The binding of Ab (IgG)-opsonized particles by FcγRs on macrophages results in phagocytosis of the particles and generation of a respiratory burst. Both IgG-stimulated phagocytosis and respiratory burst involve activation of protein kinase C (PKC). However, the specific PKC isoforms required for these responses have yet to be identified. We have studied the involvement of PKC isoforms in IgG-mediated phagocytosis and respiratory burst in the mouse macrophage-like cell line, RAW 264.7. Like primary monocyte/macrophages, their IgG-mediated phagocytosis was calcium independent and diacylglycerol sensitive, consistent with novel PKC activation. Respiratory burst in these cells was Ca2+ dependent and inhibited by staurosporine and calphostin C as well as by the classic PKC-selective inhibitors Gö 6976 and CGP 41251, suggesting that classic PKC is required. In contrast, phagocytosis was blocked by general PKC inhibitors but not by the classic PKC-specific drugs. RAW 264.7 cells expressed PKCs α, βI, δ, ε, and ζ. Subcellular fractionation demonstrated that PKCs α, δ, and ε translocate to membranes during phagocytosis. In Ca2+-depleted cells, only novel PKCs δ and ε increased in membranes, and the time course of their translocation was consistent with phagosome formation. Confocal microscopy of cells transfected with green fluorescent protein-conjugated PKC α or ε confirmed that these isoforms translocated to the forming phagosome in Ca-replete cells, but only PKC ε colocalized with phagosomes in Ca2+-depleted cells. Taken together, these results suggest that the classic PKC α mediates IgG-stimulated respiratory burst in macrophages, whereas the novel PKCs δ and/or ε are necessary for phagocytosis.

Immunoglobulin G-mediated phagocytosis is part of the normal host defense system and may also be a component of inflammatory diseases such as rheumatoid arthritis and atherosclerosis. It is initiated by the binding of IgG-opsonized particles to FcγRs on the surface of monocyte/macrophages and neutrophils. Phagocytosis requires the cells to coordinate changes in cytoskeleton and membrane structure during pseudopod extension and particle internalization. In addition to phagocytosis, FcγR ligation activates other signaling pathways, including those regulating intracellular Ca2+ flux, mitogen-activated protein kinase cascades, and the respiratory burst (1, 2). Current research has focused on identifying signaling pathways activated by FcγR. Intracellular events resulting from FcγR clustering include tyrosine phosphorylation, actin rearrangement, arachidonic acid release, and activation of signaling enzymes phosphoinositol 3-kinase, protein kinase C (PKC),3 phospholipase C, phospholipase D, and phospholipase A2 (1, 2, 3, 4). However, the specific functions of these components in FcγR-mediated signaling remain unknown.

Our previous work has confirmed a role for PKC activation during IgG-mediated phagocytosis in human monocytes and the human monocytic cell line, Mono Mac 6 (5, 6). PKC is a family of related enzymes, which is divided into three groups on the basis of structure and cofactor requirements (7). The classic PKC (cPKC) isoforms α, βI, βII, and γ require Ca2+, diacylglycerol (DAG), and phosphatidylserine (PS) for optimal activity. The novel PKC (nPKC) isoforms δ, ε, η, and θ lack the Ca2+ requirement but are activated by DAG and PS. The atypical PKC isoforms ζ and ι/λ bind PS, but are insensitive to Ca2+ and DAG. Despite the differing cofactor requirements, there is little difference between the in vitro substrate specificities of the isoforms. Because cells usually express several PKC isoforms, in vivo function is thought to be regulated by intracellular location and binding to specific targeting proteins (7).

