Published online before print June 10, 2005
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,1
* Department of Rheumatology and Inflammation Research, Göteborg University, Göteborg, Sweden;
DRDC/BBSI (UMR 5092, CEA/CNRS/UJF), Grenoble, France; and
Department of Clinical Virology, Göteborg University, Göteborg, Sweden
1Correspondence: Department of Clinical Virology, Göteborg University, Guldhedsgatan 10, S-413 46 Göteborg, Sweden. E-mail: charlotta.movitz{at}microbio.gu.se
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Key Words: granulocytes • inflammation • resolution • cell activation • signal transduction
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Recent findings suggest that annexin AI-derived peptides possess not only anti-inflammatory activities but also can activate neutrophils directly. The triggering of neutrophil function as well as the anti-inflammatory signaling have been suggested to rely on the interactions with one or possibly both neutrophil members of the formyl peptide receptor (FPR) family [8 , 12 ]. The two FPR family members expressed in neutrophils, FPR and FPR-like 1 (FPRL1), belong to the G protein-coupled seven transmembrane receptor family. FPRL1 was originally cloned from human phagocytes by low-stringency hybridization of a cDNA library with the FPR sequence and initially defined as an orphan receptor [13 ]. The first ligand shown to bind this receptor with high affinity was the lipid mediator lipoxin A4 (LXA4) [14 ]. During the past few years, several other ligands for FPRL1 have been identified [15 16 17 18 ]. It should be noted that despite the fact that different protein/peptide ligands bind to the same receptor, they do not contain any sequence homologies. Also, whereas the protein/peptide ligands have in common that they are potent activators of neutrophils [15 , 16 , 18 ], LXA4 is an inhibitor of neutrophil functions such as adhesion and chemotaxis [19 , 20 ]. The fact that the cytoplasmic regions of FPRL1 share approximately 80% identity with FPR suggests that signaling from the two receptors should be similar, but so far, all agonists described to interact with FPR trigger neutrophil activation. Recently published experiments using a desensitization protocol and receptor expressing human embryonic kidney (HEK)293 cells reveal that N-terminal annexin AI peptides activate cells through FPR and FPRL1 (and also through FPRL2, which is expressed in monocytes) [12 ]. Moreover, the inhibitory effect of such peptides has been shown to be mediated, at least in part, by FPR [8 , 21 ], although a recent study in mice suggests that the deactivating signal is independent of FPR but might involve FPRL1 [22 ]. Taken together, these data suggest that the receptor system in neutrophils responsible for the activating and deactivating actions of the annexin AI-derived peptides needs to be characterized in more detail.
In this study, we show that a peptide, derived from the N-terminal region of annexin AI, activates neutrophils to produce and release superoxide anions. The signals activating the NADPH-oxidase were generated by FPR. We also show that the inhibitory signal generated in neutrophils by the same peptide was transduced by a not-yet identified receptor distinct from FPR and FPRL1 and affects the NADPH-oxidase activity induced by several different chemoattractants.
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Isolation of human neutrophils
Neutrophil granulocytes were isolated from buffy coats obtained from healthy adults. After dextran sedimentation at 1 g, hypotonic lysis of the remaining erythrocytes, and centrifugation in a Ficoll-Paque gradient [25
], the neutrophils were washed twice and resuspended (1x107 cells/ml) in KRG phosphate buffer, containing glucose (10 mM), Ca2+ (1 mM), and Mg2+ (1.5 mM; pH 7.3). The cells were stored on melting ice until used.
Stable expression of FPRs in undifferentiated HL-60 cells
The stable expression of FPR, FPRL1, and FPRL2 in undifferentiated HL-60 cells has been described previously [16
, 18
]. The control experiments with the specific agonists (fMLF for FPR and WKYMVM/m for FPRL1 and FPRL2) were performed at each experimental event and turned out positive. Cells were cultured in RPMI-1640 medium/glutaMax I, and the maximal density was maintained below 2 x 106 cells/ml.
