The regulation of neutrophil functions by Type I cGMP-dependent protein kinase (cGKI) was investigated in wild-type (WT) and cGKI-deficient (cGKI−/−) mice. We demonstrate that murine neutrophils expressed cGKIα. Similar to the regulation of Ca2+ by cGKI in other cells, there was a cGMP-dependent decrease in Ca2+ transients in response to C5a in WT, but not cGKI−/− bone marrow neutrophils. In vitro chemotaxis of bone marrow neutrophils to C5a or IL-8 was significantly greater in cGKI−/− than in WT. Enhanced chemotaxis was also observed with cGKI−/− peritoneal exudate neutrophils (PE-N). In vivo chemotaxis with an arachidonic acid-induced inflammatory ear model revealed an increase in both ear weight and myeloperoxidase (MPO) activity in ear punches of cGKI−/− vs WT mice. These changes were attributable to enhanced vascular permeability and increased neutrophil infiltration. The total extractable content of MPO, but not lysozyme, was significantly greater in cGKI−/− than in WT PE-N. Furthermore, the percentage of MPO released in response to fMLP from cGKI−/− (69%) was greater than that from WT PE-N (36%). PMA failed to induce MPO release from PE-N of either genotype. In contrast, fMLP and PMA released equivalent amounts of lysozyme from PE-N. However, the percentage released was less in cGKI−/− (∼60%) than in WT (∼90%) PE-N. Superoxide release (maximum velocity) revealed no genotype differences in responses to PMA or fMLP stimulation. In summary, these results show that cGKIα down-regulates Ca2+ transients and chemotaxis in murine neutrophils. The regulatory influences of cGKIα on the secretagogue responses are complex, depending on the granule subtype.

The neutrophil plays a critical role in immune defense. To be effective, the neutrophil is involved in a multistep process that leads it via multiple signals from the peripheral blood to sites of inflammation. These steps involve adhesion of circulating neutrophils to the vascular wall, diapedesis, and extravasation through the basement membrane toward sites of inflammation where it eliminates foreign microorganisms through phagocytosis, generation of toxic oxygen radicals, and release of microbicidal agents from granules (1, 2, 3).

Neutrophils are recruited to sites of inflammation by a variety of chemoattractants including complement factors (C5a), IL-8, arachidonic acid (AA)3 metabolites (leukotriene B4), and bacterial peptides (fMLP) that are generated and released locally at sites of injured tissue. Chemotactic receptors couple to heterotrimeric guanine nucleotide-binding proteins (G proteins) and elicit a wide range of responses in leukocytes including chemotaxis, respiratory burst, and granule secretion (4, 5, 6, 7). C5a, IL-8, and leukotriene B4 are classical chemoattractants, whereas fMLP functions as a secretagogue as well. Evidence is convincing that activation of chemotactic receptors increases the concentration of cytosolic calcium ([Ca2+]i) and cGMP (7, 8, 9, 10). Rat and human peripheral blood neutrophils constitutively express neuronal NO synthase (11, 12); whereas, inducible NO synthase activity is only detectable during bacterial infection (13). The findings that neutrophils generate NO (14, 15, 16), a molecule that stimulates guanylyl cyclase, and express type I cGMP-dependent protein kinase (cGKI) (17) prompted new interest in the NO/cGMP signaling pathway and its role in regulating neutrophil functions. The precise function of cGMP-elevating agents and NO donors in regulating neutrophil chemotaxis, granule secretion, and the respiratory burst is controversial (8, 10, 18, 19, 20, 21). NO has diverse effects on neutrophil functions, because a concentration-dependent biphasic effect has been reported for chemotaxis and granule secretion (22, 23, 24). Interestingly, high concentrations of NO-releasing compounds and cGMP analogues inhibit chemotaxis, whereas lower concentrations facilitate this response (20, 22, 23, 24). Evidence suggests that exogenous NO or l-Arg regulates chemotaxis and granule secretion via a cGKI-dependent mechanism (9, 22, 23, 24, 25). cGKI is an established downstream target of the NO/cGMP signaling cascade. For example, granule secretion of human neutrophils stimulated with fMLP or A23187 is associated with elevated cGMP levels and the phosphorylation of vimentin by cGKI (9, 10, 25). Similarly, during neutrophil adhesion and spreading, cGMP levels are elevated, and vasodilator-stimulated phosphoprotein (VASP), a membrane-associated focal adhesion protein, is phosphorylated by cGKI (26). Furthermore, when neutrophils are stimulated with LPS, p38 MAPK is phosphorylated upon accumulation of cGMP (27). These findings suggest that cGKI is a regulator of neutrophil functions. On the basis of studies of neutrophils and other cell types, cGKI may regulate receptor activation and signaling pathways downstream from receptor activation (see Refs.28, 29, 30, 31), which may have an impact on the cytoskeleton (9, 10, 25, 26, 32).

Part of the controversy regarding the signal transduction pathway for cGMP/cGKI in human neutrophils may be due to the reliance on pharmacological agonists and inhibitors. The specificity of these compounds has been challenged and is questionable (33, 34). Rodent neutrophils respond to chemoattractant signaling, secrete granule contents, generate a respiratory burst, generate NO, and kill microorganisms in a manner that is both qualitatively and quantitatively similar to human neutrophils (35, 36). Therefore, to gain further insight into the regulatory role of cGKI in neutrophils, we used bone marrow-derived neutrophils and extravasated neutrophils from wild-type (WT) and cGKI-deficient (cGKI−/−) mice to investigate the impact of cGKI on neutrophil function. The results suggest that cGKI regulates mouse neutrophil chemotaxis and granule secretion, and that, as with other cell types, cGKI lowers agonist-induced increases in [Ca2+]i. In addition, cGKI may dampen the respiratory burst over time. These findings strengthen the notion and confirm previous work that cGKI is an important regulator of neutrophil functions.

The following Abs were purchased from BD Biosciences Pharmingen: FITC-rat anti-mouse CD11b (clone M1/70), FITC-anti-rat IgG2b (isotype control, clone A95-1); R-PE-rat anti-mouse TER-119/erythroid cells (Ly-76), PE-rat anti-mouse Ly-6G (Gr-1), and Ly6C (RB6-8C5); and cytochrome c rat anti-mouse CD45 (Ly-5, clone 30-F11), anti-CD16/CD32 (Mouse Fc Block, clone 2.4G2).

The generation of the cGKI-knockout (KO) mouse has been described previously (37). The cGKI−/− mice were kept on the 129/Sv background and were further backcrossed either to BALB/c or to C57BL/6NN background for nine generations. The animals were kept under standard housing conditions (21 ± 1°C, 55 ± 10% relative humidity, 12-h dark-light cycle, pathogen free). All investigations were performed on 4- to 6-wk-old mice that were housed in standard (type 2) hanging rodent cages (Ehret) or static microisolator cages (Ehret) with food and water ad libitum. If not otherwise indicated, in most experiments 7–10 mice per group were studied per strain (129/Sv, BALB/c, C57BL/6NN). The animal experiments were conducted in accordance with the European Communities Council Directive of November 24th 1986 and were approved by the Government of Upper Bavaria, Germany. In addition, United States of America animal experiments were conducted in accordance with the National Research Council Guide for the Care and Use of Laboratory Animals and were approved by the University of North Carolina’s Institutional Animal Care and Use Committee.

