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J Neurosci. Author manuscript; available in PMC 2008 Nov 28.
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PMCID: PMC2518401

Stress Induces a Switch of Intracellular Signaling in Sensory Neurons in a Model of Generalized Pain


Stress dramatically exacerbates pain in diseases such as fibromyalgia and rheumatoid arthritis, but the underlying mechanisms are unknown. We tested the hypothesis that stress causes generalized hyperalgesia by enhancing pro-nociceptive effects of immune mediators. Rats exposed to non-habituating sound stress exhibited no change in mechanical nociceptive threshold, but showed a marked increase in hyperalgesia evoked by local injections of prostaglandin E2 or epinephrine. This enhancement, which developed more than a week after exposure to stress, required concerted action of glucocorticoids and catecholamines at receptors located in the periphery on sensory afferents. The altered response to pronociceptive mediators involved a switch in coupling of their receptors from predominantly stimulatory to inhibitory G-proteins (Gs to Gi), and for prostaglandin E2, emergence of novel dependence on protein kinase C epsilon. Thus, an important mechanism in generalized pain syndromes may be stress-induced co-activation of the hypothalmo-pituitary-adrenal and sympatho-adrenal axes, causing a long-lasting alteration in intracellular signaling pathways, enabling normally innocuous levels of immune mediators to produce chronic hyperalgesia.

Keywords: Fibromyalgia, G-protein, hyperalgesia, protein kinase C-epsilon, Catecholamine, Glucocorticoid

Stress exacerbates generalized pain syndromes such as fibromyalgia (Nilsen et al., 2007b), irritable bowel syndrome (Bach et al., 2006), and interstitial cystitis (Temml et al., 2007), and painful inflammatory diseases like rheumatoid arthritis (Zautra et al., 2007). However, mechanisms mediating the impact of stress on pain are poorly understood.

Clinical studies of the role of stress in the pathogenesis of chronic pain syndromes have implicated the hypothalamo-pituitary-adrenal axis in, for example, fibromyalgia and chronic fatigue syndrome (Crofford et al., 1996; Neeck and Crofford, 2000). Although better known for their anti-inflammatory actions, glucocorticoids (principal stress mediators released by the hypothalamo-pituitary-adrenal axis) have recently been reported to enhance pain, both in animal models of peripheral neuropathy (Wang et al., 2004; Takasaki et al., 2005) and when used for the treatment of osteoarthritis in humans (Wollstein et al., 2007). In addition, we previously reported that stress enhances the hyperalgesic effect of the cytokine bradykinin in the rat (Khasar et al., 2005), an effect mediated by the sympatho-adrenal stress axis mediator, epinephrine.

The present study tested the hypothesis that stress-induced activation of the hypothalamo-pituitary-adrenal and sympatho-adrenal axes exacerbates pain by enhancing the pro-nociceptive effects of immune mediators produced in peripheral tissues. Specifically, we examined the effect of stress on hyperalgesia evoked by prostaglandin E2 (PGE2) or by a mediator of neurogenic inflammation, epinephrine.



Experiments were performed on adult male Sprague Dawley rats (250–350 g; Charles River, Hollister, CA). Rats were housed in the Laboratory Animal Resource Center of the University of California, San Francisco, under a 12-hour light/dark cycle. Animal care and use conformed to NIH guidelines. The University of California, San Francisco Institutional Animal Care and Use Committee approved experimental protocols. Concerted effort was made to reduce the suffering and number of animals used.


Stock solutions of PGE2 (Sigma, St. Louis, MO) were 4 mg/ml in 10% ethanol, with subsequent dilutions made in 0.9% saline. Stock solutions of epinephrine ((-)-epinephrine bitartrate; Sigma) were prepared daily as 4 mg/ml in distilled water, with 4 mg/ml ascorbic acid to minimize oxidation. This stock solution was further diluted in 0.9% saline for use. Working solutions of epinephrine and PGE2 were kept on ice and in subdued light during experiments. Pertussis toxin (Bordetella pertussis; CalBiochem, La Jolla, CA), dissolved in 0.9% saline, was injected intradermally, on the dorsum of the hind paw. Nociceptive thresholds were measured before and 30 minutes after its injection. PGE2 or epinephrine was injected at the same site, 30 minutes after pertussis toxin and nociceptive thresholds were again measured, 10, 15 and 20 minutes later. Corticosterone (HBC complex; Sigma) was also dissolved in 0.9% saline and injected intradermally, 24 hours prior to nociceptive testing or implanted as subdermal pellet. The protein kinase C epsilon translocation inhibitor (PKCεI) peptide (CalBiochem) was injected intradermally at the site of nociceptive testing at a concentration of 1 µg/2.5 µl in distilled water. Nociceptive thresholds were measured before and 30 minutes after its injection. PGE2 was then injected at the same site and nociceptive thresholds measured 10, 15 and 20 minutes later. Injections of PKCεI were immediately preceded by injection of distilled water (2.5 µl), in the same syringe, which produces hypo-osmotic shock, facilitating cell penetrance by peptides (Taiwo and Levine, 1989; Khasar et al., 1995).