Although PKC activation is necessary for phagocytosis in monocyte/macrophages, it is not known which PKC isoform(s) function in FcγR-mediated signaling (5, 6, 8). Both cPKC (α and β) and nPKC (ε) isoforms have been reported to translocate to membranes during FcγR cross-linking or phagocytosis (9, 10), but there is no direct evidence that these isoforms are required for FcγR-mediated signaling. Phagocytosis proceeds normally in the absence of a Ca2+ signal (11, 12) and can be increased by the treatment of cells with the PKC activators PMA and DAG (5, 6). These characteristics are consistent with the cofactor requirements for nPKC isoforms, i.e., Ca2+ independent and DAG sensitive. Therefore, we tested the hypothesis that one or more of the nPKC isoforms is required for phagocytosis.

IgG-mediated phagocytosis is accompanied by the generation of a respiratory burst; however, the role of specific PKC isoforms in respiratory burst is not certain. Respiratory burst can be activated by PMA in both neutrophils and monocyte/macrophages (13, 14, 15). In neutrophils, IgG-stimulated respiratory burst is decreased by pharmacological inhibition of PKC (16) or by selective antisense down-regulation of cPKC β (14), indicating a requirement for PKC. In comparison with neutrophils, studies in monocyte/macrophages have not been as consistent. In human monocytes, respiratory burst stimulated by opsonized zymosan was reduced to background levels by selective down-regulation of cPKC α (17). However, in guinea pig macrophages, pharmacological inhibition of PKC produced only a modest decrease in FcγR-mediated respiratory burst (15). We addressed the question of whether the mouse macrophage cell line RAW 264.7 was similar to human monocytes in requiring cPKC α for FcγR-mediated respiratory burst.

We have previously demonstrated that IgG-mediated phagocytosis requires PKC and that PKC activation is upstream of mitogen-activated protein kinase and Ca2+-independent phospholipase (2, 5). Phagocytes also require PKC to produce the respiratory burst that accompanies IgG-mediated phagocytosis (1). To determine the functions of PKC in these two signaling pathways, it is necessary to identify the relevant PKC isoform(s). Activation of cPKC and nPKC is associated with their translocation to specific sites (6). Therefore, we examined the translocation of cPKC and nPKC isoforms during phagocytosis.

EDTA, EGTA, BSA, thimerosal, ammonium persulfate, Triton X-100, sucrose, DTT, HRP, and NaOH were obtained from Sigma (St. Louis, MO). Homovanillic acid and DMSO were obtained from Aldrich (Milwaukee, WI). Tris base, SDS, CaCl2, and MgCl were purchased from Mallinckrodt Baker (Paris, KY). PBS was purchased from BioWhittaker (Walkersville, MD). Calphostin C, staurosporine, PMA, and 4α-PMA were obtained from LC Laboratories (Woburn, MA). DAG was purchased from Avanti Polar Lipids (Alabaster, AL). GF109203X was obtained from Biomol Research (Plymouth Meeting, PA). CGP 41251 was a gift from Novartis (Basil, Switzerland). Gö 6976 was purchased from Alexis Biochemicals (San Diego, CA).

The RAW 264.7 mouse macrophage cell line was maintained in RPMI 1640 media (Life Technologies, Grand Island, NY) plus sodium pyruvate, nonessential amino acids, glutamate (BioWhittaker), and 10% newborn calf serum (HyClone, Logan, UT). For phagocytosis experiments, cells were seeded in 24-well culture plates at 6 × 105 cells per well. For PKC translocation experiments, cells were grown in 100-mm culture plates and used when confluent. Unless otherwise specified, cells were incubated in above media without the 10% calf serum overnight before each experiment.

HBSS (Life Technologies) buffer was composed of: 4 mM sodium bicarbonate, 10 mM HEPES, containing either 1.5 mM each CaCl2 and MgCl2 (HBSS2+) or 2 mM MgCl2 and 1 mM EGTA (Mg/EGTA). Lysis buffer was composed of: 25 mM Tris-HCl, pH 7.4, 0.25 M sucrose, 2.5 mM DTT, and 2.5 mM EDTA, and contained the following inhibitors: 5 mM benzamidine, 50 μg/ml leupeptin, 50 μg/ml aprotinin, 50 μg/ml trypsin inhibitor, 5 μg/ml pepstatin, 1 mM PMSF, 20 mM NaF, 1 mM Na3VO4, 1 mM para-nitrophenylphosphate, and 5 mM imidazole (Sigma). DNA assay buffer was composed of: 2 M NaCl, 2 mM EDTA, and 50 mM Na2HPO4. TBST was composed of: 50 mM Tris, pH 7.5, 150 mM NaCl, 0.05% Tween 20, and 0.01% thimerosal.

