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Anesth Analg 2004;98:723-729 © 2004 International Anesthesia Research Society doi: 10.1213/01.ANE.0000099369.23397.D7
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Abstract |
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The functional integrity of the immune system is essential for peripheral antinociception. Previous studies have demonstrated that immune cells elicit potent antinociception in inflamed tissues and that corticotropin-releasing factor-induced antinociception is significantly inhibited in animals that have undergone cyclosporin A (CsA)-induced immunosuppression. In this study, we examined the effect of a single bolus of CsA on inflammatory nociception. CsA-treated rats had substantially increased nociception compared with nonimmunosuppressed rats, consistent with a reduction in circulating and infiltrating lymphocytes. Furthermore, CsA-treated rats had inhibition of corticotropin-releasing factor-induced immune-derived antinociception, which was dose-dependently reversed by IV injection of concanavalin A-activated donor lymphocytes (1.0–7.0 x 106 cells/0.1 mL). In conclusion, our findings provided further evidence that opioid-containing immune cells are essential for peripheral analgesia. It is evident from these findings that control of inflammatory pain relies heavily on a functioning immune system.
IMPLICATIONS: The immune system not only contributes to inflammation, but also provides localized analgesia. A depleted immune system results in a reduction of immune-derived analgesia and a potentiation of inflammatory pain. Donor activated lymphocytes reverse these effects, highlighting the importance of a functional immune system in inflammatory pain.
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Introduction |
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Several studies have highlighted the important link between the immune system and peripheral opioid antinociception (1–3). Immune cells such as T and B lymphocytes, neutrophils, eosinophils, monocytes, and macrophages can express, synthesize, and contain opioid peptides and, in addition, deliver these active opioid peptides to inflamed tissues (4–6). Once released, these opioid peptides selectively bind to opioid receptors on sensory nerve terminals to decrease neuronal excitability, inhibit the release of proinflammatory factors, and reduce nociception (7,8). More importantly, localized administration of exogenous interleukin-1ß (IL-1ß) and corticotropin-releasing factor (CRF) has been shown to elicit potent antinociception in Freund’s complete adjuvant (FCA)-induced hind paw inflammation of immunocompetent animals (7,9,10). These cytokines and their receptors are well known for their potent proinflammatory action (11). In this instance, antinociception is produced from the endogenous release of opioids from resident immune cells within inflamed tissue via activation of IL-1ß and CRF receptors on these immune cells (7,10). Furthermore, the administration of cyclosporin A (CsA) dose-dependently inhibits immune-derived antinociception (10). These findings have important implications for clinical conditions such as acquired immune deficiency syndrome, cancer, and organ transplantation, for which immunosuppression is a factor.
In this study, we investigated the effect of CsA-induced immunosuppression on baseline inflammatory nociception in an attempt to extend our previous studies (7). Then we investigated the role of donor concanavalin A (Con-A)-activated lymphocytes in the recovery of immune-derived antinociception lost with immunosuppression.
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Methods |
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Reagents included CRF, (3-[125I]iodotyrosyl27) ß-endorphin (END), rabbit anti-END, FCA, and gelatin (Sigma, Sydney, Australia); CsA (Affinity Bioreagents Inc., Golden, CO); Con-A (Sigma-Aldrich Pty. Ltd., Castle Hill, New South Wales, Australia); isoflurane (Abbott Australasia Pty. Ltd., Kurnell, Australia); and bovine serum albumin (BSA) (Research Organics, Cleveland, OH). Doses were calculated as the free base, and drugs were dissolved in the following vehicles: 7:3 sterile water for injection and 95% alcohol for intraperitoneal (IP) CsA, sterile isotonic saline for CRF, and sterile water for injection for Con-A. Routes and volumes of drug administration were i.pl. 0.15 and 0.1 mL, respectively, and IV via lateral tail vein (100 µL) for Con-A lymphocytes.
Experiments were conducted in male Wistar rats (200–300 g) housed individually in cages lined with cat litter bedding and were provided food and water ad libitum. The study was approved by the Animal Experiment and Ethics Committee for the University of Queensland.
To induce inflammation, rats received 0.15-mL intraplantar (i.pl.) FCA injections into the right hind paw footpad under isoflurane anesthesia. The course of inflammation was monitored by paw volume (PV) by using a plethysmometer (Ugo Basile). The inflammation remained confined to the treated paw throughout the observation period. Immunosuppression was induced at Day 3–4 postinduction of inflammation.