Mechanical threshold in the skin

Nociceptive thresholds were quantified in lightly restrained rats using an Ugo Basile Algesymeter (Stoelting, IL), which applies linearly increasing mechanical pressure to the dorsum of the hind paw. To decrease variability of measurements, the rats were trained for the paw-withdrawal reflex test at 5-minute intervals for 1 hour each day for a period of 3 days (Taiwo et al., 1989). Mechanical nociceptive thresholds were determined both before and 10, 15, and 20 minutes after the intradermal injection of epinephrine or PGE2, in a volume of 2.5 µl. For each dose of an agent, the mean of the nociceptive threshold at the 3 time points was determined and the percentage change in nociceptive threshold calculated as (post-injection nociceptive threshold)-(pre-injection nociceptive threshold) / (pre-injection nociceptive threshold) × 100.

Mechanical threshold in the gastrocnemius muscle

Since fibromyalgia syndrome affects the muscles, we tested whether there is increased sensitivity to immune mediator, PGE2 in muscle in the sound stressed rats compared to non-stressed controls. Mechanical nociceptive threshold in the gastrocnemius muscle was quantified using a digital force transducer (Chatillon, Model DFI2, Amtek Inc., Largo, FL) (Dina et al., 2008). Rats were restrained in a vented tubular Plexiglas® holder with slated openings on the side that allow easy access to the hind limb. A 6-mm diameter probe attached to the transducer was applied to the gastrocnemius muscle to deliver an increasing compression force, and the nociceptive threshold defined as the force, in Newtons (N), at which the rat withdrew its hind leg. Baseline withdrawal threshold was defined as the mean of 2 readings taken at 5-minute intervals before administration of test agent. Each experimental manipulation was performed on a separate group of rats.

Intramuscular injection of agents

Increasing doses of PGE2 (0.1–1000 ng) were administered cumulatively, at 25-minute intervals, in a volume of 10 µl, into the belly of the gastrocnemius muscle. Injection sites were marked on the skin, using an indelible pen, so that the same site on the muscle could be tested for mechanical nociceptive threshold. Mechanical nociceptive thresholds were measured again, 15 and 20 minutes after injection of PGE2.


Exposure to sound stress occurred over 4 days as initially described by Singh et al. (Singh et al., 1990) and previously used in our laboratory (Strausbaugh et al., 2003; Khasar et al., 2005). Animals were placed 3 per cage 25 cm from a speaker that emitted a 105-dB tone of mixed frequencies (11 to 19 kHz). Over a period of 30 minutes, rats were exposed to 5- or 10-second sound epochs each minute at random intervals during the minute. Sham stressed animals were placed in the sound chamber for 30 minutes, but without exposure to the sound stimulus. Following sound stress, rats were returned to the animal care facility in their home cages. Animals were exposed to the stressor on days 1, 3, and 4. This 4-day sound stress protocol is schematically illustrated in Figure 1A. Rats were used for nociceptive studies 24 hours, 7 or 14 days after the last exposure to sound stress.

Figure 1
Sound stress enhances hyperalgesia induced by prostaglandin E2 and epinephrine

Adrenal adrenalectomy and medullectomy

Rats were anesthetized with isoflurane (2.5% in 97.5% O2). The adrenal glands were located through bilateral incisions in the abdominal wall, and either removed (for adrenalectomy) or the capsules incised and the medullae removed (for adrenal medullectomy) (Wilkinson et al., 1981; Miao et al., 1992). Fascia were closed with suture and the skin closed with wound clips. Rats were provided with 0.45% saline to drink for the first 7 days following surgery. Adrenal medullectomy was performed at least 5 weeks prior to further experimental procedures to allow maximum recovery of HPA axis function (Wilkinson et al., 1981).

Measurement of epinephrine

One day prior to initiation of sound stress and 1, 7 or 14 days following its completion, whole blood samples (0.5 ml) were collected from the tail veins of anesthetized rats, and immediately placed on ice. Plasma was isolated by centrifugation and stored at −80°C until analysis. Catecholamines were extracted from plasma by alumina adsorption, and levels determined using high performance liquid chromatography with electrochemical detection (5200 electrochemical detector; ESA Inc, Chelmsford, MA) (Khasar et al., 2003a).