Ig-G opsonized, radiolabeled erythrocytes were referred to as EIgG. SRBC (Crane, Syracuse, NY) were radiolabeled by incubation (45 min, 37°C) with 51Cr (New England Nuclear, Boston, MA) and opsonized with rabbit anti-SRBC IgG (Cappel, Durham, NC) (18). Glass beads (Duke Scientific, Palo Alto, CA) were opsonized by sequential incubation with poly-l-lysine, BSA, and anti-BSA IgG (Sigma) as previously described (5), and referred to as BIgG. Control beads were incubated with poly-l-lysine and BSA only (BBSA). For confocal analysis, the BSA-coating step included 5 μg of Alexa 568 (Molecular Probes, Eugene, OR)-conjugated BSA. BSA was conjugated to the Alexa dye according to manufacturer’s directions.

If cells were described as Ca depleted, they were incubated (45 min, 37°C) in Mg/EGTA buffer (−Ca cells), otherwise the assay was performed in HBSS2+ (+Ca cells). When used, inhibitors were added for the last 30 min of the incubation. EIgG (6 × 106) were added to each well for the designated time at 37°C. The cells were washed twice with 0.83% NH4Cl, once in PBS, and solubilized in 2 M NaOH. Phagocytosis is defined as the cell-associated radioactivity after hypotonic lysis of noninternalized targets and expressed as a percentage of phagocytosis in control cells. Nonspecific binding was determined using nonopsonized SRBC and subtracted from the phagocytosis values.

To determine the time course of phagocytosis of BIgG, 2 × 105 cells were plated on 13-mm glass coverslips (Ernest F. Fullam, Latham, NY) in 24-well plates. The cells were incubated in buffer as described above and then chilled on ice for 20 min. The buffer was removed, and 0.5 ml of ice-cold buffer containing 1 × 106 BIgG was added. After 5 min on ice for target binding, the plates were transferred to a 37°C water bath, and, at each time point, phagocytosis was stopped by fixing the cells with 3.7% formaldehyde. The coverslips were washed three times with PBS and blocked with 10% sheep serum (30 min, 21°C). The following incubations were performed with Ab diluted in 10% sheep serum (30 min, 21°C): 1) rabbit anti-BSA, 1:250; 2) FITC-conjugated goat anti-rabbit, (Rockland, Gilbertsville, PA), 1:200; 3) permeabilize 5 min in 0.01% Triton X-100; 4) block as above; 5) rabbit anti-BSA; 6) Texas-red conjugated goat anti-rabbit, (ICN Pharmaceuticals, Aurora, OH), 1:250; and 7) Hoechst nuclear stain (Molecular Probes). Coverslips were washed three times with PBS after each incubation. After washing, the coverslips were mounted on glass slides with Prolong antifade medium (Molecular Probes) and viewed with a triple-band filter. The cell nuclei stained blue, internalized beads were red, and external beads or portions of beads were yellow/green. The number of completely internalized beads were counted in a minimum of 100 cells and expressed as the phagocytic index: (no. of red beads/number of cells counted) × 100.

H2O2 production was determined as described previously (18). Briefly, 1.2 × 106 cells were incubated for 1 h in 1 ml HBSS containing 100 nM homovanillic acid and 1 IU HRP, with 1 × 107 BIgG (see Figs. 5 and 7) or 50 ng PMA (see Fig. 6) as the stimulus. The homovanillic acid oxidation product was measured fluorometrically.

FIGURE 5.

