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Published as doi: 10.1096/fj.06-6077fje.
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(The FASEB Journal. 2006;20:2627-2629.)
© 2006 FASEB

Pain control by CXCR2 ligands through Ca2+-regulated release of opioid peptides from polymorphonuclear cells

Heike L. Rittner*,1, Dominika Labuz*, Michael Schaefer{dagger}, Shaaban A. Mousa*, Stefan Schulz§, Michael Schäfer*, Christoph Stein* and Alexander Brack*

* Klinik für Anaesthesiologie und Operative Intensivmedizin and

{dagger} Institut für Pharmakologie, Charité–Universitätsmedizin Berlin, Campus Benjamin Franklin, Berlin, Germany; and

§ Department of Pharmacology, Otto-von-Guericke-University, Magdeburg, Germany

1Correspondence: Klinik für Anaesthesiologie und Operative Intensivmedizin, Charité–Universitätsmedizin Berlin, Campus Benjamin Franklin, Hindenburgdamm 30, D-12200 Berlin, Germany. E-mail: heike.rittner{at}charite.de

ABSTRACT

Leukocytes counteract inflammatory pain by releasing opioid peptides, which bind to opioid receptors on peripheral sensory neurons. In the early phase of inflammation, polymorphonuclear cells (PMN) are the major source of opioids. Their recruitment is governed by ligands at the chemokine receptor CXCR2. Here, we examined whether chemokines can also induce opioid peptide secretion from PMN and thus inhibit inflammatory pain. In rats with hindpaw inflammation, intraplantar injection of CXCL2/3, but not of the CXCR4 ligand CXCL12, elicited naloxone-reversible (i.e., opioid receptor mediated) mechanical and thermal analgesia, which was abolished by systemic PMN depletion. Both CXCR1/2- and CXCR4-ligands induced PMN chemotaxis, but only CXCR1/2 ligands triggered opioid release from human and rat PMN in vitro. This release was unaltered by extracellular Ca2+ chelation, was mimicked by thapsigargin and was blocked by inhibitors of the inositol 1,4,5-triphosphate receptor (IP3) and by intracellular Ca2+ chelation, indicating that it required Ca2+ from intracellular but not extracellular sources. Furthermore, release was partially reduced by phosphoinositol-3-kinase (PI3K) inhibitors. Adoptive transfer of allogenic PMN into PMN-depleted rats reconstituted CXCL2/3-induced analgesia, which was inhibited by prior ex vivo chelation of intracellular Ca2+. These findings demonstrate that, beyond cell recruitment, CXCR2 ligands induce Ca2+-regulated opioid release from PMN and thereby inhibit inflammatory pain in vivo.—Rittner, H. L., Labuz, D., Schaefer, M., Mousa, S. A., Schulz, S., Schäfer, M., Stein, C., and Brack, A. Pain control by CXCR2 ligands through Ca2+-regulated release of opioid peptides from polymorphonuclear cells.


Key Words: chemokine • analgesia • inflammation • secretion • neutrophil

INFLAMMATORY PAIN CAN be effectively controlled by peripheral opioid-mediated mechanisms. Endogenous opioid peptides such as Met-enkephalin (ENK) and ß -endorphin (END) are produced by infiltrating leukocytes, and ENK is also found in peripheral afferent neurons (1) . Opioid peptides are released from leukocytes on stimulation, bind to µ (MOR) and {delta} (DOR) opioid receptors on peripheral sensory neurons and counteract pain (2 , 3) . Opioid-containing leukocytes play an essential role in pain control since it is abolished by immunosuppression or leukocyte depletion (4 , 5) . In humans, pain after surgery is significantly intensified when this mechanism is interrupted following a local injection of the opioid receptor antagonist naloxone (NLX) (6) . Endogenous opioid peptides are continuously available because opioid producing-leukocytes are progressively recruited into inflamed tissue (7 8 9) . The activation of peripheral opioid mechanisms is advantageous, because they lack central side effects like sedation, nausea, respiratory depression, or tolerance (10) . Thus, unraveling the molecular pathways of opioid peptide release and endogenous pain control is a promising strategy for future pain therapy.

To explore mechanisms of endogenous control of inflammatory pain, we have extensively studied the model of complete Freund’s adjuvant (CFA)-induced hindpaw inflammation in rats (reviewed in (2) ). In early inflammation opioid-mediated pain control (antinociception) is generated by PMN, whereas in later stages, it is dependent on monocytes/macrophages and lymphocytes (5 , 9 , 11) . Recruitment of opioid-containing leukocytes is orchestrated by a successive expression of chemokines and adhesion molecules. These steps are essential components of peripheral opioid-mediated antinociception (4 , 5 , 12 , 13) . To elicit antinociception, infiltrating leukocytes have to be stimulated to release opioid peptides. In vivo, cold water swim stress or intraplantar (i.pl.) injection of corticotropin releasing hormone can be applied (14) . In vitro, opioid peptide secretion has been shown in mononuclear cells from lymph nodes (7 , 15) but has not been examined in detail in PMN.