Differentiation of non-transfected HL-60 cells
HL-60 cells were passed once a week in RPMI medium supplemented with 10% (v/v) heat-inactivated fetal calf serum, 100 U/ml penicillin, 100 mg/ml streptomycin, and 2 mM L-glutamine. The cells were induced to differentiate by the addition of DMSO to a final concentration of 1% (v/v). The cells were harvested after 5 days of cultivation, washed, and resuspended in KRG. If the cells were cultivated in the presence of fMLF and WKYMVm, 10–7 M fMLF and 10–7 M WKYMVm were added 1, 2, and 5 days after addition of DMSO.
Neutrophil NADPH-oxidase activity
Neutrophil production and release of superoxide anions were measured by an isoluminol-enhanced chemiluminescence (ECL) assay [26
]. The ECL activity was measured in a six-channel Biolumat LB 9505 apparatus (Berthold Co., Wildbad, Germany), using disposable, 4 ml polypropylene tubes with a 400-µl reaction mixture containing 4 x 105 neutrophils, HRP (4 U), and isoluminol (20 µM). boc-MFL (40 µl, 10–5 M), cyclosporin H (10–6 M), PBP10 (1 µM), or WRWWWW (5x10–6 M) was added as indicated in Results. The measuring tubes were equilibrated for 5 min at 37°C, and the cells were activated by addition of one or two peptide/protein agonists at different time-points. The light emission was recorded continuously.
Determination of changes in cytosolic calcium
Cells were loaded with 2 µM Fura-2/AM (Molecular Probes, Eugene, OR), according to the instructions given by the manufacturer. Calcium measurements were carried out as described earlier [16
] using an LC50 fluorescence spectrophotometer (LS50B, Perkin Elmer, Wellesley, MA). Intracellular calcium concentrations ([Ca2+]i) were calculated using the formula: [Ca2+]i = Kd(F–Fmin)/(Fmax–;F) with a dissociation constant (Kd) for Fura-2 of 224 nM, where Fmax is the fluorescence in the presence of 0.04% Triton X-100, and Fmin is the fluorescence after the addition of 5 mM EGTA and 30 mM Tris-HCl, pH 8.7.
Statistical analysis
Two-tailed, paired Student’s t-test was performed to determine statistical significance. A Pvalue of <0.05 was regarded as significant.
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Figure 1. Ac9–25 induced release of superoxide anions. Neutrophils were activated by addition of increasing concentrations of Ac9–25 (10–100 µM final concentration). The release of superoxide anions was measured by isoluminol-ECL. Abscissa, Time of study; ordinate, superoxide production, given as 106 counts per minute (Mcpm). The curves are from a representative experiment.
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Figure 2. Inhibition of Ac9–25 induced release of superoxide anions by receptor antagonists. (A) Neutrophils were incubated with (solid line) or without (broken line) 10–5 M boc-MLF (upper panel) or 10–6 M cyclosporin H (lower panel) for 5 min at 37°C and activated by the addition of Ac9–25 (50 µM final concentration). The release of superoxide anions was measured by isoluminol-ECL. Abscissa, Time of study; ordinate, superoxide production given as Mcpm. The curves are from a representative experiment. The effects of boc-MLF (upper inset) and cyclosporin H (lower inset) on fMLF- and WKYMVM (10–7 M final concentration)-induced radical production are expressed as percent of control cells (cells incubated in the absence of antagonist) and shown as mean ± SEM, n = 3. (B) Neutrophils were incubated with 5 x 10–6 M (broken line) or without WRWWWW (solid line) for 5 min at 37°C and activated by addition of Ac9–25 (50 µM final concentration). The release of superoxide anions was measured by isoluminol-ECL. Abscissa, Time of study; ordinate, superoxide production given as Mcpm. The curves are from representative experiments. The effects of the FPRL1 antagonist WRWWWW on WKYMVM and fMLF (10–7 M final concentration, inset)-induced superoxide anion release are included as control (data are presented as percent of control cells and shown as mean±SEM, n=3).
