Mitochondria regulate neutrophil activation by generating ATP for autocrine purinergic signaling

Abstract

Polymorphonuclear neutrophils (PMNs) form the first line of defense against invading microorganisms. We have shown previously that ATP release and autocrine purinergic signaling via P2Y2 receptors are essential for PMN activation. Here we show that mitochondria provide the ATP that initiates PMN activation. Stimulation of formyl peptide receptors increases the mitochondrial membrane potential (Δψm) and triggers a rapid burst of ATP release from PMNs. This burst of ATP release can be blocked by inhibitors of mitochondrial ATP production and requires an initial formyl peptide receptor-induced Ca(2+) signal that triggers mitochondrial activation. The burst of ATP release generated by the mitochondria fuels a first phase of purinergic signaling that boosts Ca(2+) signaling, amplifies mitochondrial ATP production, and initiates functional PMN responses. Cells then switch to glycolytic ATP production, which fuels a second round of purinergic signaling that sustains Ca(2+) signaling via P2X receptor-mediated Ca(2+) influx and maintains functional PMN responses such as oxidative burst, degranulation, and phagocytosis.

Keywords: ATP; Calcium Signaling; Inflammation; Neutrophil; Purinergic Receptor.

FIGURE 1.

fMLP stimulation causes rapid cellular ATP release. A, purified human PMNs (10⁶/ml) were stimulated with fMLP for the indicated times and extracellular ATP was measured by HPLC. ATP release was calculated using non-stimulated cells as controls. Data shown are mean values ± S.D. (error bars) of n = 4–6 separate experiments. B, for real time visualization of ATP release, PMNs were incubated with the membrane-bound ATP probe 2-2Zn(II) (500 nm), stimulated with fMLP (10 nm), and fluorescence changes of 2-2Zn(II) were recorded over time using a fluorescence microscope (scale bar, 20 μm; see also supplemental Movie S1). Gray values were analyzed with ImageJ software, and extracellular ATP (eATP) concentrations were estimated based on the signal obtained with known ATP concentrations (supplemental Fig. 1; see also supplemental Movie S2). Data are expressed as mean ± S.D. (error bars) of n = 13 cells and are representative of n = 4 separate experiments with similar results. C, PMNs stained with 2-2Zn(II) were stimulated with fMLP (1 nm) and z-stack time lapse image sequences were acquired with spinning disk confocal microscopy. Arrows indicate sites of hot spots of 2-2Zn(II) signal and asterisks indicate the site of pseudopod protrusion during FPR-induced cell polarization (scale bar, 5 μm; see also supplemental Movie S3). Data shown are representative of four to six separate experiments with similar results. *, p < 0.05 versus time = 0 s.

FIGURE 2.

Mitochondria are required for ATP release. A, PMNs (10⁶/ml) were treated for 2 h with 2-deoxy-d-glucose (2DG) (20 mm) or sodium iodoacetate (SIA) (0.5 mm) to inhibit glycolysis (black bars) or with oligomycin (oligom.) (10 μm) or CCCP (10 μm) to inhibit mitochondrial ATP production (gray bars), and iATP was measured with HPLC (upper panel). In the lower panel, cells were treated with the indicated drugs for 2 h, stimulated with fMLP (100 nm) for 15 s, and extracellular ATP in bulk media was measured with HPLC. *, p < 0.05 versus untreated controls; #, p < 0.05 versus stimulated control without drug treatment. B, PMNs were treated with CCCP (10 μm) or KCN (1 mm) for 10 min, stimulated with the indicated concentrations of fMLP for 15 s, and iATP (upper panel) and ATP release into the supernatant (lower panel) were measured by HPLC. The amount of ATP released was calculated relative to non-stimulated cells that served as controls. #, p < 0.05 versus non-stimulated controls; *, p < 0.05 versus untreated but stimulated controls. Data are expressed as mean values ± S.D. (error bars) of n = 3 separate experiments. C and D, PMNs were treated with CCCP (10 μm) or not, stimulated with fMLP (10 nm), and ATP release was visualized with 2-2Zn(II) probe (500 nm). Extracellular ATP (eATP) concentrations were estimated as described in Fig. 1 (scale bar, 10 μm; see also supplemental Movie S4).

FIGURE 3.

FPR stimulation increases Δψm in PMNs. A, PMNs were stained with JC-1 (100 ng/ml) and stimulated with fMLP (10 nm) for the indicated times, and Δψm was analyzed by flow cytometry to assess the distribution of PMNs based on their red and green JC-1 fluorescence and the fluorescence ratio (red/green). Data shown are representative of similar results obtained in three separate experiments. B, to assess the time course of changes in Δψm following stimulation with fMLP (10 nm), mean fluorescence intensity value changes were normalized to the initial fluorescence intensity at time 0 (F/F₀). C, the dose dependence of changes in Δψm following stimulation of PMNs with the indicated fMLP concentrations for 15 s was assessed as described in B. *, p < 0.05 versus non-stimulated controls. Data are expressed as mean values ± S.D. (error bars) of n = 3 separate experiments.