Mice (strain 129/Sv and BALB/c) received injections i.p. with 0.5 ml of 3% thioglycollate. After 4 h, the peritoneal cavities were lavaged with HBSS supplemented with 0.5 mM EDTA, and the peritoneal exudate cells were collected by low-speed centrifugation. The percentage of neutrophils was determined by differential count. Briefly, cytospins were stained with Diff-Quick (Dade Behring) and identified as neutrophils, eosinophils, macrophages, and lymphocytes by standard morphology. Cell counts were obtained in five random fields, each containing ∼100 cells using ×100 oil immersion objective. Typically, 75–80% neutrophils were recovered. The cells were resuspended in Gey’s balanced salt solution (GBSS) supplemented with 0.1% BSA and 20 mM HEPES and were used in the assays.

BMC of mice (129/Sv and C57BL/6NN) were flushed from femurs with ice-cold HBSS supplemented with 0.1% BSA and 2 mM EDTA. The bone marrow was freed of RBC by hypotonic lysis (62 mM NH4Cl, 20 mM NaHCO3, and 1 mM EDTA), and the neutrophil population was positively selected by FACS using FITC anti-mouse Gr1 (BD Pharmingen). The FACS-sorted, anti-Gr1-positive leukocyte population was composed of 97% neutrophils. Cell viability was 98%, as determined by trypan blue exclusion. Greater than 99% of the neutrophils were spherical, suggesting that the neutrophils were not activated during isolation.

Cell suspensions of mouse peripheral blood were stained with PE-anti-mouse CD42d Ab (BD Pharmingen) and positively selected by FACS. The population of cells was >98% platelets.

Cell suspensions of bone marrow and peripheral blood cells (strain BALB/c) were stained as follows: PE-anti-TER 199 for detection of immature erythroid cells; cytochrome c anti-CD45 for detection of total leukocytes; FITC anti-CD11b for detection of macrophages; and PE-anti-Gr1 for detection of neutrophils. Samples were analyzed by FACS with CellQuest version 3.2.1 f1 1998 software (BD Biosciences). Peripheral blood leukocyte counts were obtained by using Beckman Coulter AcT diff with veterinary software (Beckman Coulter), and differentials were counted manually.

Neutrophil cell pellets (C57BL/6NN) were freeze-thawed three times in liquid nitrogen and solubilized with sample buffer (0.2 M Tris, 5% SDS, 40% glycerine, 0.004% bromphenol blue, and 50 mM DTT). After SDS-PAGE, the proteins were transferred to polyvinylidene fluoride membranes. Membranes were blocked with 5% skim milk in TBST, and then incubated with 1/1000 rabbit anti-cGKIα or 1/100 rabbit anti-cGKIβ for 1 h at room temperature. The membranes were washed in TBST and incubated with 1/10,000 goat anti-rabbit IgG-HRP (Dianova) for 1 h at room temperature. The wash was repeated, and cGKIα and cGKIβ were detected by chemiluminescence (Amersham Biosciences). The polyclonal Abs specific for the isoforms cGKIα and cGKIβ were raised against the recombinant proteins expressed in bacteria (38).

Cytospins (129/Sv) of BMCs were fixed for immunofluorescence microscopy as described previously (10) and stained with affinity purified rabbit anti-cGKI common peptide (35-B-5-3) at 1:25 for 1 h (39). Thereafter, cells were incubated for 30 min with FITC-goat anti-rabbit IgG (Rockland). Cells were viewed on a Leitz fluorescence microscope, and immunofluorescence micrographs were acquired with an Optronics Engineering DEI-470 color video camera (Optronics International).

PE-N or BMC-derived neutrophils of 3- to 4-wk-old mice (strain 129/Sv) were used for these studies. Cells were resuspended at a concentration of 1 × 106 cells/ml in GBSS/20 mM HEPES/0.1% BSA, and chemotaxis in response to C5a (1 nM to 1 μM) or IL-8 (10–1000 ng/ml) was measured using a 48-well chemotaxis chamber with a 5-μm pore size polycarbonate membrane filter (NeuroProbe). The C5a- or IL-8-stimulated cells were incubated at 37°C for 30 or 60 min, respectively. The filters were stained with Diff-Quick (Dade Behring), and the number of cells in five random fields was counted using ×100 oil immersion objective.

The ears of 4- to 5-wk-old BALB/c mice were painted with 10 μl of (200 mg/ml) AA in acetone (left ear) and 10 μl of acetone (right ear) to serve as a control. The animals were sacrificed after 1.5 h, and punches of each ear lobe were obtained using a sterile 5-mm AcuPunch (Acuderm). The ear punches were immediately weighed and cut into small pieces for myeloperoxidase (MPO) analysis. The remaining ear was fixed in 3.7% formalin in PBS for histology.

MPO was extracted from homogenized ear punches by suspending the material in 0.5 ml of 0.5% hexadecyltrimethylammonium bromide in 50 mM potassium phosphate buffer (pH 6.0) before sonication in an ice bath for 10 s (Heat Systems; Ultrasonics). The specimens were freeze-thawed three times, after which sonication was repeated. Suspensions were centrifuged at 40,000 × g for 15 min, and 50 μl of the resulting supernatant was measured for MPO activity (see Measurement of MPO).

PE-N from BALB/c mice were used in this study. A microtiter assay measured the superoxide dismutase-inhibitable reduction of ferricytochrome c (an indirect measure of superoxide anion production) by peritoneal exudate cells (BALB/c) in response to either 100 ng/ml PMA or 1 μM fMLP in the presence of cytochalasin B (CB) as described previously (40). The change in OD550 was monitored kinetically using a Molecular Devices Vmax Kinetic Microplate Reader (Molecular Devices). The OD was recorded every 10 s following a 3 s shake. The Vmax was calculated using the slope of the ΔOD over the first 3 min for the fMLP response and through the maximum number of points before substrate exhaustion (generally 30 min) with PMA. The correlations for the curve fittings all exceeded 0.99. An extinction coefficient of 21.1 mM−1 at 550 nm was used for ferricytochrome c, and superoxide production was expressed as nanomoles of reduced cytochrome per 106 cells per min. The area under the curve (AUC) through 60 min was calculated as an estimate of the sustained production of superoxide in response to fMLP.