Measurement of Plasma Corticosterone

Rats were anesthetized in 2–3% isoflurane for at least 15 minutes prior to sample collection and all efforts were made to minimize stress prior to blood collection. Peripheral blood was collected via the tail vein in heparinized tubes, centrifuged, and plasma was stored at −80°C until analysis. Plasma corticosterone was measured (in duplicate) as per manufacturer’s instructions using a commercially available radioimmunoassay kit (Siemens Medical Solutions Diagnostics, Los Angeles, CA). Briefly, 50 µl of standard or plasma and 1 ml of 125I rat corticosterone were added to corticosterone Abcoated tubes and incubated for 2 hours. Tubes were decanted and each radiolabeled tube was read using a gamma counter for 1 minute. Corticosterone concentration is expressed as ng/ml.

Tissue preparation and immunohistochemistry

Naïve rats were deeply anesthetized with sodium pentobarbital (100 mg/kg, i.p.) and perfused through the left ventricle with ~35 ml phosphate buffered saline (PBS) containing heparin (100 U/ml), followed by 300 ml PBS (pH 7.3), containing 4 % paraformaldehyde (PFA, Sigma Aldrich, St. Louis, MO). L5 dorsal root ganglia (DRG) were immediately dissected out and sequentially transferred and stored in PBS containing 10, 20 and 30% sucrose at 4°C for 12 to 24 hours each. The DRG were embedded in Tissue tek (OCT compound, Electron microscopy sciences, Hatfield, PA) sectioned at 20 µm in a Hacker/Bright microtome cryostat (Hacker Instruments Inc., Fairfield, NJ) and mounted on microscope slides (Fisher scientific, Pittsburgh, PA).

Prior to antigen detection, tissue sections were rinsed with PBS, blocked and permeabilized for 1 hour at room temperature with PBS containing 10% normal goat serum (Jackson Immunoreasearch Laboratories, West Grove, PA) and 0.3% Triton X-100. Glucocorticoid-receptor (GCR) expression was revealed by a 12-hour incubation with a 1:500 dilution of the anti GCR antibody (Santa Cruz Biotechnology, Inc., Santa Cruz, CA) with PBS containing 10% normal goat serum (antibody dilution buffer) followed by three washing steps with PBS containing 0.3% Triton X-100 (for 10 minutes each) and a 2-hour incubation with an 1:500 dilution of an Alexa Fluor 594 conjugated anti-rabbit antibody (Invitrogen, Carlsbad, CA) in antibody dilution buffer. Tissue sections were washed three times with PBS containing 0.3% Triton X-100 and mounted with Fluoromount G (Southern Biotechnology Assoc., Inc., Birmingham, AL).

Oligodeoxynucleotide antisense

The glucocorticoid receptor antisense oligodeoxynucleotide (ODN) sequence: 5’- TGG AGT CCA TTG GCA AAT -3’ was directed against the translation initiation site of the rat mRNA ((Engelmann et al., 1998); GenBank accession number Y12264). The mismatch ODN sequence was designed by mismatching 5 bases ((Engelmann et al., 1998); denoted here by bold face type) of the antisense sequence, 5’- TGA AGT TCA GTG TCA ACT -3’. A BLAST search was performed to check for the theoretical specificity and activity of the sequences, to confirm that they were not homologous to other sequences in the rat. ODNs were reconstituted in nuclease-free 0.9% NaCl. ODN was injected at a dose of 2 µg/µl in 20 µl. As described previously (Alessandri-Haber et al., 2003; Alessandri-Haber et al., 2004; Dina et al., 2005; Malik-Hall et al., 2005), rats were anesthetized with 2.5% isoflurane (97.5% O2), a 30-gauge needle was inserted into the subarachnoid space on the midline between the L4 and L5 vertebrae, and the ODN injected at 1 µl/sec by micro-syringe. It has been shown that intrathecal administration of ODNs provides an effective mode of delivery of these molecules to the DRG neurons in vivo (Lai et al., 2002). Rats received injections of corticosterone (1 µg/2.5 µl, intradermally) into one hind paw on the sixth day of glucocorticoid receptor ODN treatment. In stressed animals, ODN treatment was initiated on the same day as sound stress and continued for 13 days after sound stress. In either case, nociceptive testing was performed the day after the last ODN treatment.

Chronic administration of epinephrine and corticosterone

Chronic administration of stress levels of epinephrine was performed by implanting Alzet® mini-osmotic pumps (model 2004; Durect, Cupertino, CA) filled with epinephrine, subcutaneously in the interscapular region to deliver epinephrine at the rate of 5.4 µg/0.25 µl/h (Khasar et al., 2005). Rats were anesthetized with isoflurane to insert implants. To produce levels of circulating corticosterone that mimic the time course of corticosterone levels induced by the sound stress protocol, a 100-mg corticosterone pellets (Akana et al., 1985) were implanted into adrenalectomized rats (to produce the normal baseline corticosterone level) and corticosterone (25 mg/kg i.p. (Kavushansky and Richter-Levin, 2006)) injected on days 1, 3, and 4 (to mimic the sound stress-induced transient increases in corticosterone). Nociceptive threshold was measured in rats with corticosterone and epinephrine implants 14 days after the last injection of corticosterone or vehicle.

