Opioid Peptide–expressing Leukocytes: Identification, Recruitment, and Simultaneously Increasing Inhibition of Inflammatory Pain

Animal protocols were approved by the Animal Care Committee of the State of Berlin (Landesamt für Arbeitsschutz, Gesundheitsschutz und Technische Sicherheit (Berlin, Germany). Male Wistar rats weighing 150–200 g were injected with 150 μl FCA (Calbiochem, La Jolla, CA) in the right-side hind paw under brief halothane anesthesia. Rats developed an inflammation confined to the inoculated paw.

Tissue samples were harvested 2, 6, and 96 h after injection of FCA. Animals were killed and popliteal lymph nodes were dissected. Subcutaneous paw tissue was obtained from the plantar surface leaving the deep flexor tendon in situ . To obtain a single cell suspension for fluorescence activated cell staining (FACS) analysis, magnetic cell separation, and radioimmunoassay (RIA), lymph nodes and subcutaneous paw tissue were cut into 1- to 2-mm pieces. Fragments were digested for 1 h at 37°C with 10 ml Roswell Park Memorial Institute (RPMI) 1640 medium (Life Technologies, Paisley, Scotland)/1 g tissue containing 30 mg collagenase (Sigma, Deisenhofen, Germany), 10 mg hyaluronidase (Sigma), and 0.5 ml HEPES, 1 m (Sigma). The digested fragments were pressed through a 70-μm nylon filter (Pharmingen/Becton Dickinson, Heidelberg, Germany) to remove particles.

Single cell suspensions obtained from one or two paws were centrifuged, decanted, and fixed with 1% paraformaldehyde in phosphate-buffered saline (PBS) for 30 min at room temperature and washed with PBS. Cells were permeabilized with saponin buffer (0.5% saponin, 0.5% bovine serum albumin, 0.05% NaN³[all from Sigma]) and subsequently stained with 20 μg/ml 3E7 mouse monoclonal immunoglobulin G^2a(IgG^2a) antibody in saponin buffer for 30 min at room temperature. This antibody recognizes the pan-opioid sequence Tyr-Gly-Gly-Phe-Met at the N-terminus of opioid peptides. 10Afterwards, cells were incubated with the secondary rat anti-mouse IgG^2a+bphytoerythrin (PE)–conjugated monoclonal antibody (1.5 μg/ml) for 15 min at room temperature. Negative controls included the replacement of the primary antibody with an isotype matched irrelevant antibody (mouse IgG^2a) and omission of the secondary PE–conjugated antibody at the same concentration.

Subsequently, cells were labeled with fluorescein isocyanate (FITC) or Cy Chrome–conjugated surface markers to differentiate subpopulations of monocytes or macrophages, granulocytes, and T cells and then fixed in 1% paraformaldehyde in PBS. To calculate absolute numbers of cells per paw, the stained cell suspension was transferred to a TruCOUNT^®tube (Pharmingen/Becton Dickinson) containing a known number of fluorescent beads. FACS events from the fluorescent TruCOUNT^®beads and stained cells were collected simultaneously using the FACScan (Pharmingen/Becton Dickinson). Numbers of CD45⁺cells per tube were calculated in relation to the known number of fluorescent TruCOUNT^®beads and extrapolated for the whole paw. At least 10,000 FACS events were collected in the lymph node, and 30,000 FACS events in the paw. Data were analyzed using CellQuest software (Pharmingen/Becton Dickinson).

To verify the specificity of staining, exogenous END or a control peptide (adrenocorticotropic hormone [ACTH]) (both from Peninsula, Merseyside, United Kingdom) was added to the lymphocytes from an inflamed lymph node after 96 h of FCA treatment before staining with 20 μg/ml 3E7. Each peptide was added at four concentrations: equimolar to 3E7 concentration, 0.5, 1.5, and 2.0 m excess of the peptide. Amounts were calculated as follows: (1) 3E7 monoclonal antibody (IgG, MW = 150,000 Da), final concentration, 20 μg/ml = 0.132 nmol; (2) END peptide (MW = 3,464), equimolar 0.132 nmol × 3,464 = 462 ng/ml; and (3) ACTH peptide (MW = 4,579): equimolar 0.132 nmol × 4,579 = 610.8 ng/ml.