General PKC inhibitors decrease phagocytosis and respiratory burst in RAW 264.7 cells. Cells were preincubated with staurosporine (A) or calphostin C (B) at the indicated doses (30 min, 37o), and phagocytosis or respiratory burst assayed as described in Materials and Methods. Cells treated with calphostin C were illuminated with fluorescent light for the last 15 min during the incubation to activate the drug. Control (100%) is defined as the phagocytosis or H2O2 release by cells treated with carrier (DMSO) only. Data are expressed as mean ± SEM, n = 3. ∗, p < 0.05.

FIGURE 5.

General PKC inhibitors decrease phagocytosis and respiratory burst in RAW 264.7 cells. Cells were preincubated with staurosporine (A) or calphostin C (B) at the indicated doses (30 min, 37o), and phagocytosis or respiratory burst assayed as described in Materials and Methods. Cells treated with calphostin C were illuminated with fluorescent light for the last 15 min during the incubation to activate the drug. Control (100%) is defined as the phagocytosis or H2O2 release by cells treated with carrier (DMSO) only. Data are expressed as mean ± SEM, n = 3. ∗, p < 0.05.

Close modal
FIGURE 7.

Macrophage functions of phagocytosis and respiratory burst can be separated by selective inhibition of cPKC. Cells were incubated (30 min, 37°C) with the indicated doses of CGP 41251 (A), Gö 6976 (B), or GF109203X (C), and phagocytosis or respiratory burst was assayed. Control (100%) is defined as the phagocytosis or H2O2 release by cells treated with carrier only. Data are expressed as mean ± SEM, n = 3. ∗, p < 0.05.

FIGURE 7.

Macrophage functions of phagocytosis and respiratory burst can be separated by selective inhibition of cPKC. Cells were incubated (30 min, 37°C) with the indicated doses of CGP 41251 (A), Gö 6976 (B), or GF109203X (C), and phagocytosis or respiratory burst was assayed. Control (100%) is defined as the phagocytosis or H2O2 release by cells treated with carrier only. Data are expressed as mean ± SEM, n = 3. ∗, p < 0.05.

Close modal
FIGURE 6.

Respiratory burst is abolished by Ca depletion of RAW 264.7 cells. H2O2 production was measured in control cells (Ca), cells Ca depleted as described (EGTA), or cells Ca depleted then resuspended in Ca-containing buffer during the stimulus of the respiratory burst (EGTA/Ca). Data are expressed as mean ± SE, n = 3.

FIGURE 6.

Respiratory burst is abolished by Ca depletion of RAW 264.7 cells. H2O2 production was measured in control cells (Ca), cells Ca depleted as described (EGTA), or cells Ca depleted then resuspended in Ca-containing buffer during the stimulus of the respiratory burst (EGTA/Ca). Data are expressed as mean ± SE, n = 3.

Close modal

For PMA stimulation, confluent cultures of RAW 264.7 cells in 10-cm dishes were treated with 100 nM PMA or an equivalent volume of DMSO (carrier) (10 min, 37°C), the media was aspirated, and the cells were scraped and sonicated in 0.4 ml of lysis buffer. Lysates were centrifuged 45 min at 100,000 × g. The supernatant (designated cytosol) was removed, and the pellet was solubilized in lysis buffer plus 0.2% Triton X-100. The samples were centrifuged again as above. The supernatant (designated membrane) was removed from the pellet (designated insoluble fraction). For phagocytosis experiments, the protocol for synchronized phagocytosis of BIgG was followed using cells in 100-mm culture dishes and 1 × 108 targets. Phagocytosis was stopped at the various time points by scraping and sonicating the cells in lysis buffer; cell lysates were processed as described above. As no PKCs were detected in the insoluble cell fractions following phagocytosis (data not shown), PKC levels were quantified only in the membrane and cytosolic fractions.

DNA was measured by the method of Labarca and Paigen (19). Briefly, aliquots of whole cell lysate were diluted with assay buffer and 2% (final concentration) bisbenzamide, and the fluorescence was compared with a standard curve prepared with calf thymus DNA.