Western blots

In anesthetized rat, the saphenous nerve was ligated with silk surgical suture (4-0) 1 cm above its knee-level bifurcation. On the fourth day following initiation of ODN treatment, a 5-mm section of saphenous nerve proximal to the ligation was harvested. Nerves were homogenized in RIPA buffer with 2X protease inhibitor by probe sonication. Cell lysates were then run on 7.5% polyacrylamide gels and transferred to the membrane. The membrane was probed with M-20 anti-glucocorticoid receptor antibody (raised in the rabbit, 1:200; Santa Cruz Biotechnology, Inc., Santa Cruz, CA) followed by incubation with horseradish peroxidase-conjugated goat anti-mouse IgG (1:5000; Pierce Biotechnology, Inc., Rockford, IL). The specificity of the anti-glucocorticoid receptor antibody was controlled for by using the corresponding blocking peptide. The antibody only detected 3 bands, one intense band at the expected size of 90 KDa, and two much fainter bands around 65 and 120 KDa. The band at the expected size was used for our statistical quantification. To normalize the amount of sample loaded, affinity purified mouse monoclonal anti-GAPDH antibody (1:5000; Abcam Inc., Cambridge, MA) was used, followed by horseradish peroxidase-conjugated goat anti-mouse IgG (1:5,000; Pierce). Membranes were incubated with enhanced chemiluminescence reagents (SuperSignal West solution; Pierce) and images of the membrane were acquired with CHEMILMAGER chemiluminescence imaging system and analyzed with AlphaEaseFC software (Alpha Innotech, San Leandro, CA).

Statistical analysis

Group data are presented as mean ± SEM and analyzed using repeated measures analysis of variance (ANOVA), 2-way or 1-way ANOVA, or Student’s t-test, as appropriate. Where the overall ANOVA showed significant differences between the groups, Scheffe’s post hoc test was used to determine the pairs of groups that were different. The accepted level of significance was p < 0.05. The p-values for main effects are from ANOVA, and all subsequent p-values are from Scheffe’s post hoc tests, unless otherwise stated.


Stress causes a delayed-onset, long-lasting increase in hyperalgesia evoked by PGE2 or epinephrine

As we reported previously (Khasar et al., 2005), 24 hours following exposure to a 4-day protocol of unpredictable sound stress (Fig. 1A), the magnitude of hyperalgesia induced by an intradermal injection of PGE2 into the hind paw was unchanged (Fig. 1B). However, in the present study we tested for effects over a much longer time period, and found that 14 days following sound stress, a marked enhancement of PGE2 hyperalgesia became apparent (40% greater decrease in the threshold for mechanically-evoked paw withdrawal; Fig. 1B). This state of greater susceptibility to PGE2-induced hyperalgesia persisted without diminution for at least 21 days (Fig. 1B). The 4-day sound stress protocol alone did not significantly affect nociceptive threshold, measured 24 hours (sham sound stress, 107.9 ± 1.1 g (n = 30); sound stress, 107.9 ± 1.4 g (n = 30)) or 14 days (Sham sound, 105.9 ± 1.7 g (n = 42); sound stress 106.7 ± 1.1 g (n = 40)) after the last sound stress.

To determine if stress-induced enhancement is restricted to hyperalgesia elicited via prostaglandin receptors, we tested the effect of sound stress on hyperalgesia elicited by the pronociceptive mediator, epinephrine. Epinephrine produces mechanical hyperalgesia via G-protein-coupled β2-adrenergic receptors on nociceptors (Khasar et al., 1999). Twenty-four hours following the end of sound stress, epinephrine-induced hyperalgesia was attenuated (Fig. 1C). However, similar to the observed enhancement of PGE2-induced hyperalgesia, 14 days after stress, epinephrine hyperalgesia was significantly enhanced (Fig. 1C). This indicates that the effect of stress is not limited to prostaglandin receptor signaling, but may be generalized to other G-protein-coupled receptors. Importantly, the concentration of immune mediator needed to induce hyperalgesia following stress is 1 to 1.5 orders-of-magnitude less than that required in the non-stressed control condition (Fig. 1B, C).

In the gastrocnemius muscle, 14 days following sound stress, PGE2 hyperalgesia was significantly enhanced compared to that in the non-stressed control group (Fig. 1D, p < 0.001).