PE-N were suspended at 1 × 107 cell/ml in GBSS containing 0.1% BSA. Cells (40 μl) were placed in a 96-well microtiter plate (Falcon) containing 160 μl of GBSS plus 0.1% BSA in the presence or absence of DMSO (1:2000) or 10 μM CB. The plate was placed in a circulating water bath at 37°C for 5 min. The cells were then stimulated with 1 μM fMLP or 20 ng/ml PMA for 15 min. The plate was centrifuged for 10 min at 4°C at 1000 rpm, and the culture supernatant was removed and placed in tubes on ice containing 1 μl of protease inhibitor mixture. The supernatant was used to measure MPO and lysozyme activity. For total enzyme activity within a population of cells, the cells were lysed on ice with 0.1% Triton X-100 containing protease inhibitor mixture. The cells were vortexed and microfuged to remove debris. The supernatant was used to measure MPO and lysozyme activity.

MPO activity of supernatants (50 μl) was measured in a microwell assay using 3,3′,5,5′-tetramethylbenzidine (70 μl) as a peroxidase substrate (Kirkegaard & Perry Laboratories). Human MPO (Athens Research & Technology) was used as a standard. The plate was read at OD650 after 15 min and was monitored kinetically using a Molecular Devices Vmax Kinetic Microplate Reader (Molecular Devices). The standard curves were fitted using the software program InPlot (GraphPad). Data are reported as units MPO/106cells released for secretion studies, and units MPO/ml for in vivo chemotaxis assays. In addition, the percentage of MPO released was calculated by dividing the amount of enzyme released by the total amount of enzyme extracted from the nonstimulated neutrophil population.

Lysozyme activity of supernatants (25 μl) was measured in a microwell assay using Micrococcus lysodeikticus (125 μl) as a substrate. A standard curve was performed using egg white lysozyme. The hydrolysis of M. lysodeikticus was monitored kinetically for 1 h at OD450 using a Molecular Devices Vmax Kinetic Microplate Reader (Molecular Devices). The standard curves were fitted using the software program InPlot (GraphPad). Data are reported as micrograms of lysozyme/106 cells. In addition, the percentage of lysozyme released was calculated by dividing the amount of enzyme released by the total amount of enzyme extracted from the nonstimulated neutrophil population.

Neutrophils (strain 129/Sv) were isolated from bone marrow and purified by FACS (as described above). Ca2+ transients were measured at room temperature in single cells loaded with the Ca2+ indicator fura 2-AM (1 μM). Ca2+ transients were elicited two times in succession by local application of C5a (0.1 μM). Cells were incubated for 5 min in the absence or presence of 8-bromo-cGMP (8-Br-cGMP) (1 μM–1 mM) before the second Ca2+ transient was elicited. Images were recorded with the vision 4.0 fluorescence imaging software program (T.I.L.L. Photonics), and the AUC was integrated using the software program Microcal Origin 6.0 (Microcal Software). Values were calculated by dividing the AUC of the second transient by the AUC of the first transient.

Data are given as mean ± SEM of n experiments. Statistical significance of differences was analyzed by nonpaired or paired, two-tailed Student’s t test. A p value <0.05 was considered significant. All statistical calculations were performed with the InStat program (GraphPad).

HBSS and GBSS were purchased from (Invitrogen Life Technologies). Unless indicated otherwise, all other chemicals were obtained from Sigma-Aldrich or from Sigma Chemical.

In agreement with our early findings in human neutrophils (17), FACS-sorted murine bone marrow neutrophils (BM-N) express cGKI (Fig. 1). Immunoblot analysis revealed that mouse neutrophils express the α isoform of cGKI (Fig. 1,A). In contrast to murine platelets, which express both the α and β isoforms of cGKI, no immunoreactivity for cGKIβ was found in neutrophils (Fig. 1 B). Neither cGKIα nor cGKIβ was detected in neutrophils or platelets of cGKI-deficient mice.

FIGURE 1.

Expression of cGKIα in FACS-purified bone marrow-derived neutrophils. A and B, Western blots of cell extracts (60 μg/lane) of neutrophils and platelets from WT and cGKI−/− mice are shown for cGKIα (A) and cGKIβ (B). Recombinant cGKIα and cGKIβ proteins (10 ng per lane) demonstrate the specificity of the Iα and Iβ Abs. The experiments were repeated at least three times with different sets of mice. The FACS-sorted cell populations of neutrophils and platelets were >95% pure.

FIGURE 1.

Expression of cGKIα in FACS-purified bone marrow-derived neutrophils. A and B, Western blots of cell extracts (60 μg/lane) of neutrophils and platelets from WT and cGKI−/− mice are shown for cGKIα (A) and cGKIβ (B). Recombinant cGKIα and cGKIβ proteins (10 ng per lane) demonstrate the specificity of the Iα and Iβ Abs. The experiments were repeated at least three times with different sets of mice. The FACS-sorted cell populations of neutrophils and platelets were >95% pure.

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Immunofluorescence microscopy of BMCs demonstrated that cGKI was expressed in neutrophils and megacaryocytes (Fig. 2,A). No staining was observed in myeloid cells or megacaryocytes of cGKI−/− animals (Fig. 2,B). Fine granular staining of cGKI was observed in the cytoplasm of neutrophils (Fig. 2 C). Staining was observed only in permeabilized cells, suggesting that cGKI is not localized on the plasma membrane.

FIGURE 2.

A, Staining of cGKI of BMCs was observed in permeabilized neutrophils (arrows) and megacaryocytes (m; control). B, No staining was observed in myeloid cells or megakaryocytes of cGKI−/− animals. C, Fine granular staining for cGKI was observed in the cytoplasm of neutrophils. The rabbit Ab used for this experiment recognized the cGKIα and cGKIβ isoforms. Magnification, ×125 for A and B and ×500 for C.

FIGURE 2.

A, Staining of cGKI of BMCs was observed in permeabilized neutrophils (arrows) and megacaryocytes (m; control). B, No staining was observed in myeloid cells or megakaryocytes of cGKI−/− animals. C, Fine granular staining for cGKI was observed in the cytoplasm of neutrophils. The rabbit Ab used for this experiment recognized the cGKIα and cGKIβ isoforms. Magnification, ×125 for A and B and ×500 for C.

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Leukocyte lineages were examined in peripheral blood and bone marrow of 4-wk-old WT and cGKI−/− littermates (Fig. 3). cGKI−/− mice had significantly more peripheral blood leukocytes (WBC) than WT controls (13.73 ± 2.08 × 103/μl vs 8.33 ± 0.82 × 103/μl; n = 9; ∗, p < 0.05). Interestingly, FACS analyses revealed a disproportionate increase in total numbers of neutrophils in the cGKI−/− animals that resulted in a significant increase in the percentage of neutrophils at the expense of the lymphocytes (Fig. 3,A). No significant differences in the percentage of monocytes, basophils, or eosinophils were observed between the two animal lines on blood smears stained with Wright-Giemsa or diaminobenzidine (MPO detection). In addition, all of the WBC exhibited normal morphology on blood smears. Examination of BMCs by FACS revealed no significant differences in the percentage of myeloid, lymphoid, or monocytoid lineages (Fig. 3 B). Thus, the cGKI−/− animals present with a granulocytosis in the peripheral blood.