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Figure 3. Desensitization of Ac9–25 induced NADPH-oxiadse activity by fMLF. Neutrophils triggered with fMLF (10–7 M, solid line, upper panel) or WKYMVM (10–7 M, solid line, lower panel) or simply preincubated without agonist (broken lines) were activated through addition of Ac9–25 (50 µM final concentration). The release of superoxide anions was measured by isoluminol-ECL. Abscissa, Time of study; ordinate, superoxide production given as Mcpm. The curves are from representative experiments.
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Figure 4. Ac9–25 induced calcium mobilization in FPR-, FPRL1-, and FPRL2-transfected HL-60 cells. Changes in intracellular calcium were measured in undifferentiated HL-60 cells stably transfected with FPR (upper panel), FPRL1 (lower-left inset), or FPRL2 (lower-right inset). Cells loaded with Fura-2 were activated through an addition of Ac9–25 (100 µM final concentration), and the change in fluorescence was measured. Abscissa, Time of study; ordinate, intracellular Ca2+ concentration in nM. The curves are from representative experiments.
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80% by raising the Ac9–25 concentration to 100 µM (Fig. 5B)
. The lower concentration of Ac9–25 (i.e., 20 µM) only had an inhibitory effect if the fMLF concentration was decreased to 10–8 M.
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Figure 5. Ac9–25 inhibits the fMLF-induced release of superoxide anions. (A) Neutrophils were incubated with (broken line) or without (solid line) Ac9–25 (50 µM final concentration) and reactivated through addition of fMLF (10–7 M final concentration). The release of superoxide anions was measured by isoluminol-ECL. The time is given on the abscissa, and the amount of superoxide on the ordinate is given as Mcpm. The curves are from a representative experiment. (B) Neutrophils were incubated with different concentrations of Ac9–25 (0–100 µM final concentration) for 5 min at 37°C and then activated through addition of fMLF (10–7 or 10–8 M final concentration). The release of superoxide anions was determined from the peak values of the responses and expressed as percent of control (i.e., the response from cells incubated in the absence of Ac9–25) and given as mean ± SEM, n = 3. **, P < 0.01, versus cells incubated in the absence of Ac9–25. (C) Neutrophils were incubated with (broken line) or without (solid line) a suboptimal concentration of fMLF (10–8 M final concentration) and activated through addition of a new dose of fMLF (10–7 M final concentration). The release of superoxide anions was measured by isoluminol-ECL. The time is given on the abscissa, and the amount of superoxide on the ordinate is given as Mcpm. The curves are from a representative experiment. The release of superoxide anions is also presented as the response expressed as percent of control cells (i.e., the response from cells incubated in the absence of 10–8 M fMLF before stimulation with 10–7 M fMLF) and is given as mean ± SEM, n = 4. *, P< 0.05.