Plasma glucocorticoids contribute to the effect of stress

We previously showed that plasma levels of glucocorticoids are increased by our unpredictable sound stress protocol (Strausbaugh et al., 2003), and others have shown that sensory neurons express glucocorticoid receptors (DeLeon et al., 1994). Therefore, we assessed the role of glucocorticoids in the stress-induced enhancement of hyperalgesia evoked by immune mediators, using a glucocorticoid receptor antagonist and intrathecal administration of oligodeoxynucleotide (ODN) antisense to the glucocorticoid receptor. As shown in Figure 2A, daily subcutaneous administration of the glucocorticoid receptor antagonist, mifepristone, during the period of sound stress exposure, and 13 days thereafter, prevented the stress-induced enhancement of epinephrine hyperalgesia compared to mifepristone vehicle treated group (p < 0.001), while not affecting baseline nociceptive thresholds (mifepristone vehicle, 107.5 ± 4.4 g (n = 4); mifepristone, 107.8 ± 2.7 (n = 12)).

Figure 2
The effect of stress is mediated by glucocorticoid receptors in sensory neurons

To determine if glucocorticoids act directly on sensory nerve fibers to enhance epinephrine-and PGE2-induced hyperalgesia, we tested if the stress-induced enhancement of epinephrine hyperalgesia is blocked by intrathecally injected ODN antisense to the glucocorticoid receptor. As shown in Figure 2B, sensory neurons express glucocorticoid receptors. As shown in Figure 2C, expression of glucocorticoid receptor protein was reduced by 33 ± 8% in the saphenous nerve in antisense- versus mismatch-treated rats. As shown in Figure 2D, glucocorticoid receptor antisense treatment eliminated the stress-induced enhancement of epinephrine hyperalgesia (tested 14 days after the last exposure to sound stress; p = 0.001, n = 12).

We determined if elevated glucocorticoids alone, acting locally at the site of nociceptive testing, are sufficient to mimic the pronociceptive effect of stress. As shown in Figure 3A, intradermal injection of corticosterone (Cort, 1 µg in 2.5 µl) at the site of mechanical nociceptive testing on the dorsum of the hind paw, increased epinephrine-induced hyperalgesia (p = 0.004 at 24 hours, p < 0.005 at 7 days). While the glucocorticoid-induced enhancement of epinephrine hyperalgesia was of more rapid onset (detectable within 24 hr) than that induced by stress, it was similarly long-lasting. Corticosterone injection had no effect on the baseline nociceptive threshold after 24 hours (Cort, 110.4 ± 1.7; vehicle, 111.3 ± 2.1 (n = 22 each)) or 7 days (cort, 107.3 ± 1.8; vehicle, 111.0 ± 1.7 (n = 12 each)), and epinephrine hyperalgesia in the contralateral, vehicle-treated paw was not enhanced (Fig. 3A). Local intradermal corticosterone treatment in sound-stressed rats did not further enhance epinephrine-evoked hyperalgesia (Fig. 3B). The increase in the hyperalgesic response produced by local administration of glucocorticoids was not limited to epinephrine-evoked hyperalgesia; it also enhanced PGE2 hyperalgesia (Fig. 4A).

Figure 3
Corticosterone mimics the ability of stress to enhance epinephrine hyperalgesia
Figure 4
Corticosterone-enhanced hyperalgesia is mediated by glucocorticoid receptors on sensory neurons

To address the possibility of an indirect effect of locally injected glucocorticoids via non-neuronal cells in the skin, antisense ODN was intrathecally administered to reduce expression of glucocorticoid receptor in primary afferents selectively among all other cells in the skin. Intrathecal administration of ODN antisense to glucocorticoid receptor blocked the enhancement of PGE2-evoked hyperalgesia induced by intradermal administration of corticosterone, compared to the effect of corticosterone in the control group (Fig. 4A, p = 0.03). As shown in Figure 4B, ODN antisense, but not mismatch, to the glucocorticoid receptor also eliminated the enhancement of epinephrine-induced hyperalgesia following intradermal corticosterone treatment (p = 0.035), but baseline nociceptive threshold was unaltered (mismatch 109.0 ± 2.0 g, vs antisense 107.2 ± 1.8 g (p > 0.05, n = 12 each)). Epinephrine-induced hyperalgesia in the vehicle-treated (contralateral) paw of control rats was not different from that of the antisense ODN-treated group (Fig. 4B, p > 0.05). Thus, we conclude that the action of corticosterone in the skin to enhance immune mediator-evoked hyperalgesia occurs directly on the peripheral nerve endings of primary afferents, presumably enhancing sensitization of nociceptors by the immune mediator.