PMN can be stimulated by several groups of chemokines. Depending on species, PMN express the receptors CXCR1, 2 and/or 4, among others. In humans, the best characterized CXCR1/2 ligand is CXCL8 [formerly interleukin (IL)-8]. CXCL8 is a potent trigger of chemotaxis, oxidative burst, and release of granules, including azurophilic (primary), specific (secondary), gelatinase (tertiary), and secretory granules (16) . In rats, CXCR2 is expressed on PMN and its ligands CXCL1 (keratinocyte-derived chemokine), and CXCL2/3 (macrophage inflammatory protein-2) are detectable in early CFA-induced inflammation (5) . In humans and rats, CXCL12 (stromal derived factor-1{alpha}) is the only known ligand for CXCR4. CXCL12 plays a role in the bone marrow retention and homing of PMN (17 , 18) . It can trigger a Ca2+ signal, phosphoinositol-3-kinase (PI3K) activation and oxidative burst in human PMN, but—in contrast to CXCR1/2 ligands—it does not induce granule release (19 20 21) .

Chemokine receptors couple to Gi proteins. After activation and dissociation of the heterotrimeric G-proteins, the resulting Gß{gamma} subunits, in turn, activate phospholipase C (PLC) generating inositol 1,4,5-trisphosphate (IP3). IP3 binds to IP3 receptors (IP3R) on the endoplasmic reticulum leading to release of Ca2+ from intracellular stores (22) . In addition, Gß{gamma} subunits activate PI3K{gamma}, which also has a role in granule release (25) . However, the PI3K pathway is separate from Ca2+ mobilization. For example, blocking PI3K does not influence the rise in intracellular Ca2+ after stimulation with CXCL8 (23 , 24) .

To examine the role of CXCR1/2 ligands in opioid peptide release from PMN and in the inhibition of inflammatory pain in detail, we determined the effects of their injection in early CFA-induced hindpaw inflammation in vivo and identified molecular mechanisms of CXCR1/2 ligand-induced opioid release from PMN by inhibiting intracellular signaling pathways in vitro. Finally, we tested the in vivo functional relevance of these mechanisms for pain control using local adoptive transfer of allogenic PMN that were pretreated with appropriate inhibitors ex vivo.

MATERIALS AND METHODS

Antibodies and reagents
Mouse CXCL12 and rat CXCL2/3 were purchased from Peprotech (London, UK), human CXCL8, human CXCL12, anti-human CXCR1, and CXCR2 Abs from R&D Systems (Minneapolis, MN) and anti-rat CXCR2 antibody (Ab) from Santa Cruz Biotechnology (Santa Cruz, CA). Thapsigargin, ionomycin, NLX, D-Phe-Cys-Tyr-D-Trp-Orn-Thr-Pen-Thr-NH2 (CTOP), and naltrindole hydrochloride (NTI) were obtained from Sigma-Aldrich Chemie (St. Louis, MO) and LY294002, wortmannin, 2-aminoethoxydiphenyl borate (2-APB) and 1,2-bis(o-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid acetoxymethyl ester (AM) (BAPTA/AM) from Calbiochem (San Diego, CA). Thapsigargin, BAPTA/AM, LY294002, wortmannin, and ionomycin were dissolved in DMSO (maximal final concentration 1%). Rabbit anti-rat CXCR4 Ab was produced in the laboratory and characterized before (26) .

Animals and CFA-induced inflammation
Male Wistar rats weighing 180–220 g were injected with 150 µl CFA (Calbiochem) in the right hind paw as described (27) . Experiments were conducted at 2 h after inoculation. All injections were performed under brief isoflurane anesthesia. Animal protocols were approved by the animal care committee of local authorities and were in accordance with the guidelines of the International Association for the Study of Pain (28) .

Measurement of mechanical and thermal hyperalgesia and antinociception
Mechanical nociceptive thresholds were assessed using the paw pressure algesiometer (modified Randall-Selitto test; Ugo Basile) (5) . Rats were handled once per day for 4 days before testing. On the day of testing, rats were held under paper wadding, and incremental pressure was applied via a wedge-shaped, blunt piston onto the dorsal surface of the hind paw by an automated gauge. The pressure required eliciting paw withdrawal, the paw pressure threshold (PPT), was determined by three consecutive trials separated by 10-s intervals. The same procedure was performed on the contralateral paw; the sequence of paws was alternated between subjects to preclude order effects. The treatments were randomized, and the experimenter was blinded to the treatment. A decrease in PPT was interpreted as hyperalgesia (pain), whereas a rise in PPT was interpreted as antinociception (analgesia).

Thermal nociceptive thresholds were measured by the Hargreaves test (29) . Animals were acclimatized to the testing apparatus. Radiant heat was applied to the plantar surface of a hind paw from underneath the glass floor with a high-intensity light bulb, and paw withdrawal latency (PWL) was measured with an electronic timer (IITC Inc/Life Science, Woodland Hills, CA). The PWL was the average of two measurements taken with 20-s intervals. The stimulus intensity was adjusted to give a 9 to 10 s PWL in noninflamed paws, and the cutoff was 20 s to avoid tissue damage. A decrease in PWL was interpreted as hyperalgesia (pain), whereas a rise in PWL was interpreted as antinociception (analgesia).

Chemokine injection
Six rats per group with 2 h CFA were injected i.pl. with 1–100 ng rat CXCL2/3 or 100 ng mouse CXCL12 dissolved in 100 µl of NaCl 0.9% or with solvent only. Mouse CXCL12 has biological activity in rat cells (30) . PPT or PWL was measured 5 min after chemokine injection. In some experiments, the opioid receptor antagonists NLX (nonselective, 0.56 ng i.pl.), CTOP (MOR selective, 2 µg i.pl.) and NTI (MDOR selective, 50 µg i.p. were injected concomitantly. Optimal doses for antagonists were determined in pilot experiments and in previous studies (5 , 31) .