FIGURE 3.

FACS of WT and cGKI−/− leukocytes. The percentage of WT (□) and cGKI−/− (▪) gated cells by FACS is shown of peripheral blood (A) and bone marrow (B) for monocytes (MN), polymorphonuclear leukocytes (PMN), and lymphocytes (LYM). No significant differences were observed in BMCs. However, a significant increase in WBC was observed in cGKI−/− peripheral blood that resulted in a significant increase in the percentage of neutrophils. Peripheral blood, WT (n = 4), and cGKI−/− (n = 3); bone marrow, WT (n = 7), and cGKI−/− (n = 6). Four week-old WT and cGKI−/− littermate mice (BALB/c) were used. ∗, Significance from WT at p < 0.05.

FIGURE 3.

FACS of WT and cGKI−/− leukocytes. The percentage of WT (□) and cGKI−/− (▪) gated cells by FACS is shown of peripheral blood (A) and bone marrow (B) for monocytes (MN), polymorphonuclear leukocytes (PMN), and lymphocytes (LYM). No significant differences were observed in BMCs. However, a significant increase in WBC was observed in cGKI−/− peripheral blood that resulted in a significant increase in the percentage of neutrophils. Peripheral blood, WT (n = 4), and cGKI−/− (n = 3); bone marrow, WT (n = 7), and cGKI−/− (n = 6). Four week-old WT and cGKI−/− littermate mice (BALB/c) were used. ∗, Significance from WT at p < 0.05.

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Increased calcium transients are a hallmark of the neutrophil response to agonists (7, 41) and are modulated by cGKI in other cells (28, 42). To determine whether cGKI modulates calcium fluxes in neutrophils, we measured [Ca2+]i transients in neutrophils that were stimulated with C5a. We used bone marrow-derived neutrophils to ensure that the cells were minimally activated, and chose C5a as an agonist because it induces a rise in intracellular calcium levels through a classic G protein-linked receptor.

In fura 2-loaded BM-Ns, C5a (0.1 μM) elicited [Ca2+]i transients in both WT and cGKI−/− neutrophils that reached a peak level within 1 min and returned to baseline levels within 5 min (Fig. 4,A). A second exposure to C5a resulted in [Ca2+]i transients of similar magnitude in neutrophils from both genotypes (Fig. 4,A). In contrast, when WT neutrophils were preincubated before the second application of C5a with 8-Br-cGMP, an activator of cGKI, the size of the second transient was reduced to 68% of the first peak (Fig. 4, B and C). The reduction in [Ca2+]i transients in WT neutrophils by 8-Br-cGMP was effective at concentrations as low as 1 μM, supporting the specificity of 8-Br-cGMP for cGKI. In contrast, there was no reduction in [Ca2+]i induced by exogenous 8-Br-cGMP in cGKI-deficient neutrophils (Fig. 4, B and C). These data are consistent with that of other cell types (29, 37, 42). They support a cGKIα-dependent regulation resulting in decreased agonist-induced Ca2+ transients in neutrophils.

FIGURE 4.

Comparison of calcium transients in WT and cGKI−/− neutrophils. Bone marrow-derived neutrophils from WT (□) and cGKI−/− (▪) mice were loaded with fura 2-AM for 45 min. [Ca2+]i transients were elicited by applying 0.1 μM C5a to adhered cells. Cells were incubated for 5 min with buffer (A) or 1 mM 8-Br-cGMP (B) before the second application of C5a. Summary of the results (C) is given as relative AUC (ratio of the second to first transient) for WT cells (control, n = 15; +8-Br-cGMP, n = 16) and cGKI−/− cells (control, n = 42; +8-Br-cGMP, n = 24). ∗, p < 0.02; ∗∗, p < 0.01.

FIGURE 4.

Comparison of calcium transients in WT and cGKI−/− neutrophils. Bone marrow-derived neutrophils from WT (□) and cGKI−/− (▪) mice were loaded with fura 2-AM for 45 min. [Ca2+]i transients were elicited by applying 0.1 μM C5a to adhered cells. Cells were incubated for 5 min with buffer (A) or 1 mM 8-Br-cGMP (B) before the second application of C5a. Summary of the results (C) is given as relative AUC (ratio of the second to first transient) for WT cells (control, n = 15; +8-Br-cGMP, n = 16) and cGKI−/− cells (control, n = 42; +8-Br-cGMP, n = 24). ∗, p < 0.02; ∗∗, p < 0.01.

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In vitro chemotaxis.

Chemotaxis of FACS-purified BM-Ns of WT and cGKI−/− littermates was investigated using the prototypic chemoattractants, IL-8 and C5a. Chemotaxis of BM-Ns to IL-8 and C5a was dose dependent, with the highest chemotactic response for both WT and cGKI−/− cells at 1 μg/ml IL-8 and 0.01 μM C5a (Fig. 5, A and B). Chemotaxis was enhanced significantly (p < 0.05) in cGKI−/− BM-Ns compared with that of the WT littermate controls by 1 μg IL-8 and 0.1 μM C5a. There were no significant differences in random migration and chemokinesis between genotypes.

FIGURE 5.

Comparison of chemotaxis of WT and cGKI−/− bone marrow-derived and peritoneal exudate neutrophils. Chemotaxis was dose dependent for IL-8 (A) and C5a (B) for WT □ and cGKI−/− ▪ bone marrow-derived neutrophils. Chemotaxis was significantly enhanced in cGKI−/− BM-Ns to IL-8 at 1 μg/ml (A) and to C5a at 0.1 μM (B). Similar to bone marrow-derived neutrophils, chemotaxis was dose dependent to C5a for WT and mutant PE-N (C). Chemotaxis was significantly enhanced in peritoneal exudate cGKI−/− neutrophils from 1 to 0.01 μM C5a. A, n = 3; B, n = 4; C, n = 2 WT and cGKI−/−. Incubation time was 30 min for C5a and 60 min for IL-8. All cells were obtained from litter-matched 129/Sv animals. Asterisks indicate a significant difference from WT littermate mice. ∗, p < 0.05; ∗∗, p < 0.01.

FIGURE 5.

Comparison of chemotaxis of WT and cGKI−/− bone marrow-derived and peritoneal exudate neutrophils. Chemotaxis was dose dependent for IL-8 (A) and C5a (B) for WT □ and cGKI−/− ▪ bone marrow-derived neutrophils. Chemotaxis was significantly enhanced in cGKI−/− BM-Ns to IL-8 at 1 μg/ml (A) and to C5a at 0.1 μM (B). Similar to bone marrow-derived neutrophils, chemotaxis was dose dependent to C5a for WT and mutant PE-N (C). Chemotaxis was significantly enhanced in peritoneal exudate cGKI−/− neutrophils from 1 to 0.01 μM C5a. A, n = 3; B, n = 4; C, n = 2 WT and cGKI−/−. Incubation time was 30 min for C5a and 60 min for IL-8. All cells were obtained from litter-matched 129/Sv animals. Asterisks indicate a significant difference from WT littermate mice. ∗, p < 0.05; ∗∗, p < 0.01.