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Ac9–25 inhibits superoxide anion production in neutrophils through a receptor distinct from FPR and FPRL1
The fact that Ac9–25 is unable to prime the fMLF-triggered activity suggests that the peptide, in addition to its activating property, also evokes an inhibitory signal that may affect the oxidase activity induced by other stimuli. We used several different approaches to determine if Ac9–25 inhibits neutrophil respiratory burst through the receptor responsible for activation, i.e., FPR. The receptor-specific antagonist boc-MLF was used to block signaling through FPR. Neutrophils were incubated with boc-MLF, treated with Ac9–25, and challenged with the FPRL1-specific agonist WKYMVM. In accordance with earlier presented data, the Ac9–25-induced respiratory burst is antagonized by the FPR antagonist (Figs. 2A
and 6A
) but has no effect on WKYMVM (Fig. 2A
, upper panel inset). The inhibitory effect that Ac9–25 possesses on the WKYMVM-triggered activity also remains in the presence of boc-MLF (Fig. 6A)
. To investigate this phenomenon in more detail, the effects of Ac9–25 on the response induced by other agonists operating through G protein-coupled receptors were investigated. The peptide WKYMVm, C5a, IL-8, and PAF evoke a respiratory burst. The NADPH-oxidase activity induced by these agonists was inhibited by Ac9–25 in the presence of boc-MLF (Fig. 6B)
. When using a stimulus such as PMA, which bypasses cell-surface receptors and directly activates protein kinase C (PKC), Ac9–25 was without effect on the respiratory burst (Fig. 6C)
. This shows that the inhibitory effect induced by Ac9–25 is not related to an effect directly on the oxidase and second, that there is no effect on signaling downstream of PKC. These results suggest that the inhibitory signal induced by Ac9–25 is generated through a receptor distinct from FPR. This suggestion gained further support by the results obtained with differentiated HL-60 cells. When these cells were differentiated in the presence of fMLF and WKYMVm (an agonist to FPRL1 and FPR), they were unable to elicit any burst activity when stimulated with the FPR-specific ligand fMLF, the FPRL1-specific ligand WKYMVM, or Ac9–25 (Fig. 7
, lower panels a–c, respectively). To study the importance of FPR and FPRL1 during Ac9–25-induced inhibitory signaling, these cells were stimulated with Ac9–25 before stimulation with C5a, which binds to the G protein-coupled C5aR. The burst induced by C5a was, however, reduced when the cells were preincubated with Ac9–25 (Fig. 7
, upper panel). This suggests that the inhibitory signal is also generated in cells lacking functional FPRs and FPRL1s. By using HL-60 cells, differentiated only in the presence of fMLF, we could also confirm our data presented in this study, showing that Ac9–25 activates neutrophils through FPR (data not shown).
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Figure 6. Ac9–25 induced inhibition of neutrophil NADPH-oxidase activity in the presence of an FPR antagonist. (A) Neutrophils incubated with (broken line) or without (dotted line) boc-MLF (10–5 M final concentration) were activated through the addition of Ac9–25 (50 µM final concentration). The cells were then challenged with the FPRL1 agonist WKYMVM (10–7 M final concentration). The control (solid line) was preincubated in the absence of boc-MLF and Ac9–25 and challenged only with WKYMVM. The release of superoxide anions was measured by isoluminol-ECL. The time is given on the abscissa, and the amount of superoxide is on the ordinate given as Mcpm. The curves are from representative experiments. (B) Neutrophils were incubated with boc-MLF (10–5 M final concentration) for 5 min at 37°C and stimulated with Ac9–25 (50 µM final concentration). The cells were reactivated through addition of one of five different chemoattractants [10–7 M WKYMVM, 10–8 M WKYMVm, 0.5 µg/ml IL-8, 5x10–7 M platelet-activating factor (PAF), or 0.1 µg/ml C5a, given as final concentrations]. The release of superoxide anions was determined from the peak values of the responses and expressed as percent of control (i.e., the response from cells incubated in the absence of Ac9–25) and is given as mean ± SD (n=3–4). *, P < 0.05; **, P < 0.01, versus cells incubated in the absence of Ac9–25. (C) Neutrophils were activated by addition of Ac9–25 (50 µM final concentration, broken line) and then reactivated by PMA (5x10–8 M final concentration). The nonpretreated control cells (solid line) were activated with PMA in parallel with the Ac9–25-treated cells. The release of superoxide anions was measured by isoluminol-ECL. The time is given on the abscissa, and the amount of superoxide on the ordinate is given as Mcpm. The curves are from a representative experiment.