For PMA experiments, protein in cytosolic and membrane fractions was quantified by the Bradford protein assay. Different proportions of the cell fractions were loaded to obtain a measurable signal; however, the amount in each fraction was matched between treatments to allow comparison. This corresponded to ∼6% of the cytosol, 25% of the membrane, and 25% of the insoluble fractions. For phagocytosis experiments, equal volumes of the membrane fractions were loaded. Proteins were transferred to nitrocellulose membranes, and standards were located with Ponceau S stain. The membranes were blocked with 3% BSA in TBST and probed with the appropriate primary and secondary Ab in 1% BSA in TBST. For the primary Ab, mAb against PKCs α, δ, ζ, θ, and η, (Transduction Laboratories, San Diego, CA) and polyclonal Ab against PKCs βI, βII, and ε (Santa Cruz Biotechnology, Santa Cruz, CA) were used. For the secondary Ab, goat anti-rabbit HRP (Santa Cruz Biotechnology) and rabbit anti-mouse HRP (The Jackson Laboratory, Bar Harbor, ME) were used. Bands were detected with Ultra Supersignal ECL reagent (Pierce, Rockford, IL) and were quantified by densitometry. In phagocytosis experiments, the densities of the PKC bands were normalized for cell DNA before comparison by ANOVA.

The construction and characterization of the green fluorescent protein (GFP) PKC α and ε plasmids have been previously described (20). These constructs are enzymatically active and translocate to membranes in response to a variety of stimuli. DNA (14 μg/6 × 106 cells) was transfected into RAW 264.7 cells using Superfect (Qiagen, Valencia, CA) as per manufacturer’s instructions. Cells were exposed to DNA for 3 h, then washed and incubated for an additional 12–18 h to allow maximal expression of fluorescence.

Transfected cells were replated onto 13-mm coverslips (5 × 105 cells/coverslip) for 3 h. Media were removed and the cells were washed and incubated in either HBSS2+ or Mg/EGTA (45 min, 37°C). Cells were cooled on ice (30 min), and ice-cold Alexa 568-conjugated BIgG were added at a 5:1 BIgG/cell ratio. Following target binding (15 min on ice), cells were placed in a 37°C water bath for 0–15 min. At varying times, the cells were fixed (5 min, 3.7% formaldehyde), mounted in Prolong Antifade (Molecular Probes), and analyzed by confocal microscopy. Z series images of GFP-expressing cells were analyzed on a Noran-OZ (Noran Instruments, Middleton, WI) confocal laser scanning microscope, interfaced with a Nikon Diaphot 200 inverted microscope (Nikon, Melville, NY) equipped with a PlanApo ×60, 1.4 NA oil-immersion objective lens. GFP and Alexa 568 were independently imaged using excitation/emission wavelengths of 488/500–550 nm band pass and 568/590 nm long pass, respectively. Images were analyzed using Noran InterVision software.

Unless otherwise stated, all measurements were made in triplicate on at least three separate cell preparations. Data are expressed as the mean ± SEM. Comparisons were made by ANOVA. Results with p ≤ 0.05 were considered significant.

We identified the PKC isoforms expressed in RAW 264.7 cells by Western blot analysis with isoform-specific Abs (Fig. 1). cPKC α, cPKC βI, nPKC δ, nPKC ε, and atypical PKC ζ were detected in RAW 264.7 cell lysates. cPKC γ, nPKC θ, and nPKC η could be detected only in positive controls but not in RAW 264.7 cell lysate (data not shown). PKC βII could not be reproducibly detected in cell fractions.

FIGURE 1.

Detection of PKC isoforms in RAW 264.7 cells. Cell lysate was prepared and analyzed by Western blotting as described. PKCs α, βI, δ, ε, and ζ were detected.

FIGURE 1.

Detection of PKC isoforms in RAW 264.7 cells. Cell lysate was prepared and analyzed by Western blotting as described. PKCs α, βI, δ, ε, and ζ were detected.