Plasma epinephrine augments the glucocorticoid-mediated effect

To investigate the contribution of circulating epinephrine in stress-induced hyperalgesia, we first tested if sound stress does, in fact, elevate the serum level of epinephrine. Plasma levels of epinephrine were significantly increased 24 hours after the final sound stress exposure compared to those in sham-stressed rats (Table 1), and plasma epinephrine levels were still significantly elevated 21 days following sound stress (Table 1).

Table 1
Plasma epinephrine and corticosterone levels in sound stressed male Sprague Dawley rats.

As shown in Figure 5A, adrenal medullectomy was performed in two groups of rats and one group stressed, to test if release of epinephrine from the adrenal medulla is necessary for stress-induced enhancement of hyperalgesia evoked by immune mediators. In adrenal-medullectomized rats, hyperalgesia evoked by intradermal injection of epinephrine was not significantly different in stressed compared with non-stressed animals (p > 0.05), suggesting that both glucocorticoids and epinephrine are necessary for the initiation of the process leading to enhancement of hyperalgesia later on.

Figure 5
Epinephrine augments the glucocorticoid-mediated enhancement of epinephrine hyperalgesia

We performed further experiments to examine the influence of sympatho-adrenal-released epinephrine on the glucocorticoid-mediated enhancement of immune mediator hyperalgesia. First, rats were adrenalectomized to eliminate the release of both epinephrine and glucocorticoids by the sympathoadrenal and HPA axes, respectively. Corticosterone (100 mg) fused pellets as well as epinephrine-containing osmotic mini-pumps were implanted in adrenalectomized rats. In addition to the implants, a subgroup of the rats was given 3 injections of corticosterone (25 mg/kg body weight, i.p., (Kavushansky and Richter-Levin, 2006)) over a 4-day period, on days 1, 3 and 4, to produce stress levels of corticosterone, similar to the sound stress protocol. One hour after the last injection of corticosterone, plasma levels were (640.5 ± 103.5 ng/ml (n = 6)). Plasma levels for corticosterone 1 hour after sound stress in the current study were (268.3 ± 25.9 ng/ml (n = 12; Table 1)). As shown in Figure 5B, rats exposed to stress plasma levels of corticosterone for 3 days and epinephrine for 14 days exhibited an enhanced epinephrine-evoked compared to either naïve or rats injected with corticosterone vehicle (p < 0.001).

Stress alters intracellular signaling pathways in nociceptors

We hypothesized that the initial decrease of sensitivity to intradermal epinephrine (measured 24 hours after stress) reflects desensitization of β2-adrenergic receptors in response to the high plasma levels of epinephrine. However, the enhancement of epinephrine hyperalgesia, first observed 14 days after sound stress, which also occurs in the presence of sustained elevation of plasma epinephrine, would seem to require some other mechanistic change in the signaling pathways underlying PGE2- and epinephrine-induced hyperalgesia. Therefore, we tested the hypothesis that the enhancement of epinephrine-induced hyperalgesia involves a change in intracellular signaling pathways by which epinephrine sensitizes nociceptors.

In cardiac cells, chronic exposure to catecholamines can induce a switch in the a subunit of the G-protein complex associated with β2-adrenergic receptors, from αs to αi (Gs to Gi/o) (Lamba and Abraham, 2000). To test for a similar change in intracellular signaling pathways in cutaneous nociceptors caused by stress, we assayed the sensitivity of epinephrine-evoked hyperalgesia to pertussis toxin, a Gi/o protein inhibitor. As shown in Figure 6A, in non-stressed rats, approximately 50% of epinephrine-induced decrease in paw withdrawal threshold was blocked by pertussis toxin (100 ng in 2.5 ml intradermal); following sound stress, however, approximately 80% of the epinephrine-induced hyperalgesia became sensitive to this dose of pertussis toxin (p < 0.001).

Figure 6
Adrenal medulla is required for stress-induced enhancement of hyperalgesia and pertussis-toxin sensitivity

To determine if the stress-induced increase in signaling via Gi/o is due to release of epinephrine by the sympathoadrenal stress axis, we performed adrenal medullectomy to eliminate the effect of neuroendocrine epinephrine. Hyperalgesia evoked by intradermal injection of epinephrine was not inhibited by pertussis toxin in both stressed as well as non-stressed adrenal-medullectomized animals (Fig. 6B). Thus, even levels of adrenal-derived epinephrine in the systemic circulation affect the G-protein dependence of signaling pathways by which epinephrine produces hyperalgesia, with basal levels of epinephrine maintaining approximately 50% dependence on Gi/o and stress levels markedly increasing this dependence to 80%.