Nociceptive thresholds in subjects were evaluated by using a standard Analgesy-Meter (Ugo Basile). Rats were gently restrained, and incremental pressure (maximum of 250 g) was applied onto the footpad of the hind paw. The pressure required to elicit paw withdrawal, the paw pressure threshold (PPT), was determined. The mean of three consecutive measurements separated by 10 s was recorded. The sequence was altered between the contralateral and inflamed paw to preclude order effects. Nociceptive readings were taken at baseline (before intraplantar (i.pl.) FCA injection) and 3–4 days post-FCA injection (before pretreatment with CsA or vehicle).
Lymphocytes were harvested from nontreated adult male Wistar rats. The procedure involves lightly anesthetizing rats with CO2/oxygen. The rats were then decapitated, and trunk blood was collected (7–15 mL) into tubes containing 2 mL of heparinized saline. Blood was diluted with an equal volume of Hanks’ balanced salt solution (HBSS) (1:1) and 4 mL layered over 3 mL of Lympholyte®-Mammal (Cedarlane Laboratories Ltd., Hornby, Canada). After centrifugation (2500g, 20 min, 24°C), the lymphocyte layer was carefully removed, resuspended in HBSS (1:1), and centrifuged (3500g, 10 min), and the supernatant was decanted. The pellet was resuspended in 1 mL of HBSS and centrifuged at 5000g for 5 min. The supernatant was decanted and the pellet reconstituted with 2–4 mL of HBSS. An aliquot (20 µL) of the suspension was removed for determination of cell numbers and viability (>95%) by using the trypan blue exclusion method.
The lymphocyte suspensions were placed in sterile well plates with Con-A (100 ng/mL for every 1–2 x 106 lymphocytes), sealed, and incubated overnight on a moderate-speed rotator at room temperature to induce lymphocyte activation. After lymphocyte activation, cell numbers were determined and cell viability was assessed. The lymphocyte suspensions were reconstituted with HBSS.
Radioimmunoassay (RIA) was performed to quantify the END content in nonstimulated and Con-A-stimulated lymphocytes and in inflamed and noninflamed paw tissues. Inflamed and noninflamed subcutaneous paw tissues were removed from the hind paws, placed in 0.32 M sucrose solution, and stored at -80°C for further processing.
On the day of assay, the frozen tissue or lymphocyte samples were thawed at 4°C, the wet mass was weighed, and samples were made up to 3–5 mL with pH 7.4 phosphate-buffered normal saline (PBS) and 0.3% bacitracin. Samples were then homogenized (1 min) and centrifuged at 5000g for 5 min. The supernatant was removed and diluted 1:100 for processing by the RIA method described below.
On Day 1 of RIA, tubes were prepared in duplicate containing 200 µL of END standards (100–10,000 pg/200 µL prepared by serial dilution from 1 µL/mL stock) or diluted samples. Anti-END antibody 100 µL (10 µL of product per milliliter of 2% BSA in PBS) was added to the tubes. Tubes were parafilmed and incubated overnight at 4°C. On Day 2 of RIA, 125I-END tracer (10,000 disintegrations per minute) was added to the tubes, and tubes were parafilmed and incubated overnight at 4°C. On Day 3 of RIA, the binding reaction was terminated by the addition of 1.5 mL of polyethylene glycol (33% polyethylene glycol and 0.13% glacial acetic acid) at 4°C. After the addition of the polyethylene glycol/glacial acetic acid solution, the radioactive tubes were centrifuged at 5000g for 20 min, and the tubes were allowed to drain for 12 h. The radioactive pellets were counted on a PerkinElmer gamma counter.
The colocalization of END+ and CD4+ cell distribution was examined in control and CsA-induced immunosuppressed animals for both inflamed and noninflamed paw tissues. Frozen stored hind paw tissue was embedded into OCT Compound 4583 (Tissue Tek®) and snap-frozen, and 10-µm coronal sections were cut by using a Leica cryostat. Sections were lifted onto gelatin-coated slides (Sigma, Sydney, Australia), and immunolabeling was performed as described previously (7). Hydrophobic wells were made around cryostat tissue sections with a PAP pen (Sigma, Sydney, Australia), tissue-fixed with acetone (10 s, 4°C), and then washed with PBS (pH 7.4, 10 min). To block nonspecific reactive groups, sections were incubated in 0.5% BSA-PBS for 30 min. Between each two incubation steps, the sections were washed three times with PBS for 15 min. Antibodies were applied to tissue sections and incubated in a humidity chamber for the following durations: rabbit anti-END at 1:800 PBS-BSA, 16 h at 4°C; anti-rabbit fluorescein-5-isothiocyanate (FITC)-conjugated secondary antibody at 1:200 PBS-BSA, 2 h at room temperature; and preconjugated monoclonal anti-CD4 with phycoerythrin at 1:250 PBS-BSA, 2 h at room temperature. Sections were coverslipped with the aid of a SlowFade® Antifade kit (Molecular Probes) and viewed with a high-speed digital fluorescence microscope with Meta-Morph® and Metafluor® imaging software. Specificity of anti-END was verified by preabsorption of the antibody with END (10 µM, 24 h, 4°C), followed by the same immunolabeling procedure as outlined previously.
