldehyde and mercaptoethanol and assayed using an HPLC system (Gilson) linked to a fluorescence detector (Waters 420; Milford, MA) as described by Moghaddam (1993).

Plasma and tissue nitrite and nitrate. The stable NO metabolites nitrate and nitrate (NO_x) were measured using the Griess reaction. Briefly, nitrate was reduced stoichiometrically to nitrite by incubating sample aliquots (100 µl) for 20 min at 37°C in the presence of 0.1 U/ml nitrate reductase, 50 µM FAD and 50 µM NADPH. Lactate dehydrogenase (10 U/ml) and 10 mM sodium pyruvate were then added to oxidize NADPH to avoid any interference with the following nitrite determination. Total nitrite was then determined spectrophotometrically using a microplate reader (Bio-Rad 550; Hercules, CA).

Plasma cortisol. Cortisol (17-hydroxycorticosterone) was determined by radioimmunoassay (Immunotech, Coulter, Fullerton, CA).

LDH. The viability of the synaptosomes was determined by measuring the activity of LDH (Cancela and Beley, 1995). Briefly, synaptosomes were incubated for 5 min at 37°C in incubation buffer that did not contain glutamate. Synaptosomes were then pelleted by centrifugation at 10,000 × g for 30 min at 4°C. Supernatants were assayed for LDH activity, and the pellet was resuspended with 1 ml of 0.32 M sucrose and pelleted again. The pellets were sonicated in 1 ml of phosphate buffer (KH₂PO₄/K₂HPO₄ 50 mM, pH 7.5) and used to measure LDH activity in the synaptosomes.

Vascular contractility. After bleeding the animal by heart puncture, the thoracic aorta was removed, trimmed of adhering fat and connective tissue and cut into 3-mm long transverse rings. The aortic rings were mounted on stainless-steel hooks under 1.5 g resting tension in 20 ml organ baths and bathed at 37°C in Krebs solution containing (mM) NaCl 116, KCl 5.4, CaCl₂ 1.2, MgCl₂ 2, Na₂PO₄ 1.2, EDTA 0.023, glucose 10 and NaHCO₃ 19 and aerated with 95% O₂ and 5% CO₂. Tension was recorded isometrically with Grass FTO3C transducers and displayed on a Digi-Med tissue force analyzer (model 210) which was linked to an IBM compatible computer. Data were collected and analyzed using an DMSI 210/4 data reduction program (Micro-Med). Contractions to phenylephrine (1 nM-10 µM) were examined. The EC₈₀ contraction by phenylephrine was used to examine endothelium-dependent acetylcholine- (1 nM-10 µM) induced relaxation.

Platelet aggregation. Platelet aggregation was measured in whole blood using a whole blood platelet aggregometer (Chrono-Log, Havertown, PA). Aggregation was initiated by collagen (2-20 µg/ml) and analyzed using an Aggro-Link data reduction system.

Protein assay. Proteins were measured using a Bio-Rad kit.

Statistical analysis. Data are expressed as mean ± S.E.M. and comparisons between groups were performed using analysis of variance followed by Newman-Keuls's test, with P < .05 considered as significant.

Chemicals. Unless otherwise stated, all drugs and compounds were purchased from Sigma Chemical Co. (St Louis, MO).

	Results

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Body weight and food ingestion was not modified during 4, 9 or 14 days of repeated stress in both cp/cp and +/+? rats.

Brain. A long-lasting (14 days) immobilization stress induced a decrease in glutamate uptake in forebrain synaptosomes in lean, but not in obese (fig. 1a) rats. This impairment correlated with an increase in the release of LDH from synaptosomes (fig. 1a). In addition, the levels of glutamate significantly increased in hippocampus after 14 days of repeated immobilization in lean (control: 1883.2 ± 113; 14 days: 2294.3 ± 116 pmol/mg tissue, P < .05, n = 4-6), but not in obese rats (control: 2345.1 ± 105; 14 days: 2298.5 ± 108 pmol/mg tissue). The obese rats showed higher control concentrations of glutamate than lean rats (P < .05, n = 4-6).