To determine if the stress-induced enhancement of PGE2 hyperalgesia also involves a similar switch to Gi/o–dependent mechanisms, we assayed the effect of pertussis toxin on PGE2 hyperalgesia. In sham-stressed control rats, pertussis toxin actually enhances PGE2-evoked hyperalgesia (Fig. 6C) (a result similar to that previously observed in naive rats, presumably due to the toxin attenuating some baseline inhibition of adenylyl cyclase by Gi/o (Taiwo and Levine, 1991)). After exposure to sound stress, however, pertussis toxin no longer enhanced PGE2 hyperalgesia, rather it significantly inhibited it (Fig. 6C, p = 0.002). This suggests that, as for β2-adrenergic receptors, sound stress induces a switch in G-protein signaling, enhancing a hyperalgesic, pertussis toxin-sensitive pathway linked to PGE2 receptors. It also suggests that signaling downstream from Gi/o switches from predominantly inhibiting adenylyl cyclase to an alternative pathway that sensitizes primary afferent nociceptors.

We have shown previously that in hyperalgesic priming, a state induced by prior inflammatory lesion, PGE2 hyperalgesia switches to PKCε-dependent mechanisms (Aley et al., 2000). Therefore we investigated whether stress-enhanced PGE2 hyperalgesia, in addition to switching to a Gi/o–dependent pathway, also gains a new linkage to PKCε. An inhibitor of PKCε was administered intradermally before administration of PGE2 at the same site. The PKCε translocation inhibitor peptide had no effect on PGE2-evoked hyperalgesia in sham-stressed rats, but it abolished the enhancement of PGE2 hyperalgesia caused by sound stress (Fig. 6D). This reduction in stress-induced enhancement of PGE2 hyperalgesia was similar in magnitude to the reduction caused by the Gi/o inhibitor, pertussis toxin.


We have identified a stress-induced prolonged enhancement of mechanical hyperalgesia mediated by neuroendocrine release of both glucocorticoids and catecholamines, which act by modifying signaling pathways in primary afferent nociceptors. This phenomenon may explain the prominent influence of stress in chronic generalized pain syndromes, such as fibromyalgia (Nilsen et al., 2007a) and irritable bowel syndrome (Bach et al., 2006), as well as in other illnesses, like post-traumatic stress disorder (Otis et al., 2003) and depression (Blackburn-Munro and Blackburn-Munro, 2001)

The pro-nociceptive effect of stress that we observed differs markedly from previously reported models of “stress-induced hyperalgesia” (Imbe et al., 2006). Most notably, our protocol of unpredictable sound stress by itself did not cause hyperalgesia (there was no change in the baseline nociceptive threshold following stress). Instead, the effect of stress was only manifest as an enhancement of the hyperalgesia evoked by immune mediators. Specifically, stress caused a leftward shift in the dose-response relation for immune mediator hyperalgesia by 1.5 orders-of-magnitude. Therefore, we speculate that in disease states associated with increased cytokines, even at low levels normally insufficient to produce hyperalgesia, stress (also at levels normally insufficient to produce hyperalgesia) could precipitate a long-lasting hyperalgesic condition.

Our model also differs from other models of stress-induced hyperalgesia in the very long delay (7 to 14 days) between exposure to stress and the expression of enhanced immune mediator hyperalgesia. The present results do not implicate a mechanism for the delay, but it may be instructive to note that a similar delay in enhancement of hyperalgesia occurs in rats after subdiaphragmatic vagotomy (Khasar et al., 1998b), an effect also dependent on neuroendocrine stress axes (Khasar et al., 1998a; Khasar et al., 2003b). Furthermore, whereas hyperalgesia is a component of inflammation, in our earlier studies on the effect of non-habituating sound stress in a model of inflammation, bradykinin-induced plasma extravasation from the knee joint, we found no evidence of the involvement of the HPA axis in sound stress-induced suppression of plasma extravasation (Strausbaugh et al., 2003). This lack of involvement of the HPA axis suggests differences in the mechanisms mediating the effect of non-habituating sound stress in the inflammation model (Strausbaugh et al., 2003) and enhancement of immune mediator hyperalgesia in the current study.

Previous investigations have implicated changes in central nervous system circuitry as underlying mechanisms of stress-induced hyperalgesia (Imbe et al., 2006). In contrast, while the stress-induced hypersensitivity to immune mediator hyperalgesia that we observed could potentially also involve changes in the central nervous system, it clearly depends on changes within the primary afferent nociceptor. Thus, the effect of stress could be mimicked by local administration of corticosterone in the skin, an effect attenuated by intrathecal administration of antisense to the glucocorticoid receptor. These results suggest that glucocorticoids act at receptors located on sensory nerve fibers to participate in stress-enhancement of epinephrine hyperalgesia.