Opioid peptide release
Human PMN were obtained from healthy blood donors using dextran sedimentation, Ficoll separation, and hypotonic lysis (all Amersham Biosciences, Piscataway, NJ). Rat peritoneal PMN were harvested by lavage of the peritoneal cavity with 10 ml PBS/2 mM EDTA 4 h after i.p. injection of 20 ml 1% oyster glycogen in PBS, as described before [Invitrogen (Carlsbad, CA) and Sigma-Aldrich Chemie] (32) .

For measurement of opioid peptide release 5 x 107, PMN were first preincubated with 5 µg/ml cytochalasin B for 5 min in HBSS containing the proteinase inhibitors 5 µg/ml bestatin, 40 µg/ml aprotinin, and 100 µM thiorphan (all Sigma-Aldrich Chemie), as described (7 , 33) . In some experiments, cells were concomitantly incubated with inhibitors, as described in the Results section. Subsequently, cells were stimulated with chemokines, ionomycin, or thapsigargin. Control samples with the solvent DMSO did not induce significant release. Release was terminated after 7 min by rapid cooling, centrifugation, and harvesting of the supernatant. Samples were stored at –20°C until further analysis.

Measurement of opioid peptides by RIA
Opioid peptides were measured in the supernatant using commercially available kits for rat and human END and ENK (Bachem, King of Prussia, PA) (9 , 34) . To prevent peptide degradation, proteinase inhibitors were added after thawing of the samples in addition to the above proteinase inhibitors.

Polymerase chain reaction for chemokine receptor mRNA
RNA from 1 x 106 peritoneal PMN was extracted using Qiagen Mini Kit (Qiagen, Valencia, CA) and transcribed into cDNA using 10 U avian myeloblastosis virus (AMV) Reverse Transcriptase (Roche Diagnostics, Basel, Switzerland). For PCR, the following primers were used: CXCR4 5'GGG CAT CGT CAT CCT GTC CTG and 5'GAG TGT CCA CCC CGT TTC CCT corresponding to bases 633 and 1004 (NIH accession No. U90610); CXCR2 5'CCC ATC TTC ATT CTT CGG AC, and 5'GAC AAT GTT GTA GGG AAG CCA G corresponding to bases 693 and 969 (NIH accession No. U70988); ribosomal protein L-19 (RPL-19), 5'AAT CGC CAA TGC CAA CTC TCG, and 5'TGC TCC ATG AGA ATC CGC TTG corresponding to bases 1521 and 3274 (NIH accession No. X82202). cDNA was amplified with Taq polymerase (Roche Diagnostics) under optimized PCR conditions in a Gene Amp PCR System 9700 Thermocycler (Applied Biosystems, Foster City, CA). Amplification was performed using 40 cycles (RPL-19: 30 cycles), consisting of 45 s at 94°C, 45 s at the gene-specific temperature for primer annealing, followed by 45 s at 72°C. Primer annealing was performed at 68°C for RPL-19, 64°C for CXCR4, and 63°C for CXCR2. As a control for the PCR reaction, RT-negative samples were used.

Chemotaxis assay
Rat CXCL2/3 or mouse CXCL12, dissolved in sterile HBSS/0.5% BSA (Sigma-Aldrich Chemie) were placed in the lower well of a Boyden chamber (Costar) and 4 x 105 rat peritoneal PMN in the upper well. After 1 h incubation at 37°C and 5% CO2, the cell suspension from the lower well was quantified by flow cytometry using fluorescent beads (see below). The migration index was calculated by dividing the number of cells migrating toward CXCL2/3 or CXCL12 by the number of migrating cells using HBSS/BSA alone (34) .

Immunohistochemical staining
Cytospins from 5 x 104 rat peritoneal PMN were prepared using Cytospin 3 (Shandon-Thermo, Waltham, MA). Staining was performed with an avidin-biotin peroxidase complex kit (vectastain, Vector Laboratories, Burlingame, CA), as described previously (35) . Slides were incubated with 0.3% H2O2 and 10% methanol in PBS for 45 min blocking endogenous peroxidase. To reduce nonspecific binding, slides were covered with 0.3% Triton X-100, 1% BSA, 4% goat serum and 4% horse serum. The sections were incubated in four steps with 1) rabbit anti-rat CXCR2 polyclonal Ab at 4°C overnight (1:200 dilution, Santa Cruz Biotechnology) or rabbit anti-rat CXCR4 polyclonal Ab (1 µg/ml) (26) , 2) biotinylated goat anti-rabbit Ab (Vector Laboratories) for 1 h, 3) avidin/biotinylated peroxidase complex for 45 min, and 4) 3,3'-diaminobenzidine (Sigma-Aldrich Chemie) containing 0.01% H2O2 in 0.05 M Tris-buffered saline (pH 7.6, all Vector Laboratories) for 3–5 min. Slides were washed with PBS after each incubation step. Slides were washed with tap water, dehydrated in alcohol, cleared in xylene, and mounted in DPX (Merck Eurolab).