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Figure 7. Ac9–25 inhibits C5a-induced superoxide anion release in FPR/FPRL1-desensitized HL-60 cells, which were differentiated in the presence of fMLF (10–7 M final concentration) and WKYMVm (10–7 M final concentration) for 5 days. These cells were unresponsive with respect to superoxide release when triggered by the agonists fMLF (lower panel a), WKYMVM (lower panel b), or Ac9–25 (lower panel c). Cells, differentiated in the presence of fMLF/WKYMVm, released superoxide anions when triggered by C5a (upper panel, solid line), whereas cells preincubated with Ac9–25 (50 µM final concentration) before C5a stimulation responded with a reduced superoxide anion release (dotted line). The release of superoxide anions was measured by isoluminol-ECL. The time is given on the abscissa, and the amount of superoxide is on the ordinate given as Mcpm. The curves are from representative experiments.
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Figure 8. The FPRL1 antagonist WRWWWW is without effect on Ac9–25-mediated inhibition of superoxide release. Neutrophils incubated with (solid line) or without (dotted line) the FPRL1 antagonist WRWWWW (5x10–6 M final concentration) were activated through an addition of Ac9–25 (50 µM final concentration). The cells were then challenged with the FPR agonist fMLF (10–7 M final concentration). The control cells (broken line) were preincubated in the absence of WRWWWW and Ac9–25 and challenged only with fMLF. The release of superoxide anions was measured by isoluminol-ECL. The time is given on the abscissa, and the amount of superoxide is on the ordinate given as Mcpm. The curves are from a representative experiment.
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Figure 9. The PIP2-binding peptide PBP10 is without effect on Ac9–25-mediated inhibition of superoxide release. Neutrophils were preincubated with (dotted line) or without (solid line) PBP10 (1 µM final concentration) for 5 min at 37°C and activated with Ac9–25 (50 µM final concentration) followed by fMLF (10–7 M final concentration). The amount of superoxide anion release, in the absence of Ac9–25, and the effects of PBP10 are shown for the FPRL1 agonist WKYMVM (lower left panel) and for the FPR agonist fMLF (lower right panel). The release of superoxide anions was measured by isoluminol-ECL. The time is given on the abscissa, and the amount of superoxide is on the ordinate given as Mcpm. The curves are from representative experiments.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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We used receptor-specific agonists and antagonists to identify the receptors involved in Ac9–25 signaling. The formylated peptide fMLF, a classical high-affinity ligand for the chemoattractant receptor FPR, was used together with the antagonists boc-MLF and cyclosporin H (reviewed in ref. [33 ]). FPRL1, originally defined as an orphan receptor, is a promiscuous receptor to which the peptide WKYMVM is one of the more potent high-affinity and specific ligands [16 ]. Recently, the peptide WRWWWW was shown to be a FPRL1-specific antagonist [23 ]. This antagonist inhibits several different FPRL1 agonists and even agonists that do not share identical binding domains in FPRL1 [23 , 34 ]. This peptide was used, together with WKYMVM, to evaluate the role of FPRL1 in neutrophil triggering by Ac9–25. Receptor desensitization experiments using agonists specific for FPR and FPRL1, respectively, strongly suggest that the signal for activation of the neutrophil NADPH-oxidase by Ac9–25 originates from FPR and not from FPRL1. This conclusion was further supported from experiments with the receptor-specific antagonists cyclosporin H and boc-MLF. These antagonists totally block the Ac9–25-induced respiratory burst, whereas WRWWWW was without any effect. Our results were finally confirmed using FPR-, FPRL1-, and FPRL2-expressing, undifferentiated HL-60 cells. Calcium mobilization experiments reveal that Ac9–25 triggers a response in FPR-expressing cells but not in those expressing FPRL1 or FPRL2. These results are not in agreement with earlier data presented using stably transfected HEK293 cells [8 ]. At the present time, we don’t know the reason for this, but the important finding is that our peptide retains the inhibitory effect but lacks the ability to activate through FPRL1.
A large number of ligands have been identified for FPRL1, including a number of peptides with microbial origin as well as the endogenous acute-phase reactant serum amyloid A [17 ] and the cathelicidin LL-37 [35 ]. For FPR, conversely, only a few ligands, in addition to the formylated peptides, have been identified [8 , 36 ]. It may, however, be that FPR is also promiscuous with respect to ligand binding, and more proteins/peptides will be added in the future.