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Chemotaxis of PE-N was also investigated for comparison. The PE-N of both genotypes were more responsive than bone marrow-derived neutrophils to C5a based upon the number of cells that migrated in 30 min. Similar to bone marrow-derived neutrophils, chemotaxis of PE-N to C5a was dose dependent for both WT and cGKI−/− cells with the maximal chemotactic response at 0.01 μM C5a for WT and cGKI−/− cells (Fig. 5,C). Chemotaxis of cGKI−/− neutrophils was significantly increased when compared with that of neutrophils from WT littermate controls over a range of concentrations from 0.01 to 1 μM C5a (Fig. 5 C). There was minimal random migration. Migration of cells that were incubated in the presence of 0.1 μM C5a (chemokinesis) was similar in WT and cGKI−/− neutrophils (data not shown). In conclusion, chemotaxis was greater in cGKI−/− animals than their WT littermate controls with both quiescent neutrophils (bone marrow-derived) and elicited neutrophils (peritoneum-derived).

In vivo chemotaxis.

To determine whether chemotaxis is also enhanced in vivo, we used an AA-induced inflammatory ear model. When applied topically, AA induces an acute inflammatory response involving both vascular leakage and cellular components that are highly dependent on leukotrienes (35). A significant increase in ear weight was observed in cGKI−/− mice compared with WT controls at 1.5 h after topical AA treatment (7.89 ± 0.15 vs 4.85 ± 0.26 mg, respectively; p < 0.001) (Fig. 6 A). A negligible response was observed in the vehicle control ears with no differences between the genotypes. This increase in ear weight in the cGKI−/− mice after AA treatment indicates enhanced vascular permeability in the cGKI−/− mice.

FIGURE 6.

Acute inflammatory response to topical AA in WT and cGKI−/− mice. The ears of BALB/c WT □ and cGKI−/− ▪ mice were analyzed for edema (ear weight; A) and for MPO content, an index of neutrophil influx (B). Differences in weight and MPO content between AA-treated and control (acetone) ear punches were measured. A significant increase in ear weight and MPO content was observed in cGKI-deficient mice. Asterisks indicate a significant difference from WT mice ∗∗∗, p < 0.001; n = 7.

FIGURE 6.

Acute inflammatory response to topical AA in WT and cGKI−/− mice. The ears of BALB/c WT □ and cGKI−/− ▪ mice were analyzed for edema (ear weight; A) and for MPO content, an index of neutrophil influx (B). Differences in weight and MPO content between AA-treated and control (acetone) ear punches were measured. A significant increase in ear weight and MPO content was observed in cGKI-deficient mice. Asterisks indicate a significant difference from WT mice ∗∗∗, p < 0.001; n = 7.

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The MPO activity can be used as an indicator of the cellular response to this acute inflammatory stimulant. An increase in MPO activity is accepted as an indicator of increased number of neutrophils in the inflamed tissue (35). MPO activity was negligible in the ears of both animals after topical treatment with acetone (vehicle). However, after AA stimulation, MPO activity was significantly increased in the cGKI−/− mice compared with control mice (4.42 ± 0.31 vs 1.16 ± 0.11 U MPO/ml lysate; p < 0.001) (Fig. 6,B). Histological examination of the tissues indicated that the neutrophil was the predominant cell type elicited in response to AA treatment in both WT and cGKI−/− mice (Fig. 7). Thus, the increased MPO activity in the inflamed tissue was associated with an enhanced influx of neutrophils and not monocytes or eosinophils. The significant increase in MPO concentration in the tissues of the cGKI−/− mice may be due to increased numbers of recruited neutrophils, consistent with the in vitro chemotaxis data, but also may be associated with the relatively increased MPO content of extravasated cGKI−/− neutrophils (see below).

FIGURE 7.

Histology of ears after topical application of AA in WT and cGKI−/− mice. Histological sections of ears after topical application of acetone (control) (A and B) and AA (C and D) are shown in WT (A and C) and cGKI−/− (B and D) mice. Samples given are representative for each group of seven WT and seven cGKI−/− animals, from experiments performed on three different days. An influx of neutrophils was observed after AA treatment in both WT and cGKI−/− ears. Magnification, ×40.

FIGURE 7.

Histology of ears after topical application of AA in WT and cGKI−/− mice. Histological sections of ears after topical application of acetone (control) (A and B) and AA (C and D) are shown in WT (A and C) and cGKI−/− (B and D) mice. Samples given are representative for each group of seven WT and seven cGKI−/− animals, from experiments performed on three different days. An influx of neutrophils was observed after AA treatment in both WT and cGKI−/− ears. Magnification, ×40.

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The effects of deleting the cGKI gene on the regulation of granule secretion and respiratory burst (below) were performed using elicited PE-N. The percentage of neutrophils in the harvested exudates was not significantly different between WT and cGKI-deficient cells. We used fMLP for these studies because it is an established activator of both granule secretion and the respiratory burst in mouse and human neutrophils, and similar to C5a and IL-8, fMLP binds to a G protein-coupled receptor for activation (43, 44). Comparative studies of fMLP with PMA, a direct activator of protein kinase C (PKC), were performed using the same population of PE-N from each animal. We used lysozyme to follow granule secretion in general, because it is packaged in primary, secondary, and tertiary granule types (45). In addition, MPO was used as a marker to measure the secretion of primary (azurophilic) granules, which are released by fMLP but not PMA (PKC-independent). We calculated the amount and the percentage of enzyme released.

For these studies, the PE-N were preincubated with CB and then stimulated with 1 μM fMLP for two reasons. First, granule secretion was not measurable in either the WT or the cGKI−/− PE-N with fMLP in the absence of CB or with CB alone. Second, whereas actin is not a substrate for cGKI, the intermediate filament protein vimentin is phosphorylated by cGKI in human neutrophils activated with fMLP (9). We have shown that cytochalasins alter the organization of both microfilaments and intermediate filaments during granule secretion (10). Therefore, CB offers an opportunity to directly investigate the effects of deleting the cGKI gene on granule secretion independent of the cytoskeleton.