The time-course effect of immunosuppression on inflammatory nociception was determined in animals with 3–4 days of post-FCA-induced inflammation. Nociception was determined prior to and at 6, 24, 48, 72, and 96 h after IP CsA (3 mg/mL) or IP vehicle (28.5% ethanol in water; 1 mL). Circulating lymphocyte numbers were determined in control (n = 3) and CsA-treated (n = 3) groups. END content was determined for inflamed and noninflamed paw tissues of control and CsA-treated groups.
To investigate whether bolus injection of Con-A-activated donor lymphocytes provides recovery of antinociceptive responses from direct injections of CRF, all treatment groups were pretreated with IP CsA (3 mg/mL), and 2 days after pretreatment, rats received tail vein injections of 1) HBSS (100 µL; n = 7), 2) donor nonactivated lymphocytes (2.58 x 106 cells per 100 µL; n = 4), or 3) donor activated lymphocytes (1.37, 2.58, or 6.60 x 106 cells per 100 µL; n = 6–8 per group). Nociceptive testing commenced 24 h after donor cell injection; PPT and PV were then taken before and 5 min after i.pl. CRF.
PPT values are given as raw values (mean ± SEM) and were analyzed by Mann-Whitney U-test for independent data and by Wilcoxon’s signed rank test for dependent data (P < 0.05). Analysis of variance (ANOVA) and a subsequent linear regression ANOVA were performed to test the zero slope hypotheses for dose curves.
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Results |
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Nociceptive thresholds (PPT; mean ± SEM) were significantly lower in inflamed than noninflamed paws of CsA and control groups at all time points (P < 0.05; Wilcoxon’s signed rank test; Fig. 1). Comparison between inflamed paws of control and CsA groups showed markedly lower PPT in the CsA group at 24, 48, 72, and 96 h (P < 0.05; ANOVA; Fig. 1), while PPTs were similar at 0 and 6 h after CsA (P > 0.05; ANOVA; data not shown). No changes were observed in the contralateral paw throughout any of the treatment paradigms.
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; n = 9) and CsA-treated (•; n = 13) groups remained significantly lower than for the contralateral noninflamed paws (
; n = 22) at all time points (P < 0.0001, Wilcoxon’s signed rank test). The paw pressure threshold for inflamed paws of the CsA-treated group was significantly lower than for inflamed paws of the vehicle-treated groups at 24, 48, 72, and 96 h (*P < 0.05; analysis of variance) but was not significantly different at 0 h.Lymphocyte counts in normal (n = 4) and CsA-treated (n = 4) rats revealed an approximately 2-fold reduction in circulating lymphocyte numbers in CsA-immunosuppressed rats over immunocompetent rats (CsA treated, 16.12 ± 1.81 x 106 cells; vehicle treated, 29.20 ± 5.80 x 106 lymphocytes). Swelling was confined to the right paw throughout the treatment period. Although PV was significantly larger in inflamed paws than noninflamed paws (P < 0.05; Wilcoxon’s signed rank test; Fig. 2), PV between inflamed paws of vehicle-treated (n = 11), CsA-treated (n = 9), and donor lymphocyte-treated (n = 14) groups were not significantly different (P > 0.05; Mann-Whitney U-test).
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; n = 14), and vehicle-treated (
; n = 9) rats were significantly larger than for noninflamed paws (Lymphocytes incubated overnight with Con-A, a T cell-activating mitogen, showed significantly higher levels of END content by RIA than unactivated lymphocytes (6.0 ± 2.0 µg/106 cells and 1.3 ± 0.7 µg/106 cells, respectively). More than 95% of lymphocytes were still viable after overnight incubation verified by the trypan blue exclusion method.