Ca2+ imaging
Fluorescence imaging was performed with a monochromator and a cooled CCD camera (TILL-Photonics) connected to an inverted epifluorescence microscope (Axiovert 100; Carl Zeiss). All imaging experiments were performed in a HEPES-buffered solution containing 128 mM NaCl, 6 mM KCl, 1 mM MgCl2, 1 mM CaCl2, 5.5 mM glucose (Glc), 10 mM HEPES (pH 7.4), and 0.2% (w/v) BSA. For determination of [Ca2+]i, rat peritoneal PMN were placed on dishes coated with poly-L-lysine and then loaded with 4 µM Fura-2/AM (Molecular Probes-Invitrogen) for 30 min at 37°C modified as described previously (36) . After basal recordings, cells were stimulated by subsequent addition of 100 nM CXCL12 and 100 nM CXCL2/3. Fura-2-loaded cells were alternately excited at 340 and 380 nm, and fluorescence was detected through a 505-nm filter. Calibration of [Ca2+]i was performed as described (36) .

Flow cytometry
Cell suspensions were prepared and stained, as described previously (5 , 9 , 37) . Samples were stained with PE-Cy5-conjugated mouse antirat CD45 (4 µg/ml, BD Biosciences, Franklin Lakes, NJ) to label all hematopoetic cells. For intracellular stains, cells were fixed with 1% paraformaldehyde and permeabilized with saponin buffer (0.5% saponin, 0.5% BSA, 0.05% NaN3 in PBS, all Sigma-Aldrich Chemie). Permeabilized cells were incubated with PE-conjugated mouse anti-rat RP-1 (recognizing PMN, 12 µg/ml, BD Biosciences) and FITC-conjugated mouse anti-rat CD68 (formerly named ED1, recognizing monocytes/macrophages, 2 µg/ml, Serotec, Raleigh, NC). Replacement of the primary antibodies with isotype-matched antibodies was used for negative controls.

Absolute numbers of cells were calculated using TruCOUNT tubes with known numbers of fluorescent beads. Data were acquired using a FACSCalibur and analyzed using the CellQuest software (all BD Biosciences).

PMN depletion and adoptive transfer of allogenic PMN
For PMN depletion 6 rats per group were injected i.v. with 80 µl rabbit anti-rat PMN serum or control rabbit serum diluted in 420 µl 0.9% NaCl [Accurate Chemical & Scientific Corporation (Westbury, NY) and Sigma-Aldrich Chemie] 15–18 h before i.pl. injection of CFA, as described previously (5) . After 2 h of inflammation, tissue was collected for flow cytometry, or PPT was measured before and after 100 ng CXCL2/3 i.pl. injection. Reconstitution with PMN was achieved by local injection of 0.1–1 x 106 allogenic PMN, isolated from rats after intraperitoneal (i.p.) glycogen injection (see above) and resuspended in cold PBS. After 15 min, opioid peptide release was stimulated by i.p. CXCL2/3. In some experiments PMN were pretreated with 100 µM BAPTA/AM for 10 min and subsequently washed and resuspended in PBS.

Statistical analysis
Data are presented as raw values (means±SEM). Normally distributed data were analyzed by t test or Mann-Whitney U test. Multiple measurements were analyzed by one-way ANOVA or by one-way ANOVA on ranks in case of not normally distributed data. If necessary, repeated measurement one-way ANOVA was used. Post hoc comparisons were performed by the Student-Newman-Keuls, Dunnett or Dunn method, respectively. Differences were considered significant if P < 0.05. Dose dependency was evaluated by linear regression analysis.

RESULTS

Effects of local CXCR2- and CXCR4-ligands and opioid receptor antagonist on nociceptive thermal and mechanical thresholds
Two hours after i.pl. injection of CFA mechanical nociceptive thresholds (PPT) were reduced in comparison to noninjected paws, demonstrating the presence of local inflammatory pain (Fig. 1 A). The CXCR2 ligand CXCL2/3, but not the CXCR4 ligand CXCL12, injected i.pl. into inflamed hindpaws, elicited significant and dose-dependent antinociception, as indicated by a rise in PPT (Fig. 1A ). Therefore, CXCL12 was used as a control in the following experiments. CXCL2/3-induced PPT elevation was maximal after 5 min, still significant after 10 min, and disappeared after 20 min (Fig 1B ). No change of PPT was seen in the noninflamed paw. To analyze whether local opioid receptors are involved in CXCL2/3-induced antinociception, nonselective (NLX) and selective antagonists at MOR (CTOP) and DOR (NTI) were coadministered. All antagonists significantly reduced CXCL2/3-induced PPT elevation at systemically ineffective doses (5 , 31) (Fig. 1C-E ). NLX-reversible antinociception of CXCL2/3 was also demonstrated by measuring thermal nociceptive thresholds (PWL) (Fig. 1F ).


Figure 1
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  		Figure 1. Local injection of CXCL2/3 but not of CXCL12 blocks inflammatory pain by a peripheral, opioid receptor-dependent mechanism. Rats received i.pl. injections of CXCL2/3 (cross-hatched bars), CXCL12 (gray bar) and normal saline (solvent, open bar) into inflamed paws (2 h post CFA) (A). PPT was determined 5 min after injections. PPT in noninflamed paws is shown for comparison (noninfl) (n=6, *P<0.05 one way ANOVA, Dunnett method vs. solvent control as well as P<0.001 linear regression analysis for dose-dependency of CXCL2/3-induced antinociception). Time course of CXCL2/3-induced antinociception (n=7, *P<0.05 repeated measurement ANOVA, Dunnett method vs. baseline control) (B). C–E) Effect of peripheral opioid receptor blockade by concomitant i.pl. naloxone (0.56 ng, NLX), naltrindole hydrochloride (50 µg, NTI) or D-Phe-Cys-Tyr-D-Trp-Orn-Thr-Pen-Thr-NH2 (2 µg, CTOP) on CXCL2/3 (100 ng)-induced antinociception (control: saline, open bar; n=6, *P<0.05 one-way ANOVA, Student-Newman-Keuls method). F) PWL (2 h post-CFA) before (baseline) and after i.pl. CXCL2/3 (100 ng) and the effect of concomitant injection of NLX (0.56 ng, solvent control: saline; n=6, *P<0.05 t test compared to baseline). Data are expressed as means ± SEM.