The granule content of the PE-N of cGKI−/− animals was different from their WT littermates (Tables I and II). Although the total extractable lysozyme was similar between animals, there was more than a 2-fold increase in the total MPO extracted from the cGKI−/−-elicited peritoneal exudate cell population compared with that of WT (p < 0.05). fMLP/CB stimulated the release of MPO and lysozyme by WT and cGKI−/− peritoneal exudate cells. There was more than a 4-fold increase in the quantity of MPO released by cGKI−/− peritoneal exudate cells (Table I). This increase was not simply due to the increased availability of MPO content, because there was also a significant increase in the percentage of MPO released from peritoneal exudate cells of cGKI−/− (69 ± 10%) vs that of WT (36 ± 4%) animals. Consistent with primary granule packaging, MPO was not released in response to PMA with peritoneal exudate cells from either genotype (Table I).

Table I.

Secretion of MPO by WT and cGK1−/− neutrophils stimulated with fMLP or PMA

Total MPO (U/106 cells)fMLPPMA
MPO (U/106 cells)% MPOMPO (U/106 cells)% MPO
WT 0.786 ± 0.103a 0.264 ± 0.031a 36 ± 4a n.d.b n.d.b 
cGKI−/− 1.681 ± 0.42 1.154 ± 0.351 69 ± 10 n.d.b n.d.b 
Total MPO (U/106 cells)fMLPPMA
MPO (U/106 cells)% MPOMPO (U/106 cells)% MPO
WT 0.786 ± 0.103a 0.264 ± 0.031a 36 ± 4a n.d.b n.d.b 
cGKI−/− 1.681 ± 0.42 1.154 ± 0.351 69 ± 10 n.d.b n.d.b 
a

, p < 0.05 between WT and cGKI−/−; n = 8 WT and seven cGKI animals.

b

n.d., not detectable.

Table II.

Secretion of lysozyme by WT and cGKI−/− neutrophils stimulated with fMLP or PMA

Total Lysozyme (μg/106 cells)fMLPPMA
Lysozyme (μg/106 cells)% lysozymeLysozyme (μg/106 cells)% lysozyme
WT 2.850 ± 0.536 2.590 ± 0.539 90 ± 5a 2.500 ± 0.505 86 ± 4a 
cGKI−/− 4.256 ± 0.916 2.793 ± 0.753 63 ± 7 2.622 ± 0.719 61 ± 7 
Total Lysozyme (μg/106 cells)fMLPPMA
Lysozyme (μg/106 cells)% lysozymeLysozyme (μg/106 cells)% lysozyme
WT 2.850 ± 0.536 2.590 ± 0.539 90 ± 5a 2.500 ± 0.505 86 ± 4a 
cGKI−/− 4.256 ± 0.916 2.793 ± 0.753 63 ± 7 2.622 ± 0.719 61 ± 7 
a

p < 0.05 between WT and cGKI−/−; no significance in lysozyme secretion between fMLP and PMA in WT or cGKI−/− neutrophils; n = 8 WT and seven cGKI animals.

In contrast to MPO, PMA stimulated the release of lysozyme from the elicited peritoneal exudate cell populations of both genotypes (Table II), consistent with secondary granule packaging. Furthermore, similar to PMA, stimulation with fMLP/CB yielded equivalent quantities of released lysozyme. Lysozyme released as a percentage of the total available lysozyme revealed an interesting difference between the two genotypes. Virtually all of the lysozyme of WT PE-N was released after stimulation with fMLP (90 ± 5%) or PMA (86 ± 4%). Conversely, significantly less lysozyme, 63 ± 7% (fMLP/CB) and 61 ± 7% (PMA), was exocytosed by cGKI−/− PE-N (p < 0.01).

The effects of the deletion of the cGKI gene on the respiratory burst were performed using PE-N. As with the granule secretion studies, the PE-N were preincubated with CB and then stimulated with 1 μM fMLP. Comparative studies of fMLP with PMA were performed using the same populations of neutrophils. There is an initial response to fMLP that is sustained for ∼4 min, followed by a rapid decline in the rate of superoxide production. To determine the initial rate, the Vmax was determined using the values in the first 3 min and the Vmax software of Molecular Devices. The AUC over 60 min was calculated using the Molecular Devices software to capture the sustained production of superoxide. In contrast to fMLP, PMA activates NADPH oxidase through direct activation of PKC, resulting in a constant rate of cytochrome c reduction that is limited only by the availability of substrate. Therefore, the PMA-induced NADPH oxidase activity was estimated as a Vmax using the maximum number of data points before substrate limitation. Because AUC resulting from PMA stimulation would be directly proportional to the Vmax, AUC for PMA response was not a meaningful calculation. The elicited peritoneal exudate cell populations from both WT and cGKI−/− gave similar kinetics in response to both fMLP/CB and PMA.

As with other studies, there was no measurable respiratory burst response with fMLP in the absence of CB or with CB alone (no fMLP) with either the WT or the cGKI−/− PE-N. After fMLP/CB stimulation, there was an initial rate that was sustained for ∼4 min (Vmaxinit) followed by a reduced rate through 60 min. The kinetics of the response from 0 to 60 min was captured by transforming the data to the AUC. As can be seen in Table III, there was a significant difference in the sustained release of O2 (AUC) in WT vs cGKI−/− PE-N. In contrast, the differences in the Vmaxinit values of the WT vs the cGKI−/− PE-N responses to fMLP failed to reach significance (p = 0.29).

Table III.

Comparison of cytochrome c reduction by WT and cGKI−/− PE-N

fMLPPMA
Vmax initial (ΔOD/min)AreaVmax initial (ΔOD/min)
WT 3.99 ± 0.26 250 ± 29a 7.35 ± 2.45 
cGKI−/− 5.67 ± 1.36 392 ± 33 13.79 ± 3.57 
fMLPPMA
Vmax initial (ΔOD/min)AreaVmax initial (ΔOD/min)
WT 3.99 ± 0.26 250 ± 29a 7.35 ± 2.45 
cGKI−/− 5.67 ± 1.36 392 ± 33 13.79 ± 3.57 
a

p < 0.01 between WT and cGKI−/−; AUC was calculated from 0 to 60 min; n = 6 WT and six cGKI−/− animals.

The combination of these data is consistent with the cGKI−/− cells having a greater sustained production of superoxide than that of WT. This suggests that cGKI serves to dampen the second phase of the respiratory burst evoked by fMLP/CB.

PMA induced a sustained rate of superoxide dismutase-inhibitable reduction of cytochrome c by peritoneal exudate cells from both WT and KO mice (Table III). The differences in responses did not reach significance (p = 0.12), suggesting that cGKI does not influence the NADPH oxidase signaling pathway downstream from PKC with extravasated neutrophils.