The following antibodies were used: mouse monoclonal anti–pan-opioid 3E7 (subtype IgG^2a; Gramsch Laboratories, Schwabhausen, Germany); mouse IgG^2aand rat anti-mouse IgG^2a+bPE-conjugated (both from Pharmingen/Becton Dickinson); mouse anti-rat CD3 FITC/PE (T-cell marker), mouse anti-rat granulocyte FITC (HIS48), mouse anti-rat CD45 Cy Chrome and streptavidin–conjugated mouse anti-rat CD45 (all hematopoetic cells); mouse anti-rat ED1 FITC (monocyte/macrophage marker) (Pharmingen/Becton Dickinson and Serotec, Oxford, United Kingdom).

To obtain a sufficient number of CD45⁺cells to measure END by RIA, cells from several paws had to be pooled. After 2 h of inflammation, 16 paws; after 4 h, 6 paws; and after 96 h, 3 paws were processed and measured together. Cell suspensions were stained with streptavidin-conjugated mouse anti-rat CD45 monoclonal antibody and subsequently labeled with avidin-coupled magnetic beads (Milteny Biotec, Bergisch-Gladbach, Germany) in PBS (0.5% bovine serum albumin [BSA], 2 mm EDTA). CD45⁺cells were isolated using a VS⁺column (Milteny Biotec). Purified CD45⁺cells were pelleted, lysed in 250 μl RIA buffer (0.1 m sodium phosphate, 0.05 m NaCl, 0.01 m NaN³, 0.1% BSA, and 0.1% Triton X-100) containing aprotinin (all from Sigma), and shock-frozen until further used. Purity of the cells was greater than 95% as measured by FACS after appropriate staining.

After the above-described cell separation, END peptide content in CD45⁺cells was measured by rat END RIA (Peninsula) as described previously. 9 

Rats were anesthetized with halothane 2, 6, and 96 h after FCA inoculation and perfused through the ascending aorta with 50 ml PBS and 400 ml Zamboni fixative (4% paraformaldehyde in PBS and 14% picric acid). Subcutaneous paw tissue was stained as previously described using a primary rabbit anti-rat END (1:1,000 dilution, Peninsula) and secondary biotinylated anti-rabbit antibody, Vectastain avidin-biotin-peroxidase complex, and 3,3′-diaminobenzidine as a substrate (all from Vector Laboratories, Burlingave, CA). 2 

Paw pressure testing was performed 2, 6, and 96 h after FCA injection as described previously. 4In brief, rats were handled before testing and gently restrained under paper wadding, and pressure was applied onto the dorsal surface of each hind paw. The pressure threshold (cut-off at 250 g) for paw withdrawal was determined using an algesiometer (Ugo Basile, Comerio, Italy) in three consecutive measurements separated by 10-s intervals to calculate mean baseline values. To test stress-induced opioid analgesia, rats were exposed to ice-cold water for 1 min (CWS) and gently dried, and pressure threshold values were obtained after 1 min as described previously. 4 

Data obtained at three time points (2, 6 and 96 h) were compared by analysis of variance (ANOVA) if the normality test and the equal variance test were passed. Otherwise, the Kruskal–Wallis ANOVA on ranks was used. Post hoc  multiple pairwise comparisons were performed by the Student–Newman–Keuls method. If group sizes were unequal, post hoc  comparisons were performed by Dunn test. Data of two cellular subpopulations (granulocytes vs.  monocytes) were compared by the t  test. If data were not normally distributed, the Mann–Whitney U test was used. A value of P < 0.05 was considered significant.