Effect of CXCR1/2- and CXCR4-ligands on opioid peptide release from human PMN
On the basis of these opioid-dependent antinociceptive effects of a CXCR2 ligand and the lack of effect of a CXCR4 ligand, we hypothesized that CXCR2 ligands can release opioid peptides from PMN. Incubation of human PMN with the CXCR1/2 ligand CXCL8 stimulated a significant dose-dependent release of END and ENK (Fig. 2 A), which was significantly reduced by preincubation with blocking antibodies against CXCR1 and CXCR2 (Fig. 2B ). As a control, no opioid peptide release was observed after CXCL12 incubation.


Figure 2
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  		Figure 2. CXCL8 but not CXCL12 elicits opioid peptide release from human PMN via CXCR1/2 stimulation. A) Human PMN were stimulated with increasing doses of CXCL8 and CXCL12 for 7 min. ENK (left) and END (right) release was measured in the supernatant by RIA (n=9–15). B) Effect of PMN preincubation with anti-CXCR1 Ab (aCXCR1, 10 µg/ml) and anti-CXCR2 Ab (aCXCR2, 100 µg/ml) for 25 min (hatched bars) on CXCL8-induced opioid release (50 nM, cross-hatched bar, n=10–14; *P<0.05 vs. respective controls, all one-way repeated-measures ANOVA, Dunnett and Student-Newman-Keuls method, respectively). Data are expressed as means ± SEM.

Dependence of opioid peptide release on intra- vs. extracellular Ca2+ and PI3K activation in human PMN
To analyze the signaling pathways involved in opioid peptide release, human PMN were incubated with the Ca2+ ionophore ionomycin and with thapsigargin. Thapsigargin blocks the sarcoendoplasmic reticulum Ca2+ ATPases, resulting in passive Ca2+ leak from intracellular stores (38) . Both treatments, as well as CXCL8 significantly stimulated opioid peptide release (Fig. 3 A, B). The removal of extracellular Ca2+ did not affect CXCL8-induced opioid peptide release (Fig. 3B ). Preincubation with either thapsigargin (for 10 min in the absence of extracellular Ca2+ to deplete intracellular Ca2+ stores; Fig. 3D ), or with BAPTA/AM (a membrane permeable intracellular Ca2+ chelator; Fig. 3C ) abrogated the CXCL8-induced release of opioid peptides. Blockage of IP3R by 2-APB also resulted in a significant reduction of CXCL8-induced release (Fig. 3E ). Taken together, although direct entry of extracellular Ca2+ elicited opioid release, CXCL8-induced release was independent of extracellular Ca2+ but required IP3R-triggered Ca2+ release from intracellular stores. To test the possible involvement of PI3K, we applied the two PI3K inhibitors wortmannin and LY294002. Both inhibitors significantly reduced CXCL8-stimulated END and ENK release but did not completely block the secretion (Fig. 3F ).


Figure 3
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  		Figure 3. Secretion of opioid peptides from human PMN depends on Ca2+ release from intracellular stores and on PI3K activation. A) Release of ENK and END after 7 min incubation with ionomycin (iono, 10 µM) or thapsigargin (thap, 1 µM) (n=7–11; open columns = solvent only in all subsequent experiments). B) CXCL8 (50 nM, 7 min stimulation, cross-hatched bars in all subsequent experiments) induces opioid peptide release in the presence (open bars) and absence (gray bars) of extracellular Ca2+ (-[Ca2+]e; n=6–8). C) CXCL8-induced opioid release was blocked by the intracellular Ca2+ chelator BAPTA/acetoxymethyl ester (100 µM, hatched bars) after 10 min preincubation with thapsigargin in Ca2+-free medium (thap, 1 µM, gray bars; thapsigargin alone, gray cross-hatched bars: thapsigargin+CXCL8, n=8–12) (D), the IP3R blocker 2-APB (100 µM, hatched bars, n=7–14) (E) and partially by the PI3K-inhibitors LY294002 (LY, 100 µM) and wortmannin (wort, 100 nM, n=10–15) (F). *P < 0.05 statistically significant difference compared to respective controls (all one-way repeated-measures ANOVA, Student-Newman-Keuls method). Data are expressed as means ± SEM.