The regulation of neutrophil functions by cGMP/cGKI has not been adequately defined using pharmacological inhibitors, because these compounds elicit effects unrelated to cGKI (33, 34). To gain further insight into the signaling pathway of cGKI, we used neutrophils from genetically modified mice that lacked the cGKI gene. In agreement with our studies of human neutrophils, murine neutrophils express cGKI (9, 17, 25). We further demonstrated that murine neutrophils express cGKIα, the isoform with the highest affinity for cGMP (31). The murine neutrophils used in this investigation were obtained from 28- to 35-day-old mice. The cGKI-KO mice die within 42 days of age, although the exact cause of death is undetermined. The mice are notably smaller than their littermates from birth until death, and have hypertension, increased platelet adhesion and aggregation, splenomegaly, intestinal dysmotility, and stenosis of the pylorus and ileo-caeco-colic valves (37, 46). There were no differences in the relative numbers of myeloid cells in the bone marrow between WT and cGKI−/− mice. Examination of the peripheral blood showed that the cGKI−/− mice have a mild anemia and mild reticulocytosis. In addition, there was an elevated WBC and a granulocytosis with a significant elevation of segmented and band neutrophils, suggesting inflammation. However, necropsies of the cGKI mice revealed no evidence of inflammation throughout their life span.

Similar to other cell types (28, 29, 37, 42), cGMP decreased agonist-induced Ca2+ transients in WT, but not cGKI−/− bone marrow-derived neutrophils. This observation is noteworthy, because Ca2+ is regarded as an important second messenger in response to agonists in neutrophils (41). Thus, when cytosolic levels of cGMP are increasing, for example, during an inflammatory episode (8), cGKIα may be activated by NO/cGMP signaling to phosphorylate a calcium regulatory protein that functions to lower [Ca2+]i (47) and the binding subunit of myosin phosphatase (48) that results in an increased calcium sensitivity of the contractile machinery (49). In addition, modification and inhibition of the small GTPase Rho has been reported (50). The phosphorylation of these various targets may explain why moderate chelation of cytosolic calcium has been reported to have a minimal or no effect on membrane ruffling, actin polymerization, and other cytoskeletal functions (51). We conclude that deletion of cGKI can affect neutrophil motility and function by several mechanisms.

Chemotaxis to IL-8 and C5a was greater in cGKI-deficient neutrophils than WT controls, suggesting that cGKIα down-regulates neutrophil chemotaxis. Both WT and cGKI-deficient PE-N were more responsive to C5a than the bone marrow-derived neutrophils, consistent with the possibility that C5a receptors were up-regulated by degranulation of secretory vesicles, gelatinase granules, and/or specific granules in both control and mutant neutrophils during transmigration to the peritoneum.

The in vitro results were confirmed in the AA ear model. Again, more neutrophils migrated into the cGKI-deficient ear than in the WT ear, as determined by MPO activity (4-fold increase) and histology. The MPO levels of control ear punch homogenates were similar in WT and cGKI−/− mice, indicating that cGKI-deficient neutrophils migrate preferentially to sites of inflammation. Of particular relevance was the observation that extravasated cGKI-deficient neutrophils contained twice the quantity of MPO than WT peritoneal exudate controls (see secretion section below). However, even when assumptions are made that extravasated cGKI-deficient neutrophils have twice the quantity of MPO than controls, the 4-fold increase in MPO activity and the predominance of neutrophils in the ear punches over WT controls suggests that chemotaxis is greater in the cGKI−/− animals.

Interestingly, the ear weight was significantly increased in the cGKI−/− animals, suggesting enhanced vascular permeability in the cGKI−/− animals. Release of eicosanoids has been shown to elevate cGMP levels in neutrophils and other cells (52, 53, 54). Activation of cGKI reduces vascular tone by a number of different mechanisms (28, 29, 48, 49). Therefore, it is conceivable that the increased tone of vascular smooth muscle in cGKI−/− cells facilitated the increased permeability. Further studies are required to investigate cGKI signal transduction in regulating vascular permeability.

Chemotactic processes are, at least partly, dependent upon intracellular Ca2+. Neutrophils undergo repeated Ca2+ influx events as chemotaxis proceeds (55, 56, 66). It is interesting that chemotaxis is enhanced, and there is a cGMP-dependent suppression in [Ca2+]i transients in C5a-stimulated BM-Ns. A correlation between the [Ca2+]i and the speed of neutrophil migration has been reported (55). Therefore, it is possible that cGKI may regulate neutrophil motility by lowering agonist-induced [Ca2+]i transients. In addition, cGKI may regulate neutrophil motility by affecting myosin phosphatase activity (48), thereby affecting filament formation (e.g., pseudopodia). Furthermore, VASP, a member of the actin-binding protein family Drosophila Enabled/VASP, also plays a key role in cell motility (57). In neutrophils, VASP is phosphorylated by cGKI and colocalizes with F-actin in filopodia and focal adhesions during spreading, a time when cGMP levels are elevated (26). Fibroblasts deficient in mammalian homologue of Drosophila Enabled/VASP move faster than controls (58, 59), suggesting that a possible mechanism for the increase in motility of cGKI−/− neutrophils is the reduced phosphorylation of VASP by cGKI. It is possible that the cGKI-deficient neutrophils may move faster because agonist-evoked [Ca2+]i transients remain high, thus leading to a faster turnover of focal adhesions, and that VASP and myosin phosphatase are targets regulated by cGKI (for detailed discussion, see Ref.31). Further studies are required to determine the mechanism for the abnormal chemotaxis response of cGKI-deficient neutrophils.

Extravasated neutrophils were used to investigate granule secretion and the respiratory burst. Comparative studies with BM-Ns were not feasible, because large numbers of neutrophils were required for experimentation, and the survival of the cGKI-deficient animals is poor. However, in light of the fact that neutrophils perform their primary function at sites of inflammation, the PE-N offered the opportunity to investigate the responsiveness of a population of neutrophils that were recruited in vivo to an inflammatory site.

We were surprised to find that extravasated neutrophils from cGKI-deficient mice contain a significant 2-fold increase in the concentration of total extractable MPO, but not lysozyme. Therefore, this increase in MPO suggests that either there is a packaging defect for MPO during myelopoiesis, or that cGKI−/− PE-N retained a population of MPO-containing granules that was released by WT cells en route from the peripheral blood to the peritoneum. MPO was undetectable after stimulation with PMA, a specific granule agonist, for both cGKI−/− and WT neutrophils. If MPO packaging were defective, we might expect that MPO would be released with lysozyme after PMA stimulation. It is also possible that MPO is synthesized in greater quantity during granulogenesis, but is packaged appropriately in the primary granules. Unfortunately, we were unable to determine the MPO content of bone marrow-derived neutrophils. It is also possible that there is a subset of MPO-containing granules that is under the regulation of cGKI and is released in transit from the peripheral blood to the peritoneum. This hypothesis would predict that the WT, but not the cGKI−/− neutrophils have released a subset of MPO-containing granules before arriving in the peritoneum.