Immune cells isolated from the hind paw were pregated by staining with anti-CD45 monoclonal antibody to exclude debris and nonhematopoietic cells such as fibroblasts or lipocytes (fig. 1A). CD45⁺Cells were further differentiated into hematopoetic lineages. In early FCA inflammation (2 and 6 h) the majority of infiltrating cells were HIS48⁺granulocytes, whereas ED1⁺monocytes or macrophages were significantly less abundant (HIS48⁺granulocytes, 65–70%; ED1⁺monocytes, 24–38%[P < 0.05]) (figs. 1B and C). At 96 h of FCA inflammation, the percentage of HIS48⁺granulocytes decreased and ED1⁺monocytes or macrophages were the prominent cell type (35% HIS48⁺cells vs.  60% ED1⁺cells [P > 0.05, t  test]) (fig. 1D). The percentage of CD3⁺lymphocytes remained unchanged and relatively low (2–4%). The total number of infiltrating CD45⁺cells increased significantly over time (2.5-fold, 2 vs.  6 h, and threefold, 6 vs.  96 h [P < 0.001, Kruskal–Wallis;P < 0.05 at all pairwise comparisons by Student–Newman–Keuls]) (figs. 1B–D).

An intracellular stain for opioid peptides was established. Lymphocytes obtained from an inflamed lymph node were stained with 3E7 or an isotype matched control monoclonal antibody and quantified by flow cytometry (fig. 2A). Specificity of the staining was demonstrated by dose-dependent and significant inhibition of 3E7 staining by preincubation with END but not with ACTH (P < 0.001, ANOVA;post hoc  multiple comparisons by Student–Newman–Keuls:P < 0.05 for 1.5 and 2 m END vs.  without peptide competition [NONE] and 1.5 and 2 m END vs.  2 m ACTH;P > 0.05 for 2 m ACTH 2 vs.  NONE) (fig. 2B). The 3E7 staining was reduced by 60% at 2 m excess of exogenous END.

Single cell suspensions from paws obtained 2, 6, and 96 h after FCA-induced inflammation were double-stained for 3E7 and CD45 and analyzed by flow cytometry (fig. 3A). The absolute number of 3E7-positive CD45⁺cells increased significantly from 18,200 ± 4,000 cells at 2 h to 40,900 ± 8,100 cells at 6 h and to 102,300 ± 35,100 cells at 96 h after injection of FCA (P < 0.001 Kruskal–Wallis, P < 0.05 for all pairwise comparisons by Student–Newman–Keuls). The noninflamed paw was not analyzed because of the absence of CD45⁺cells in the FACS analysis.

To confirm these findings, histologic sections from the inflamed hindpaw were stained with polyclonal anti-END at 2, 6, and 96 h after FCA injection. Specificity of staining was demonstrated by preabsorption of the primary antibody with excess of rat END peptide (data not shown). The number of END-positive immune cells increased substantially with the duration of inflammation (fig. 3B).

To quantify the total END content in CD45⁺cells per paw at various intervals after FCA injection, cells were purified by magnetic beads and END content was measured by RIA. Before selection, CD45⁺cells accounted for 36% of total FACS events collected at 96 h (fig. 4A). After magnetic cell separation, positively selected cells were greater than 95% pure. The END content in purified CD45⁺cells increased in a time-dependent manner during the course of inflammation from 9.3 ± 2.1 pg/paw at 2 h to 17.3 ± 4.1 pg/paw at 6 h and to 36.9 ± 17.1 pg/paw at 96 h, confirming the results obtained by flow cytometry (P < 0.05, Kruskal–Wallis ANOVA on ranks, post hoc  pairwise comparison by Dunn test:P < 0.05 for 2 vs.  96 h, P > 0.05 for 2 vs.  6 h and 6 vs.  96 h) (fig. 4B).