All cell treatment groups (0.00, 1.37, 2.58, and 6.60 x 10 cells/100 µL), except the control group, received CsA treatment. Nociceptive results (PPT; mean ± SEM) in inflamed paws before cell donation were similar for all treatment groups, including the control group (P > 0.05; Mann-Whitney U-test). However, a dose-dependent linear increase in PPT for inflamed paws (P < 0.05; linear regression ANOVA; Fig. 2) was observed 24 h after cell injection (before i.pl. CRF); the 6.60 x 106 lymphocyte group had values similar to the control group (52 ± 17 g and 59 ± 4 g, respectively). CRF injection into inflamed paws produced a further dose-dependent increase in antinociception for the 1.37, 2.58, and 6.60 x 106 lymphocyte groups and reached a plateau at 2.58 x 106 lymphocytes (P < 0.05; linear regression ANOVA). No antinociceptive effect for i.pl. CRF was observed in the 0.00 x 106 lymphocyte group (P > 0.05; Mann-Whitney U-test). CRF-induced antinociception for the 2.58 and 6.60 x 106 lymphocyte groups was similar to that for the non-CsA-treated group (PPT was 95 ± 5 g, 89 ± 14 g, and 89 ± 5 g, respectively, for the 2.58 and 6.60 x 106 lymphocyte and non-CsA groups; Fig. 3). The CsA treatment group that received unactivated lymphocytes (2.58 x 106 cells/100 µL) had PPT responses not significantly different from those of the CsA-treated group that received only HBBS instead of lymphocytes (PPT was 31 ± 8 g and 24 ± 3 g, respectively; P > 0.05; Mann-Whitney U-test).
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; n = 9) and the CsA-treated rats that received 6.60 x 106 donor lymphocytes (*P > 0.05). The i.pl. injection of CRF into the inflamed paws of treated rats induced a corresponding dose-dependent antinociception, and the plateau effect started at 2.58 x 106 lymphocytes (•; P < 0.05; linear regression ANOVA); this was consistent with the maximal antinociceptive effect from CRF in non-CsA-treated rats (
P > 0.05; n = 9). CRF-induced antinociception was not observed in the 0.00 x 106 (n = 9) donor lymphocyte group.No immunoreactivity for END or CD4 was observed in the noninflamed hind paw (data not shown). Inflamed hind paws from nonimmunosuppressed animals exhibited larger numbers of END+ (green) and CD4+ (red) immunoreactive cells colocalized (yellow) (Fig. 4A–C) than in the immunosuppressed rat paw tissue. Cell counts from immunosuppressed versus nonimmunosuppressed animals revealed a difference in numbers of immunolabeled END+/CD4+ (Fig. 5). The controls, in which the END antibody was preabsorbed with END peptide (10 µM), yielded no specific END immunolabeling to sections (data not shown).
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The END content of inflamed paw tissue from CsA-treated and untreated rats was determined by RIA. Consistent with immunohistochemical studies, CsA-treated rats’ hind paw tissue measured END content was less in the CsA-treated group compared with the control group (P < 0.05; Mann-Whitney U-test; Fig. 6).
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Discussion |
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END-containing immune cells migrate through inflamed tissue, where they release END and then travel to local lymph nodes depleted of END (6,7). Local proinflammatory factors such as CRF release END from immune cells in inflamed tissue providing potent localized antinociception (7). In this study, a bolus injection of CsA interrupted this process of recirculation of END-containing cells, leading to depleted numbers of END-containing lymphocytes in inflamed tissue, increased nociception in inflamed paws, and decreased antinociception from i.pl. CRF. Our previous studies in rat models of inflammatory nociception have demonstrated a reduction in immune-derived antinociception with immunosuppression produced from both x-radiation and CsA (10). Furthermore, injection of Con-A-activated lymphocytes from normal rats reversed the loss of i.pl. induced antinociception observed in immunosuppressed rats. Hence, reduced immune system function through immunosuppression contributes to increased nociception. This study is the first to demonstrate a link between increased inflammatory nociception and immunosuppression.
Bolus injections of IP CsA to rats with established inflammation (three days after FCA) decreased circulating lymphocytes and increased nociception in the inflamed paw for at least four days. Concurrent with the decrease in nociceptive thresholds in the inflamed paws of immunosuppressed rats was the corresponding reduction in antinociception provided by direct injection of CRF into the inflamed paw. Previous studies have shown that CsA-induced immunosuppression inhibits IL-1ß-, CRF-, and stress-induced antinociception (3,10). However, contrary to the findings of this study, no change was observed in nociceptive thresholds of inflamed paws with CsA treatment (3). Moreover, no baseline nociceptive threshold changes have been seen in MOR knockout mice with inflammatory hyperalgesia (12), although the presence of a low endogenous opioid tone has been well characterized in acute nociceptive responses (13). It is of consequence that opioid endogenous tone has been shown to be most evident in mice with multiple opioid receptor knockout and varies depending on the stimuli (13).