View larger version (23K): [in this window] [in a new window]   Fig. 1.   Stress-induced hippocampus damage in JCR:LA-cp rats and its prevention by NOS inhibition with L-NAME. A, Glutamate uptake by synaptosomes from rats subjected to immobilization stress; open bars, +/+ rats; solid bars, cp/cp rats. B, Effect of L-NAME on uptake of glutamate by synaptosomes from +/+ rats subjected to immobilization stress for 14 days; open bars, control; shaded bars, 14 days of restraint. The LDH released from the synaptosomes is indicated below the relevant bar. The values are mean ± S.E.M. (n = 5-16), expressed as % of glutamate uptake (0.167 ± 0.01 µM/mg protein) and LDH release (49.868 ± 0.05 absorbance units) by control synaptosomes. ** P < .01, stress vs. control.

View larger version (25K): [in this window] [in a new window]   Fig. 2.   Increased generation of NO and the corresponding cGMP levels in the brain during chronic stress. Calcium-dependent NOS activity (bars), protein content (insert blots) and cGMP content (points and line) in hippocampus from +/+ rats subjected to immobilization stress. There were no significant changes in the expression of nNOS protein (control: 547.6 ± 214; 15 min: 431 ± 309; 4 days: 886.3 ± 363, 9 days: 550 ± 190; 14 days: 540.2 ± 181, arbitrary units, n = 2-5, Western blot shown in inserts). Values are mean ± S.E.M., n = 5-16. ** P < .01, * P < .05, vs. control.

View larger version (12K): [in this window] [in a new window]   Fig. 3.   Stress-induced accumulation of NO metabolites (nitrite and nitrate, NOx) in the forebrain. Values are mean ± S.E.M., n = 5-16. ** P < .01, * P < .05, vs. control.

Cardiovascular system. In nonstressed lean animals blood pressure was 76 ± 1 mmHg (n = 4-6). The aortic rings from these rats showed endothelium-dependent relaxation to acetylcholine with an EC₅₀ of 27 ± 1.5 nM and calcium-dependent NOS activity of 9.7 ± 2.1 pmol citrulline/min/mg protein (n = 4-6). The activity of calcium-dependent NOS in the myocardium was 7.7 ± 3.3 pmol/min/mg protein (n = 6-14). In all animals nonstressed and stressed lean and obese, blood pressure, endothelium-dependent relaxation, and aortic and myocardial NOS activities were similar (P > .05, n = 4-14).

Platelets. Chronic stress caused a decrease in calcium-dependent NOS activity in the platelets of both lean and obese rats (fig. 4). Collagen induced platelet aggregation both in lean and obese rats with EC₅₀ values of 11.2 ± 0.2 and 10.8 ± 0.6 µg/ml respectively, which were not significantly different from each other (P > .05, n = 4-6), and were not modified (P > .05, n = 4-6) by the exposure of animals to stress.

View larger version (14K): [in this window] [in a new window]   Fig. 4.   Stress-induced reduction of calcium-dependent NOS in platelets of +/+ (open bars) and cp/cp (solid bars) rats. Values are mean ± S.E.M., n = 5-16. * P < .05, vs. control.

Adrenal glands. Acute stress (15 min of restraint) increased the activity of calcium-dependent NOS in both lean and obese rats. In contrast, chronic stress (14 days) led to a reduction of NOS activity in the adrenal glands (fig. 5). Both acute and chronic stress resulted in increased cortisol levels in plasma, an effect that was not modified by the treatment with L-NAME (table 1).

View larger version (13K): [in this window] [in a new window]   Fig. 5.   Biphasic changes (stimulation by acute and inhibition by chronic stress) in calcium-dependent NOS activity in the adrenal glands of +/+ (open bars) and cp/cp (solid bars). Values are mean ± S.E.M., n = 5-16. ** P < .01, * P < .05, vs. control.

Plasma nitrite and nitrate. The levels of nitrite and nitrate in plasma increased in correlation with the duration of stress peaking at 4 to 9 days in lean and at 9 days in obese rats. The treatment with L-NAME for 15 days significantly attenuated the stress-induced increase in nitrite and nitrate in lean rats (fig. 6).