To assess hematopoietic origin of opioid peptide–producing immune cells, triple-color flow cytometry was performed. All samples were stained with the CD45 and 3E7 monoclonal antibodies. For lineage assignment, cells were additionally labeled with HIS48 or ED1 monoclonal antibody, respectively. Stains were pregated on CD45⁺cells as shown in figure 2Aand subsequently analyzed for 3E7 expression and for hematopoietic lineage markers (fig. 5A). The 3E7⁺HIS48⁺granulocytes increased significantly during early inflammation, whereas minor and nonsignificant changes occurred between 6 and 96 h (fig. 5B) (P < 0.01, Kruskal–Wallis;post hoc  pairwise comparison by Student–Newman–Keuls:P < 0.05 for 2 vs.  96 h and 2 vs.  6 h, P > 0.05 for 6 vs.  96 h). In contrast, 3E7⁺ED1⁺cells increased significantly between all time points (P < 0.001, Kruskal–Wallis;P < 0.05 at all pairwise multiple comparisons comparison by Student–Newman–Keuls). However, although the initial increase in recruitment of monocytes or macrophages was small (1.8-fold, 2 vs . 6 h), a dramatic 4.4-fold increase in recruitment occurred later (6 vs.  96 h) (fig. 5C). Opioid peptide–producing cells were predominantly HIS48⁺granulocytes in early inflammation (at 2 h, 66% of total 3E7⁺cells, P > 0.05 by t  test [HIS48⁺vs.  ED1⁺]; at 6 h, 63%, P < 0.05 by t  test [HIS48⁺vs.  ED1⁺]). At later stages of inflammation the pattern changed, and the majority of 3E7⁺cells were ED1⁺monocytes or macrophages (at 96 h, 73% of total 3E7⁺cells, P < 0.05 by t  test [HIS48⁺vs.  ED1⁺]).

To examine the functional relevance of the increase in the number of opioid peptide–producing immune cells and in END content, endogenous analgesia was tested by CWS. The CWS-induced pressure threshold elevation increased significantly with the duration of inflammation (P < 0.01, ANOVA;post hoc  pairwise comparisons by Student–Newman–Keuls:P < 0.05 for 2 vs.  96 h and 6 vs.  96 h;P > 0.05 for 2 vs.  6 h (figs. 6A–C). Consistent with the absence of opioid-containing cells in noninflamed tissue, 4CWS-induced analgesia was never observed in the contralateral paw.

Our findings show that (1) the cell pattern after FCA inoculation of the rat paw is consistent with a typical nonspecific inflammatory reaction of the innate immune system; (2) the monoclonal antibody 3E7 produces specific intracellular staining and allows the quantification of opioid-producing cells in flow cytometry; (3) the number of opioid-producing cells, determined both by FACS and immunohistochemistry, increases in parallel with the END content; (4) opioid-containing cells are predominantly granulocytes during early and monocytes or macrophages during later stages; and (5) endogenous opioid pain control increases in parallel with cell recruitment and inflammation.

At early stages of inflammation (2 and 6 h), opioid peptides were mainly produced by HIS48⁺granulocytes, whereas ED1⁺monocytes or macrophages became the predominant opioid-containing cell type at later stages (96 h). CD3⁺T cells were detected in low numbers in the paw tissue during the first 4 days of this inflammation. This time-dependent pattern of immune cell recruitment to subcutaneous tissue after FCA injection is similar to the emergence of cells observed in other types of inflammation including infection by pathogens or wound healing. 11 

Importantly, our studies expand this concept in that granulocytes and monocytes are not only members of the frontline innate immunity, but they also produce and secrete opioid peptides to inhibit pain. Our findings are in agreement with prior studies examining opioid peptide production in immune cells. ENK production has been demonstrated in granulocytes, 12mononuclear cells, and T cells. 8END is detectable in a variety of cell types including ED1⁺monocytes or macrophages, 5lymphocytes, 9and mast cells. 4Furthermore, precursor mRNAs of END and ENK can be found within inflamed tissue by in situ  hybridization. 5Our previous study examined trafficking of END⁺immune cells in the circulating blood and the regional lymph nodes. 9It suggested that END⁺cells are of lymphocytic origin, migrate from the peripheral blood to the inflamed hind paw where they secrete opioid peptides, and, finally, leave the site of inflammation to the draining and inflamed lymph node. In contrast to the lymph node and the blood, opioid peptide–expressing cells in the paw are mainly of granulocytic or monocytic origin. Nonetheless, CD3⁺lymphocytes also contain opioid peptides and might be of importance in truly chronic inflammation.