CXCR2 and CXCR4 expression and effects of their ligands on rat PMN
Glycogen-elicited rat peritoneal PMN were >95% viable according to trypan blue exclusion and >95% pure as demonstrated by FACS. They expressed CXCR2 and CXCR4 mRNA and protein as confirmed by PCR and immunohistochemistry (Fig. 4 A, B). In the Boyden chamber PMN migrated in response to both CXCL2/3 and CXCL12, but a 100-fold lower concentration of CXCL2/3 compared to CXCL12 was needed for maximal chemotaxis (Fig. 4C ). While CXCL2/3 induced a robust elevation of [Ca2+]i only few cells responded to CXCL12 (Fig. 4D ). Similar to human PMN, significant dose-dependent opioid peptide release was observed in rat peritoneal PMN after stimulation with the CXCR2 ligand CXCL2/3 but not with the CXCR4 ligand CXCL12 (Fig. 5 A). CXCL2/3-induced END and ENK secretion from rat PMN was not altered in the absence of extracellular Ca2+ (Fig. 5B ), while chelating intracellular Ca2+ by BAPTA/AM abolished opioid peptide secretion (Fig. 5C ). Blocking of PI3K by LY294002 and wortmannin significantly, but only partially, reduced opioid peptide release (Fig. 5D-E ). In summary, similar to human PMN, END and ENK secretion in rat PMN was dependent on intracellular Ca2+ and was modulated by PI3K activation.


Figure 4
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  		Figure 4. Expression and functions of CXCR2 and CXCR4 in rat PMN. A) CXCR2, CXCR4 and RPL-19 mRNA, CXCR2 and CXCR4 immunoreactivity (B) (brown cells) in glycogen-elicited rat peritoneal PMN (1 and 2: PMN from different rats, + cDNA synthesis with reverse transcriptase, – cDNA synthesis without reverse transcriptase as negative control). C) In vitro migration of rat peritoneal PMN in response to different doses of CXCL2/3 and CXCL12 was measured in the Boyden chamber. The migration index was calculated relative to baseline migration (n=6, *P<0.05, one-way repeated-measures ANOVA, Dunnett method). D) PMN were loaded with Fura-2, and changes in [Ca2+]i were analyzed after the addition of CXCL12 (60 s) and CXCL2/3 (240 s, both 100 nM).


Figure 5
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  		Figure 5. Intracellular Ca2+- and PI3K-dependency of CXCL2/3-induced opioid peptide release from rat PMN. A) Rat PMN were stimulated with the indicated concentrations of CXCL2/3 and CXCL12 for 7 min. ENK (left) and END (right) release was measured in the supernatant by RIA (n=6–19). B) CXCL2/3 (1000 nM, cross-hatched bars in all experiments)-induced opioid peptide release was analyzed in the presence (open bars) and absence (gray bars) of extracellular Ca2+ (-[Ca2+]e, n=9). CXCL2/3-induced opioid peptide release was measured after preincubation of rat PMN (for 10 min) with the intracellular Ca2+ chelator BAPTA/acetoxymethyl ester (100 µM) (C) or the PI3K-inhibitors LY294002 (D) and wortmannin (E) (100 µM and 100 nM, respectively) (n=5–9). *P < 0.05 significant difference compared to respective controls (all one-way repeated-measures ANOVA, Student-Newman-Keuls method). Data are expressed as means ± SEM.

Effect of PMN depletion, allogenic PMN adoptive transfer and intracellular Ca2+ depletion on CXCL2/3-induced antinociception
PMN depletion by an anti-PMN serum resulted in over 90% reduction of PMN without affecting the number of monocytes/macrophages in the paw as quantified by flow cytometry (Fig. 6 A). Identical degrees of hyperalgesia (i.e., inflammatory pain) were detected in CFA-treated rats, regardless of PMN depletion (baseline, Fig. 6B ). Under these conditions, CXCL2/3-mediated antinociception was significantly attenuated as detected by a reduced rise in PPT in PMN-depleted rats compared to animals treated with control IgG (Fig. 6B ).


Figure 6
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  		Figure 6. Effects of PMN depletion and reconstitution with adoptively transferred PMN on CXCL2/3-induced pain control. Rats were pretreated with i.v. anti-PMN serum (gray bars); control animals received nonimmune rabbit serum (open bars). Two hours after CFA the number of leukocytes in the paw (A) were quantified by flow cytometry. Likewise, PTT (B) was measured before (baseline) and after i.pl. injection of 100 ng CXCL2/3 (n=6, *P<0.05 t test). C) Different numbers of glycogen-elicited peritoneal PMN from allogenic animals were injected into the inflamed paws of PMN-depleted rats. PPT was obtained 15 min later and again after 100 ng i.pl. CXCL2/3 (n=6, *P<0.05, one-way ANOVA, Dunnett method, and P<0.001 in linear regression analysis for dose dependency of transferred PMN, open bar: before CXCL2/3 without PMN depletion; cross-hatched bar: effect of i.pl. CXCL2/3 without PMN depletion, gray bars: PMN {conpict2}depletion, striped bars: PMN reconstitution). D) Effect of ex vivo BAPTA/acetoxymethyl ester (100 µM) or solvent pretreatment before allogenic PMN transfer. Rats were PMN-depleted and reconstituted as described in (C) using 1 x 106 PMN and CXCL2/3-induced PPT elevation was measured thereafter (n=5 or 6, *P<0.05, one-way ANOVA, Student-Newman-Keuls method). Data are expressed as means ± SEM.