View larger version (14K): [in this window] [in a new window]   Fig. 6.   Chronic stress increased the levels of NO metabolites, nitrite and nitrate (NOx), in plasma of +/+ (open bars) and cp/cp (solid bars). The increase in NOx levels in lean rats was attenuated by the treatment with L-NAME. Values are mean ± S.E.M., n = 5-16. ** P < .01, vs. control, P < .05, vs. 14 days. L-NAME: rats treated for 14 days with L-NAME.

	Discussion

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The exposure of JCR-LA-cp rats to a classical stress paradigm (restraint) resulted in a generalized increase in the generation of NO. In the CNS, increased generation of NO and cGMP preceded both functional (decrease in glutamate uptake) and structural (increase in LDH activity) neuronal damage that was detectable after 9 to 14 days of restraint. These changes correlated with chronic elevation of cortisol levels and accumulation of NO metabolites, nitrite and nitrate in plasma. A NOS inhibitor, L-NAME, attenuated stress-induced increases in nitrite and nitrate, prevented neuronal damage induced by chronic stress despite the elevated cortisol levels in plasma. These results indicate that NO plays a major pathogenetic role as a mediator of stress-induced neurotoxicity.

Under normal conditions, glutamate is removed from the synaptic cleft via reuptake into presynaptic terminal and diffusion out down the glutamate concentration gradient. Synaptosomes are a good model to study glutamate transport alterations (Nicholls and Attwell, 1990). Indeed, chronic inhibition of glutamate uptake by synaptosomes has been used to mimic slow-developing neurotoxicity in vivo. This may be a pathological mechanism involved in neurodegeneration of Huntington's disease, Alzheimer's disease and amyotrophic lateral sclerosis (Rothstein et al., 1993). Using this model, we have found that chronic stress impaired the mechanism of washing of glutamate leading to this accumulation in the extracellular milieu. This is likely to cause neurodegeneration and death.

The cellular mechanism of neurodegeneration after chronic inhibition of glutamate uptake may be receptor and glucocorticoid dependent (Virgin et al., 1991). Our results showed that inhibition of NOS by L-NAME normalized glutamate uptake impaired by the exposure of rats to chronic stress in a cortisol-independent manner. These data are consistent with previous evidences that some cellular actions of glutamate are mediated by NO (Garthwaite et al., 1988). During the exposure of organism to chronic stress the relationship between NO and glutamate is likely to operate as a positive feedback loop mechanism and stress-induced augmentation of neuronal NO may, in turn, impair the sequestration of glutamate (Pogun et al., 1994). Indeed, in rat hippocampal synaptosomes, NO donors inhibited glutamate uptake, an action reversible by the removal of NO using hemoglobin (Pogun et al., 1994).

It is unclear whether NO per se is the effector molecule of neuronal damage. A persistent elevation of NO levels can result in generation of a potent oxidant, peroxynitrite (ONOO) from superoxide and NO. Peroxynitrite is a tissue-damaging agent that acts through initiation of lipid peroxidation, oxidation of sulfhydryl groups and nitrosation of tyrosine-containing molecules and these effects may account for inhibition of EAA reuptake by ONOO (Trotti et al., 1996). We have also found that increased generation of NO in the hippocampus is paralleled by elevated cGMP formation. The role of cGMP in neurodegeneration is unclear: cGMP, stimulating the release of EAA from neurons (Garthwaite et al., 1988) could further contribute to the mechanism of NO/ONOO-mediated neuronal damage, but it could act also as a neuroprotective agent (Garthwaite, 1982).

We have found that the changes in glutamate and NO pathways are largely confined to the hippocampus and were not detectable in cortex, hypothalamus and cerebellum. Interestingly, in the hippocampus, NO has been identified as the retrograde mediator of long-term potentiation (Bohme et al., 1991), a phenomenon in which glutamate uptake inhibition contributes to the strengthening of glutamate neurotransmission and to the development of learning and memory. Stressful stimuli interfere with learning and memory (McEwen, 1995). It is, therefore, plausible that these stimuli exert their effects through a NO-dependent mechanism.