The inhibitory activity induced by Ac9–25 on neutrophil superoxide release could rely on several different mechanisms; there may be a direct or indirect effect of Ac9–25 on the superoxide-generating electron-transporting system; the Ac9–25 peptide may compete with the activating agonist for binding to the specific surface receptor; or binding of Ac9–25 may switch the function of the receptor toward generation of a not-yet identified inhibitory signal [37 ]. The Ac9–25 peptide was without effect on the neutrophil NADPH-oxidase activity when the cells were triggered with PMA, a neutrophil activator that bypasses all cell-surface receptors and directly activates PKC. We can thus exclude the possibility that the peptide has a direct effect on the NADPH oxidase. The fact that the inhibitory signal affects the response, not only to fMLF but to all the G protein-coupled receptor-binding agonists tested also excludes the possibility that the Ac9–25 peptide competes with the activating agonists for binding to their respective receptor. The mechanism for inactivation is not known but might involve a general down-regulation of receptor signaling or a hierarchical cross-desensitization of receptor/signaling pathways.
Ligand occupation of FPR is associated with a rapid phosphorylation and/or cytoskeleton coupling of the occupied receptor, and this results in a physical separation of the receptor from the G protein and by that, a termination of the signaling capacity. This desensitization process makes the cells unable to generate a burst of superoxide if they are challenged with a new ligand that uses the same receptor or a hierarchically subordinated receptor (e.g., CXC chemokine receptor triggered by IL-8) [38 ]. This could be the mechanism by which Ac9–25 inhibits the response to the other chemoattractants, but the inhibitory signal could probably not be generated by FPR or FPRL1, as neither of these receptors is hierarchically superior to the other or to C5aR [38 , 39 ]. The suggestion that none of the FPR family members are involved gains further support from the results obtained with receptor-specific antagonists. The inhibitory signal generated by Ac9–25 was not reversed by cyclosporin H/boc-MLF or WRWWWW, antagonists for FPR and FPRL1, respectively. Moreover, HL-60 cells allowed to differentiate in the presence of fMLF and WKYMVm display a functional NADPH-oxidase but become "nonresponders" to all agonists (including Ac9–25) operating through FPR or FPRL1. Also in these cells, Ac9–25 exercises an inhibitory effect illustrated by the fact that the response to C5a is reduced in cells challenged with the annexin AI peptide. These data strongly suggest that the inhibitory signal induced by Ac9–25 is transduced through a receptor distinct from the members of the FPR family.
We thus suggest that the inhibitory signal induced by Ac9–25 is generated by a receptor distinct from FPR and FPRL1, and this not-yet identified receptor shares some functional similarities to the neutrophil histamine receptor and the LXA4receptor. Earlier studies have revealed that histamine inhibits the NADPH oxidase-dependent formation of superoxide in a way similar to Ac9–25 [40 ]. The receptor that transduces this inhibitory signal is the H2-type histamine receptor, but the precise signal transduction mechanism triggered by this receptor has not been defined. The fact that ranitidine, an antagonist specific for the H2receptor has no effect on the Ac9–25-induced inhibition of superoxide production (data not shown) suggests that this peptide and histamine work through different receptors. Also, LXA4 has been shown to inhibit neutrophil function through FPRL1 [20 ]. We have not been able to confirm these results, but we have shown that LXA4 triggers HL-60 cells expressing FPRL1 as well as those lacking this receptor [41 ], and our earlier data thus suggest that LXA4 has a receptor that is distinct from FPRL1.
In summary, some important features of the annexin AI-derived peptide Ac9–25 and its receptors are that the peptide activates the neutrophil NADPH oxidase through FPR but not through FPRL1, the peptide inhibits the neutrophil release of superoxide anions when triggered by other chemoattractants, and the peptide most likely inhibits cell function through a receptor distinct from FPR and FPRL1.
Received March 16, 2005; revised April 25, 2005; accepted May 2, 2005.
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