Subsequent adoptive transfer of allogenic PMN by i.pl. injection dose-dependently reestablished CXCL2/3-induced antinociception in the inflamed paw (Fig. 6C ). Baseline PTT at 2 h CFA before injection of CXCL2/3 was not affected by the different treatments (control: 35.8±1.4g, anti-PMN: 36.2±1.2g, anti-PMN+ transfer 1x105 PMN: 37.2±1.5g, anti-PMN+transfer 3x105 PMN: 38.0±1.3g; anti-PMN+transfer 1 x 106 PMN: 38.3±1.0g, n=6, mean±SEM). To analyze dependency of CXCL2/3-induced antinociception on intracellular Ca2+ in vivo, allogenic PMN were pretreated ex vivo with BAPTA/AM. This treatment significantly reduced CXCL2/3-induced antinociception following PMN reconstitution (Fig. 6D ). No change in baseline PTT at 2 h CFA before injection of CXCL2/3 was seen between the different treatment groups (control: 36.6±0.9g, anti-PMN: 38.3±1.1g, anti-PMN+transfer PMN treated with solvent: 38.6±0.9g, anti-PMN+transfer PMN treated with BAPTA: 38.8±1.2g, n=6, mean±SEM). In summary, antinociception elicited by CXCL2/3-induced opioid release from PMN was dependent on intracellular Ca2+ in vivo.

DISCUSSION

In this study, we demonstrated that local injection of CXCR1/2 but not CXCR4 ligands decrease inflammatory pain through release of opioid peptides from PMN. CXCR1/2 ligand-triggered opioid peptide secretion from human PMN in vitro depended on intra- but not extracellular Ca2+ availability. The rise of intracellular Ca2+ necessary for opioid peptide secretion was accomplished by IP3R stimulation and subsequent release of Ca2+ from intracellular stores. Activation of PI3K partially supported secretion. Release of opioid peptides in response to CXCR2 stimulation of rat PMN required the same intracellular signals. The in vivo relevance of these findings was confirmed since CXCL2/3-induced antinociception 1) was abolished by selective PMN depletion, 2) was reconstituted by local adoptive transfer of allogenic PMN in depleted rats, and 3) was blocked following intracellular Ca2+ chelation before adoptive PMN transfer. This approach allowed studying intracellular signaling requirements in leukocytes in vivo without affecting other resident cell populations such as peripheral sensory neurons.

Other groups have shown that chemokines like CXCL12 or CXCL1 can directly activate nociceptive neurons and cause pain (hyperalgesia) (39 , 40) . In contrast, local CXCL2/3 injection resulted in specific PMN recruitment but did not induce pain (29) . These studies were performed in the absence of inflammation. In models of inflammatory pain, blockage of CXCR2 ligands yielded contradictory results (5 , 41) and additional injection of CXCL2/3 into inflamed paws did not aggravate pain despite increased PMN recruitment (29 , 37) . In our present study CXCL2/3, but not CXCL12, injected into inflamed paw tissue resulted in local opioid-dependent mechanical and thermal analgesia (Fig. 1A ). In line with our previous studies (31) , analgesia was mediated by the two opioid receptors MOR and DOR (Fig. 1C-E ). Although CXCL2/3 completely reversed mechanical hyperalgesia, it was less efficient in thermal hyperalgesia (Fig. 1F ). These results are supported by C fiber recordings showing that opioids suppress mechanical stimuli more effectively than thermal stimuli (42) . The CXCR2-mediated antinociceptive effects are an extension of our previous studies that demonstrated opioid-mediated pain control by mediators such as corticotropin releasing factor (14) . In contrast to corticotropin-releasing factor, CXCR2 ligands mainly target PMN (43 , 44) , as confirmed by PMN depletion in our experiments (Fig. 6A, B ). Furthermore, we previously showed that CXCR2 ligands orchestrate the recruitment of opioid-containing PMN in early inflammation (5) . Taken together, CXCR2 ligands appear to play a key role in the control of inflammatory pain by the release of opioid peptides from PMN.

In agreement with the effects in vivo, we showed that CXCR1/2 ligands induce opioid peptide release from PMN in vitro, while the CXCR4 ligand had no effect (Fig. 2 and 5) . CXCL8-induced release was partially but not completely blocked by preincubation with antibodies to CXCR1 and CXCR2 (Fig. 2) , in line with previous findings (45) . This might be due to incomplete blockade of the ligand binding groove, in contrast to small molecule antagonists. Although in vitro stimulation with both CXCR1/2 and CXCR4 ligands (CXCL8 and CXCL12) has been shown to increase intracellular Ca2+ in human PMN (20 , 46) , no granule release in response to the CXCR4 ligand has been demonstrated so far. Consistently, CXCL12 did not produce release in our in vitro and in vivo experiments, and only a few cells responded to CXCL12 with an increase in intracellular Ca2+ concentration (Fig. 4) . Both chemokines, however, were chemotactic in vitro, albeit with different concentration requirements (Fig. 4) , indicating the presence of functional CXCR2 and 4 receptors on rat PMN. The release of granule subpopulations is dependent on the intracellular Ca2+ concentration (47) . Tertiary granules are released by small Ca2+ elevations while primary granules require larger Ca2+ concentrations (16) . Our own unpublished data show that opioid peptides are contained in primary granules, suggesting that CXCL12-induced intracellular Ca2+ elevation might be insufficient for opioid peptide release.

We further examined signal pathways for the secretion of opioid peptides and showed dependency on intracellular Ca2+ release and PI3K activation (Figs. 3 and 5) . No significant difference was seen between the two peptides END and ENK or between signaling pathways in human and rat PMN. However, the absolute opioid release from human peripheral blood PMN was higher than from peritoneal rat PMN. This may be explained by the fact that rat PMN were recruited by i.p. glycogen injection. This recruitment is CXCR2 ligand-dependent and results in CXCR2 down-regulation (48 , 49) .