The hypothalamus-pituitary-adrenal axis function is an important determinant of stress. Calzà and colleagues (1993) reported an increase in hypothalamic NOS mRNA in rats after acute restraint stress indicating that NO could mediate ACTH release. In JCR:LA-cp rats, no significant changes in NOS could be detected in the hypothalamus. However, there was an early (15 min stress) increase in NOS activity in adrenal glands. The adrenal NO is believed to be involved in controlling catecholamine secretion by chromaffin cells through cGMP-dependent mechanism (Moro et al., 1993). However, its actions on steroidogenesis are still controversial. We have found that increased formation of NO in the adrenal gland correlates with cortisol release during acute but not chronic stress. Thus, in chronic stress, glucocorticoid generation and release may depend on factors other than NO.

The effects of stress on feeding behavior and body weight are still controversial. Although various studies indicate a decrease in these parameters (Haleem and Parveen, 1994), some recent studies indicate increase or no change (Sánchez et al., 1998). In our model, the lack of effects of stress in hypothalamic NOS is consistent with the lack of effects on feeding and body weight, indicating that this level of stress is not intense enough to disrupt hypothalamic function.

In contrast to lean animals, neuronal damage was not detectable in stressed obese cp/cp rats. This could be related to high resting levels of cortisol (20.9 ± 2.9 vs. 11.3 ± 1.8 nM, P < .05, n = 4-6) and glucose (146 ± 7 vs. 132 ± 11 mg/dl as published by Amy et al., 1988) in obese vs. lean animals, respectively. Cortisol inhibits glucose uptake in brain which, in turn, may decrease the uptake of glutamate, leading to enhanced extracellular levels of the amino acid (Virgin et al., 1991). The constant exposure of obese rats to glutamate could precondition these animals and make them less susceptible to stress-induced neuronal damage.

The cardiovascular system is also a major target of stressful reactions. We have found that the exposure of rats to stress for 4 and 9 days leads to an increase in plasma nitrite/nitrate. Interestingly, a similar transient stimulation of NO release occurs also in humans subjected to stress (Mizock, 1995). This release of NO may offset the vasoconstrictor effects of adrenal hormones in the vasculature.

The changes in NO generation detected during stress exerted no apparent effect on blood pressure or aortic contractility in vitro. In this hemodynamic response to early phases of chronic stress, JCR:LA-cp rats react in a similar way to humans (Benschop et al., 1994) but not to Wistar (Gamallo et al., 1988) or Sprague-Dawley (Blake et al., 1995) rats that show a hypertensive response during the exposure to stress.

Long-lasting stressful situations may be involved in the pathogenesis and/or development of various diseases associated with abnormal hemostasis, such as thrombosis and disseminated intravascular coagulation (Levine et al., 1985). We have found that the exposure of rats to stress for 14 days led to reduced generation of NO by platelets without concomitant up-regulation of aggregation. It is known that platelets generate NO during the process of adhesion and aggregation, and its role is to down-regulate the extent of platelet activation (Radomski et al., 1990). Furthermore, NO released by the endothelium is able to dissipate preformed platelet aggregates (Radomski et al., 1987). Thus, platelets are under the influence of the NO they produce and the endothelium-released NO. Therefore, despite the decreased generation of NO by platelets after 2 wk of restrain, because more NO is generated by the vasculature, it is likely that this is sufficient to counteract the action of pro-aggregatory mediators of stress.

This compensatory ability of the vasculature to generate NO is exhaustible, as 14 days of stress leads to a decrease in NO generation predisposing these animals to the hypertensive and thrombotic events. The mechanism of exhaustion may be related to NOS substrate depletion as L-arginine levels decrease during stress (Milakofsky et al., 1993).

In conclusion, our results indicate that systemic generation of NO plays an important role in early phases of chronic stress. In the CNS, NO appears to be detrimental. In contrast, in the cardiovascular system, the release of NO may be beneficial as they attenuate the vasoconstrictor and pro-aggregatory effects of stress hormones and mediators. Finally, the pharmacological strategies aiming at selective inhibition of nNOS and stimulation/substitution of eNOS with platelet and vascular smooth muscle-selective NO donors may represent a novel therapeutic approach to the treatment of stress-related pathologies.