The most likely explanation for the observed reduction in CRF-induced antinociception is low lymphocyte infiltration into the site of injury because of the reduction in circulating lymphocyte numbers. This is supported by the twofold reduction in the circulating lymphocyte population in immunosuppressed rats in this study and our previous studies (7). Direct evidence for reduced efficacy for immune-derived endogenous antinociception can be attributed to the small numbers of END+/CD4+ cells in the inflamed tissue of immunocompromised rats and a reduction in the overall END content in the inflamed paw tissues of the CsA-treated rats. It is unlikely that this reduction in CRF-induced antinociception is due directly to a reduction in lymphocyte END content or the related effect on the release of END from these immune cells in vitro, because our previous studies have shown that CsA did not alter the content of END in lymphocytes (7).
There was no trend toward a reduction in PV in the inflamed paws of immunosuppressed rats. CsA has been shown to have potent antiinflammatory activity by inhibition of cell recruitment (14) and neutrophil function (15). Therefore, a gross change in swelling could not be clearly demonstrated with a single CsA injection, indicating that CsA had little antiinflammatory action in this instance.
Lymphocytes stimulated with Con-A, a mitogen selective for stimulating and activating T cells, have been shown in this study to contain higher levels of END than unstimulated lymphocytes. Consistent with these findings, Con-A-stimulated, but not nonstimulated, CD4+ thymocytes expressed proenkephalin A messenger RNA (16,17). Furthermore, in this study, only the Con-A-stimulated donor cells provided recovery of CRF-induced antinociception in CsA treated rats, highlighting the importance of cell activation in immune-derived opioid antinociception. The recovery of CRF-induced antinociception with the administration of activated donor cells shows that lymphocytes are capable of reentering the circulating lymphocyte pool, migrating to the site of inflammation, and eliciting potent antinociception. It is likely that 24 hours is sufficient for the migration of enough lymphocytes into the inflamed paw of the CsA-treated rat to provide antinociception. Indeed, radiolabeled donor lymphocytes have been shown to rapidly enter the recirculating lymphocyte pool and accumulate in lymph nodes and spleen within 24 hours (18). Furthermore, this study is not the first to investigate the use of donor cells containing opioid peptides for the treatment of nociception. Potent analgesia has been observed after the spinal implantation of END and catecholamine-containing chromaffin cells into neuropathic, arthritic, and cancer pain models (19–21). Donor lymphocytes from rats of the same species as the immunocompromised recipients produce no immunogenic responses (18,19). There was no evidence of rejection in the rats that received donor activated lymphocytes in this studies.
Nociceptive baselines for inflamed paws (i.e., before stimulation by i.pl. CRF) were dose-dependently improved after IV injection of donor activated lymphocytes. Moreover, baseline nociceptive thresholds in the group of rats receiving 6.60 x 106 cells were similar to the baseline scores of the non-CsA-treated group. This indicates that the donor cell numbers were large enough to produce recovery of CsA-depleted immune-derived antinociception such that baseline nociceptive scores were similar to those of inflamed paws without immunosuppression. Consistent with improved baseline nociceptive scores, immune-derived antinociception was also increased in a dose-dependent manner with injection of donor activated lymphocytes. CRF (i.pl.) injection into donor rat groups produced antinociception, and maximal responses were obtained for 2.58 x 106 cells/100 µL. The plateau effect on the dose-response curve for donor cells may illustrate either a maximal dose-response relationship for immune-derived opioids or perhaps even a limit of cell infiltration kinetics.
The immune system is undoubtedly a major source for proinflammatory (22,23) and antiinflammatory mediators (4,7). Indeed, these cells have been shown to produce cytokines that could in turn activate receptors on these cells, causing the release of opioids within inflamed tissue. Moreover, immunosuppression decreases inflammation and, consistent with this effect, reduces nociception (24). In contrast, inhibition of cell recruitment into inflamed tissues or CsA treatment reduced immune-derived antinociception (3,6,10). Taken together, these studies demonstrate the complexity of the role of immune cells in inflammation. However, in this study, immune cell deficiency resulted in enhanced inflammatory nociception, and replacement of a CsA-depleted pool of immune cells with Con-A-activated lymphocytes resulted in the recovery of CRF-induced antinociception. This study highlights the importance of immune-derived antinociception in immunocompromised individuals.
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Acknowledgments |
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Supported by the National Health and Medical Research Council, The University of Queensland Startup Grant.
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