We further showed that incubation of PMN with thapsigargin induces opioid release, most likely by leakage of Ca2+ from intracellular stores (25) (Fig 3A ). However no thapsigargin-induced release was found in the absence of extracellular Ca2+ (Fig. 3D ). These findings are similar to thapsigargin-induced glucuronidase secretion from primary granules in PMN, which was also abolished in the absence of extracellular Ca2+ and by a blocker of receptor-mediated Ca2+ entry (50) . Therefore, it seems that thapsigargin-induced secretion of opioid peptides in our setting is dependent on both intracellular and extracellular Ca2+ sources.

This mechanism is different from CXCR1/2-induced opioid peptide secretion. Activation of CXCR1/2 is coupled to Gi proteins leading to stimulation of PLC, intracellular IP3 production and Ca2+ mobilization (51) . Our findings are consistent with these studies. However other targets, including some membrane-bound transient receptor potential channels, can also be blocked by 2-APB (52) . Since in our experiments, however, CXCL2/3-induced opioid peptide release was independent of extracellular Ca2+, the action of 2-APB was most likely due to its IP3R blocking properties. Although stimulation of PMN with platelet-activating factor, formyl-Met-Leu-Phe or leukotriene B4 induces store-operated Ca2+ entry (prolonged elevation of [Ca2+]i due to entry of extracellular Ca2+), no store-operated Ca2+ entry has been found after CXCR1 stimulation with CXCL8 despite significant release of Ca2+ from intracellular stores (24 , 53) . These results support our findings of independency of CXCL8-induced opioid peptide release from extracellular Ca2+. Other stimuli like formyl-Met-Leu-Phe or leukotriene B4, however, require extracellular Ca2+ for the release of different types of granules with the exception of secretory granules (54) . These differences in extracellular Ca2+ requirement might be related to differences in the signaling pathways involved. In contrast to our present results in PMN, we previously found that opioid peptide release from lymph node cells was dependent on extracellular Ca2+. However, apart from the different cell type, this study used corticotropin-releasing factor and IL-1ß as releasing agents (7 , 15) . In summary, our present data suggest that chemokine-induced opioid secretion from PMN would exclusively be dependent on intracellular Ca2+ release from the endoplasmic reticulum without the need of extracellular Ca2+.

In addition, our data indicate that PI3K activation might contribute to opioid peptide release (Fig. 3) . Our results are supported by other studies showing that degranulation by CXCL8 or other mediators is also dependent on PI3K activation (55 56 57) . In PMN the PI3K pathway is separate from Ca2+ mobilization because wortmannin does not influence the rise in intracellular Ca2+ after stimulation with CXCL8 (23 , 24) , and Ca2+ mobilization after leukotriene B4 stimulation is not reduced in PI3K{gamma} KO mice (24) . However, the PI3K class and subclass involved in CXCL8-induced release still remains to be elucidated because wortmannin and LY294002 inhibit all classes of PI3K. Previous studies showed that leukotriene-induced secretion of primary granules is inhibited in PI3K{gamma} KO mice (24 , 58) and that selective PI3K{delta} inhibitors blocked formyl-MetLeuPhe- and TNF-{alpha}-induced release of primary granules (59) . Further studies are needed to identify the PI3K subtype responsible for opioid peptide secretion in PMN and to elucidate the pathways downstream of PI3K activation.

To confirm the relevance of intracellular Ca2+ for opioid peptide release in vivo, we employed an approach to avoid impairment of sensory nerve functioning by signaling cascade inhibitors (60) . To this end, we established a model of ex vivo treatment and subsequent adoptive cell transfer. Allogenic PMN have the advantage that they do not elicit a graft vs. host reaction and can be used in neutropenic patients (61) . Thus we depleted rats of PMN and performed local adoptive transfer of allogenic PMN. This demonstrated dose-dependent reconstitution of CXCL2/3-induced analgesia (Fig. 6C ). Chelating intracellular Ca2+ of PMN before transfer significantly impaired CXCL2/3-induced analgesia. Because we did not observe differences in baseline hyperalgesia, our approach apparently did not compromise sensory nerve function. This model provides a novel tool to selectively study the signaling pathway requirements of release from PMN in vivo.

What could be the clinical implications of this study? Although the analgesic effect of a single dose of exogenous CXCL2/3 only lasts 10 min, endogenous CXCR1/2 ligands are continuously produced during inflammation. Therefore, it is conceivable that endogenous CXCR1/2 ligands might constantly stimulate infiltrating PMN to release opioid peptides (5 , 34) . In consequence, this might constitute an endogenous system to counteract pain. This notion is in line with data obtained in postoperative patients that have shown a role of immune cell-derived endogenous opioid peptides in pain control (6) . Interference with this system using CXCR2 antagonists (62) might have the unwanted side effect of pain aggravation. In addition, we have previously shown that coinjection of CFA and CXCL2/3 does not enhance hyperalgesia (37) and that local CXCL2/3 injection into noninflamed tissue elicits PMN recruitment without hyperalgesia (29) . Thus, additional chemokine application should not worsen pain and PMN-specific chemokines might be useful for treatment of pain states when PMN are predominant, e.g., in acute inflammation or postoperatively.

ACKNOWLEDGMENTS

This study was supported by the Deutsche Forschungsgemeinschaft (KFO 100/1) and by the European Society of Anaesthesiology (ESA). The authors wish to thank Susanne Kotré and Katharina Hopp for expert technical assistance.

Received for publication March 14, 2006. Accepted for publication July 24, 2006.

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