A disorder in the release of MPO was also observed when the extravasated cGKI-deficient neutrophils were stimulated with fMLP/CB. The cGKI−/− PE-N released more MPO than WT cells, and there was a 2-fold increase in the percentage of MPO released after fMLP/CB stimulation by cGKI-deficient PE-N vs WT controls. Interestingly, both WT and cGKI-deficient PE-N retained the same amount of MPO after stimulation with fMLP/CB (0.522 U for WT vs 0.557 U for cGKI−/−), suggesting that both WT and mutant neutrophils may be retaining the same population of MPO-containing granules after stimulation with fMLP/CB. The defensin-rich population of azurophil granules may be retained because their primary function is to kill bacteria during phagocytosis (60). Whether or not this subset of granules is the defensin-rich population remains to be investigated. The 69% release of MPO by mutant PE-N vs 36% MPO by WT cells may represent the MPO from a granule subset(s) that was released by WT cells in transit plus the remainder MPO from other granule subsets. Thus, as neutrophils transit from the peripheral blood to the peritoneum, cGKI regulates the secretion of subsets of MPO-containing granules that are activated in transit (e.g., via adhesion receptors), and activated in the peritoneum (e.g., via inflammatory mediators such as fMLP).

By comparing the responsiveness of the same population of neutrophils that were stimulated with fMLP/CB (PKC independent and dependent) and PMA (PKC dependent), we were able to determine whether there was interdependence between cGKI and PKC signaling. Similar to WT PE-N, MPO was not released by cGKI−/− cells after stimulation with PMA. Thus, secretion of MPO-containing granules is PKC independent, and cGKI is not involved in regulating MPO release downstream from PKC. However, after stimulation with fMLP/CB, both the quantity and percentage of MPO released by cGKI−/− PE-N was significantly greater when compared with WT controls. Based upon the fact that MPO is not released by WT and cGKI−/− PE-N by PMA, we suggest that the regulation of these MPO granule subset(s) by cGKI in response to fMLP/CB is PKC independent.

Unlike MPO, there were no significant differences in lysozyme content in extravasated cGKI-deficient neutrophils. Although the differences in the amounts of lysozyme secreted after fMLP or PMA were not statistically significant, the percentages of enzyme released by WT and mutant PE-N were significantly different. In WT PE-N, ∼88% lysozyme was released after fMLP/CB or PMA, whereas only 62% lysozyme was released by cGKI−/− neutrophils with both agonists. These data suggest that ∼26% of the lysozyme-containing granule population is regulated by cGKIα, and that these are specific granules (lack MPO) and are PKC dependent. We propose that the site of cGKI regulation for this subset of specific granules is downstream from PKC activation, because 26% of the lysozyme was retained in cGKI-deficient neutrophils that were stimulated with both fMLP/CB and PMA.

Both fMLP and PMA stimulate significant elevations of cGMP levels in neutrophils (10, 25, 52). We expect that cGMP levels would also be elevated in mouse neutrophils, because we and others have shown that the regulation of cGMP levels is not altered in cGKI-deficient mice (61, 62). Although the substrates for cGKI are not clear, cGKI appears to play an important role in regulating signal transduction pathways in neutrophils, particularly calcium mobilization and p38 MAPK. Calcium mobilization in and of itself induces granule exocytosis in neutrophils, and a hierarchical mobilization of neutrophil granule subsets can be achieved in vitro by gradual elevations in the intracellular Ca2+ levels (63). It is well known that the MPO-containing granules require more Ca2+ for release than the gelatinase and specific granules (60). Therefore, it is possible that [Ca2+]i remain higher after fMLP stimulation in cGKI-deficient neutrophils vs WT cells, and that these elevated calcium levels promote the enhanced secretion of MPO-containing granules. In support of this hypothesis, fMLP-induced Ca2+ transients were inhibited in rat neutrophils by YC-1, an activator of soluble guanylyl cyclase (64). Because fMLP, like C5a, is activated by G protein coupling and evokes calcium transients, it is possible that cGKI down-regulates [Ca2+]i in fMLP-stimulated neutrophils as well.

Another of the many sites of cGKI regulation may include the MAPK pathway (65), because p38 MAPK is a known substrate for cGKI in LPS-primed neutrophils that are stimulated with fMLP (27). Furthermore, there is evidence that secretion of azurophil and specific granules after fMLP stimulation is mediated by p38 MAPK that is activated via Src family tyrosine kinases (67).

Although the differences in the Vmaxinit responses of the WT vs the cGKI−/− PE-N responses to fMLP/CB failed to reach significance, measurements of the AUC revealed statistically significant differences between genotypes. The cGKI−/− PE-N appeared to take longer to shut down production of superoxide than did WT cells, suggesting that cGKI may serve to dampen the second phase of the respiratory burst evoked by fMLP/CB. When the same cell populations were stimulated with PMA, the differences between the WT and cGKI−/− PE-N in O2 production did not reach significance. These data suggest that cGKI does not affect the NADPH oxidase signaling pathway downstream from PKC with extravasated neutrophils.

In contrast to the activation of PKC by PMA, the activation of formyl peptide receptors generate multiple second messengers resulting in an increase in 1,2-diacylglycerol and inositol 1,4,5-triphosphate as well as an increase in [Ca2+]i. In contrast to the PMA-induced respiratory burst, these multiple pathways evoked by fMLP stimulation result in a complex kinetics of respiratory burst, suggesting that cGKI influences the production of superoxide over time, possibly by modulation of the Ca2+-regulating protein (31, 47).

In summary, the neutrophil is a complex cell with multiple functions, many of which are dependent upon the selective release of cytoplasmic granules as the neutrophil progresses from a quiescent cell in the bone marrow and peripheral blood, to a migratory cell as it leaves the circulation, and finally to an inflammatory cell in the extravascular tissue. Our findings suggest that cGKIα down-regulates chemotaxis and agonist-induced increases in [Ca2+]i, and that it may dampen the respiratory burst over time in extravasated neutrophils. cGKIα may regulate the release of a subset(s) of azurophil granules during transmigration and at sites of inflammation. Furthermore, extravasated neutrophils contain a subset of specific granules that appear dependent on cGKIα for release. Further studies are required to resolve the specific sites of cGKIα regulation in neutrophils.

We thank Dr. Margrith Verghese for her technical advice and helpful review of the manuscript, and Pam McElveen, Sanjeda Sultana, and Astrid Lauxen for their skillful technical assistance.

The authors have no financial conflict of interest.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1

This work was supported by grants from Bundesministerium für Bildung und Forschung, Deutsche Forschungsgemeinschaft, and North Atlantic Treaty Organization.

3

Abbreviations used in this paper: AA, arachidonic acid; [Ca2+]i, cytosolic calcium concentration; cGKI, type I cGMP-dependent protein kinase; VASP, vasodilator-stimulated phosphoprotein; WT, wild type; KO, knockout; PE-N, peritoneal exudate neutrophils; GBSS, Geys’ balanced salt solution; BMC, bone marrow cell; MPO, myeloperoxidase; CB, cytochalasin B; AUC, area under curve; 8-Br-cGMP, 8-bromo-cGMP; BM-N, bone marrow neutrophil; PKC, protein kinase C.

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