Within nociceptors, stress caused changes in second messenger signaling pathways that are activated by hyperalgesic mediators. This was indicated by a shift toward greater dependence on Gi/o signaling in nociceptors of stressed rats. This shift seems related to a previously reported shift in β2-adrenergic receptor signaling from Gs to Gi/o caused by chronic exposure to epinephrine (Lamba and Abraham, 2000). However, that shift involves -an increased role for Gi/o inhibition of adenylyl cyclase, which would be expected to attenuate hyperalgesia (because elevation of cyclic AMP causes hyperalgesia). In contrast, our finding of enhanced hyperalgesia associated with a Gs-to-Gi/o switch suggests that such inhibition of adenylyl cyclase is not the dominant effect of the stress-induced signaling switch in nociceptors. In this case, chronic exposure to elevated levels of epinephrine in peripheral tissue appears to induce novel coupling via Gi/o to second messenger pathways that produce hyperalgesia. Thus, stress caused a switch in downstream signaling pathways by which PGE2 produces hyperalgesia, from its usual dependence on a cyclic AMP/protein kinase A pathway (Levine and Reichling, 2005), to a novel additional dependence on PKCε. The emergence of PKCε dependence may be particularly significant in view of our previous finding that PKCε plays a crucial role in “hyperalgesic priming,” a model of inflammation-induced chronic enhancement of hyperalgesia (Aley et al., 2000).

While the switch in G-protein coupled receptor signaling, from Gs to Gi, and the signaling from Gi to PKCε have not been reported previously in neurons, such mechanisms are well established in another excitable tissue, the myocardium. Thus, in heart muscle cells, β2-adrenegic receptor signaling can switch from Gs to Gi (Daaka et al., 1997; Hasseldine et al., 2003; Hill and Baker, 2003; Magocsi et al., 2007), and signaling from Gi to mediators known to be upstream of PKCε Herrlich et al., 1996; Daaka et al., 1997; Steinberg, 1999; Mackay and Mochly-Rosen, 2001; Pavoine et al., 2003; Pavoine and Defer, 2005) or to PKCε (Fraser et al., 2000; Otani et al., 2003; Paruchuri and Sjolander, 2003) have been reported.

Conceivably, the interaction between the hypothalamo-pituitary-adrenal and sympathoadrenal axes that we observed occurs within the primary afferent nociceptor, potentially involving known interactions between glucocorticoid and β2-adrenergic receptor signaling pathways. Thus, epinephrine can increase the efficacy of glucocorticoid receptors, via the βγ phosphokinase pathway (Schmidt et al., 2001) which can contribute to immune mediator hyperalgesia (Khasar et al., 1995). Conversely, glucocorticoids can increase the expression of β2-adrenergic receptors (Fraser and Venter, 1980) as well as enhance their downstream second messenger signaling (Gs to adenylate cyclase to cAMP) (Aksoy et al., 2002).

Our findings may also provide an explanation for why generalized pain syndromes such as secondary fibromyalgia and irritable bowel syndrome occur at higher frequencies in diseases associated with the production of immune mediators (for example, interleukin-1β which signals through production of PGE2 (Hori et al., 2000)). Such cytokine production has been noted in several chronic conditions co-morbid with generalized pain, including rheumatoid arthritis (Petrovic-Rackov and Pejnovic, 2006) and inflammatory bowel disease (Raddatz et al., 2005), as well as in depression and in self-reported poor health (Lekander et al., 2004; Calcagni and Elenkov, 2006).

Our demonstration of enhanced hyperalgesia following administration of glucocorticoids may further explain why fibromyalgia patients treated with glucocorticoids have been reported to show a trend towards worsened outcomes (Clark et al., 1985), and why patients receiving steroid injections for arthritis sometimes report a period of increased pain (Wollstein et al., 2007). Furthermore, it is known that genes that modulate β-adrenergic receptor-mediated signaling, such as catechol-O-methyltransferase (COMT), affect pain sensitivity in vivo (Nackley et al., 2007), and genetic variants of COMT are predictive of development of temporamandibular disorder, a generalized pain syndrome (Diatchenko et al., 2005). Also consistent with this idea, the β-adrenergic receptor antagonist pindolol has shown promise in a preliminary trial for fibromyalgia (Wood et al., 2005).

In summary, our work provides a potential mechanistic explanation for the pathophysiology of enhanced pain and disease flairs in stress-sensitive pain syndromes such as fibromyalgia, irritable bowel syndrome, interstitial cystitis, and rheumatoid arthritis. Importantly, our findings provide testable hypotheses for interventions in people with generalized pain syndromes. Specifically, elucidating the complex time-dependent interactions between the HPA and sympathoadrenal axes including the switch in second messenger signaling for pronociceptive immune mediators in the primary afferent nociceptor may identify specific targets for the development of novel therapeutic strategies.


This work was funded by NIH grant AR048821.


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