To examine the functional relevance of opioid peptide–producing immune cells in pain control, we tested antinociceptive effects after a stressful stimulus (CWS). Previously, we extensively characterized this paradigm and showed that, on CWS, opioid peptides are released within the paw, then occupy opioid receptors on sensory neurons and thereby produce analgesia. 2,4,5In this study we found that intrinsic peripheral analgesia triggered by CWS correlated tightly with the number of 3E7⁺CD45⁺cells. To substantiate our newly developed triple-color flow cytometry, we showed that not only the number of 3E7⁺CD45⁺cells but also the END content in the inflamed paw increased in a time-dependent manner. The latter was quantified by radioimmunoassay, similar to previous studies. 6,9,13It seems that the END content in the inflamed paw mainly reflects the degree of migration of immune cells. At present, it is unknown whether immune cells increase their production of opioid peptides already in the circulation and then migrate to the site of inflammation or whether opioid peptide production in immune cells is primarily induced at the site of the inflammation.

Is pain control determined by the amount of opioid peptide at the site of inflammation or rather by opioid receptor expression? This question is particularly important because future therapeutic strategies in pain inhibition can be directed toward either opioid peptide–containing immune cells or neuronal opioid receptor expression. Earlier studies showed that opioid receptors are transported from the dorsal root ganglion to peripheral terminals of sensory neurons and that receptor transport is up-regulated by inflammation. 1In the current study we show that recruitment of more opioid peptide–containing immune cells led to more profound analgesia. This view is supported by a prior study in which recruitment of opioid peptide–producing immune cells was blocked by the selectin blocker fucoidin, resulting in substantially reduced analgesia. Importantly, intraplantar injection of the synthetic opioid agonist fentanyl induced an identical degree of analgesia in both control and fucoidin-treated rats, suggesting the presence of similar numbers of receptors in both cases. 2Therefore, we conclude that at the beginning of an inflammatory process intrinsic opioid analgesia is primarily dependent on the number of opioid-containing leukocytes rather than on the number of receptors.

In the current study we demonstrated that opioid-containing immune cells are HIS48⁺granulocytes during the early stages of inflammation, whereas ED1⁺monocytes or macrophages predominate later. These results predict that immunosuppression targeting granulocytes or monocytes should lead to a reduction of analgesia. Indeed, immunosuppression by whole-body irradiation 5or cyclosporine 4abolishes CWS-induced analgesia in FCA-treated rats. Similarly, immunomodulation can be achieved by selective blockage of cell migration. 2These three examples demonstrate that both suppression and modulation of the immune system have profound consequences for endogenous pain control. This is particularly important because manipulation of the immune system is a prime target in various diseases including sepsis, rheumatoid arthritis, and chemotherapy for cancer. Immunosuppression with inhibition of the function of macrophages in autoimmune disease or agranulocytosis induced by chemotherapy could have the unanticipated side effect of abolishment of endogenous pain control.

These considerations have clinical consequences: Peripheral opioid analgesia has been conclusively demonstrated in surgical patients and in patients with arthritis. 14Evidence for endogenous peripheral opioid peptide release under stressful conditions such as surgery was presented in a study showing that injection of naloxone after knee arthroscopy exacerbates pain and increases pain medication consumption. 3,15Our studies in the rat model suggest that opioid peptide-containing immune cells are recruited to sites of injury and inflammation as part of the innate immune reaction. These opioid peptides apparently mount a parallel defense against the accompanying pain response. Thus, it is conceivable that modulation of this peripheral opioid mechanism (e.g.,  by gene transfer of opioid peptide DNA) could be used in future therapeutic strategies to inhibit pain. Such strategies would have the advantage of being devoid of central opioid side effects such as sedation, nausea, dependence, or respiratory depression.

In summary, tissue insults result in the immediate recruitment of granulocytes and some monocytes that not only attempt to limit the extent of tissue destruction but also allow endogenous analgesia to occur. During the ongoing inflammation other cells such as monocytes, and probably T cells at later stages, are recruited that ensure ongoing endogenous opioid-mediated analgesia. Endogenous local analgesia after tissue destruction could be a principle of fundamental importance: On the one hand, pain at the site of damage signals the degree of injury and limits unnecessary usage. On the other hand, potent analgesia is mounted simultaneously so that the handicap of the injury can be temporarily ignored in periods of danger.

The authors thank Andreas Thiel, Ph.D., and Andreas Radbruch, Ph.D., Deutsches Rheumaforschungszentrum, Berlin, Germany, for technical advice regarding intracellular staining for flow cytometry.