FIGURE 4.

FPR stimulation triggers mitochondrial activity. A and B, PMNs were plated onto fibronectin-coated glass coverslips and stained with JC-1 (1 μg/ml; A) or with DHR 123 (2 μm; B), and mitochondrial membrane potential (A) and ROS production (B) were monitored in real time using fluorescence microscopy. JC-1 red and rhodamine 123 fluorescence changes in response to stimulation with fMLP (10 nm) were recorded over time. Cells treated with CCCP (10 μm) but not fMLP were included as controls. Data represent the means ± S.D. (error bars) of normalized gray values of 15–25 cells (scale bar, 5 μm; see also supplemental Movies S5–S7). C, mitochondrial colocalization with panx1 channels was assessed by expressing panx1-TurboGFP fusion protein in differentiated HL-60 cells, staining transfected cells with the Δψm-sensitive MitoTracker dye Red CM-H₂XRos, and stimulating cells with fMLP (10 nm). Green and red fluorescence time lapse image sets were acquired with spinning disk confocal microscopy, and colocalization of mitochondria (red) and panx1 (green) was assessed by image overlay using ImageJ software (scale bar, 5 μm; see also supplemental Movie S8). The asterisk indicates the direction of migration, and the arrow highlights a region of colocalization of panx1 and mitochondria. Data shown are representative of similar results obtained in three to six separate experiments. CCCP cont., unstimulated control cells treated with CCCP only.

FIGURE 5.

Mitochondria link purinergic signaling with Ca²⁺ mobilization. A and B, PMNs were treated or not with BAPTA-AM (10 μm) for 30 min and stimulated with fMLP (10 nm) for the indicated times and ATP release was measured with HPLC as described in Fig. 1A. Mitochondrial activation was assessed with JC-1 and fluorescence microscopy as described in Fig. 4B. C–F, to assess the role of mitochondria and purinergic signaling in FPR-induced Ca²⁺ signaling, PMNs were loaded with the Ca²⁺ probe Fluo-4 AM (5 μm), placed into the stirred cuvette of a fluorescence spectrophotometer, and Fluo-4 fluorescence changes were monitored over time. Then cells were treated for 5 min with CCCP (10 μm), EGTA (5 mm), a combination of EGTA + CCCP (10 μm and 5 mm, respectively), KCN (500 μm), rotenone (Roten.) (500 nm), oligomycin (Oligom.) (10 μm; in this case pretreatment for 45 min), carbenoxolone (CBX) (50 μm), or suramin (100 μm). Then the cells were stimulated with fMLP (10 nm), and changes in cytosolic Ca²⁺ levels were monitored for 30 min. For comparison, the Ca²⁺ response to fMLP of untreated PMNs is shown in C–F. Data are expressed as mean values ± S.D. (error bars) of three to six individual experiments.

FIGURE 6.

Mitochondria regulate functional PMN responses. A, to determine how mitochondrial ATP production contributes to the oxidative burst response, purified human PMNs were pretreated for 5 min with the indicated concentrations of CCCP (1–20 μm) and stimulated with fMLP (100 nm or the indicated concentrations), and oxidative burst was assessed after 20 min using DHR 123 and flow cytometry (left panel). The dose response to CCCP was calculated based on changes of the average mean florescence intensity (MFI) values of treated cells versus untreated controls (center panel), and the effect of CCCP (50 μm) on cells stimulated with the indicated concentrations of fMLP was determined by calculating the percentage of cells with increased rhodamine fluorescence (Gate 1; left panel). B, to assess how mitochondria contribute to PMN degranulation, cells were treated with fMLP and CCCP as described in A and fixed. CD11b expression on the cell surface was measured as a marker of degranulation. C, the role of mitochondrial ATP formation in bacterial phagocytosis was assessed by treating PMNs with the indicated concentrations of CCCP, adding Alexa Fluor 488-conjugated E. coli, and assessing bacterial uptake with flow cytometry. Data are shown as the percentage of PMNs with bacteria in Gate 1 as opposed to cells without bacteria or with bacteria adhered to their surface. Data are expressed as mean values ± S.D. (error bars) of three to six individual experiments performed ≥3 different times with similar results. *, p < 0.05 versus untreated but stimulated controls. cont., control cells exposed to bacteria for <10 s.

FIGURE 7.

Proposed role of mitochondria in PMN activation. Stimulation of FPRs triggers a first phase of Ca²⁺ mobilization, mitochondrial activation, and ATP production followed by rapid ATP release via panx1 channels. This initial burst in ATP release promotes a second round of Ca²⁺ signaling triggered by P2Y₂ receptor activation. A switch to ATP production via glycolysis results in further ATP release, activating P2X receptors that contribute to a third phase of Ca²⁺ signaling due to influx from the extracellular space. This second ATP signaling phase sustains intracellular Ca²⁺ levels and maintains functional PMN responses following FPR stimulation.