Close modal

Earlier studies demonstrated that IgG-mediated phagocytosis is Ca2+ independent in monocytes and macrophages (11, 12). Ca2+ depletion did not affect the extent or rate of IgG-mediated phagocytosis by RAW 264.7 cells (Fig. 2), indicating that this is an appropriate model for studying Ca-independent phagocytic signaling. The cells were Ca2+ depleted by incubation in Mg/EGTA buffer (45 min, 37°C). We have previously shown that this treatment virtually eliminates free intracellular calcium concentration ([Ca2+]i) in monocytes as determined by fura-2 fluorescence in cells stimulated with IgG-opsonized particles (12). Similar results were obtained with RAW 264.7 cells stimulated with either platelet-activating factor or immune complexes (data not shown).

FIGURE 2.

IgG-mediated phagocytosis is Ca independent in RAW 264.7 macrophages. Phagocytosis assays were conducted as described in Materials and Methods, with time points of 0, 5, 15, 30, 45, and 60 min. Control (100%) is defined as the cell-associated radioactivity at 60 min in Ca-containing cells. Data are expressed as mean ± SEM, n = 3.

FIGURE 2.

IgG-mediated phagocytosis is Ca independent in RAW 264.7 macrophages. Phagocytosis assays were conducted as described in Materials and Methods, with time points of 0, 5, 15, 30, 45, and 60 min. Control (100%) is defined as the cell-associated radioactivity at 60 min in Ca-containing cells. Data are expressed as mean ± SEM, n = 3.

Close modal

Because Ca2+ depletion did not alter phagocytosis, we examined its affect on PKC translocation. We tested the hypothesis that Ca2+ depletion would inhibit membrane localization of PKCs α and βI in response to PMA, a potent stimulus of PKC translocation. The baseline distribution of PKC isoforms was the same in +Ca and −Ca cells, indicating that Ca2+ depletion alone did not alter PKC location (data not shown). PKCs α, βI, δ, and ε are primarily cytosolic in unstimulated cells (Fig. 3). Although the intensity of the bands in the cytosol and membrane fractions for βI appear similar, the differences in loading must be taken into account. As ∼25% of the membrane fraction vs 6% of the cytosol was loaded, the majority of PKC βI, like the other isoforms, is cytosolic.

FIGURE 3.

Depletion of intracellular calcium prevents the PMA-induced translocation of PKC α but not PKC δ or ε. Western blots show the distribution of PKCs between cytosol (C), membrane (M), and insoluble (I) fractions in control cells (+Ca, DMSO), Ca-containing cells treated (10 min at 37o) with 100 nM PMA (+Ca, PMA), and cells that were Ca depleted (as described in Materials and Methods) and treated (10 min at 37o) with 100 nM PMA (−Ca, PMA). The proportion of recovered protein loaded for each fraction was different (C: ∼6%, M and I: ∼25%), but the amount in each fraction was the same among treatments. Data are representative of three experiments.

FIGURE 3.

Depletion of intracellular calcium prevents the PMA-induced translocation of PKC α but not PKC δ or ε. Western blots show the distribution of PKCs between cytosol (C), membrane (M), and insoluble (I) fractions in control cells (+Ca, DMSO), Ca-containing cells treated (10 min at 37o) with 100 nM PMA (+Ca, PMA), and cells that were Ca depleted (as described in Materials and Methods) and treated (10 min at 37o) with 100 nM PMA (−Ca, PMA). The proportion of recovered protein loaded for each fraction was different (C: ∼6%, M and I: ∼25%), but the amount in each fraction was the same among treatments. Data are representative of three experiments.

Close modal

PMA treatment of Ca-containing cells caused a >6-fold decrease in cytosolic PKC α (Fig. 3, lane 1 vs 4) and a substantial increase in the particulate (membrane + insoluble) fractions (>12-fold; lanes 2 and 3 vs lanes 5 and 6). In contrast, Ca2+ depletion prevented the translocation of PKC α to membrane and insoluble fractions in response to PMA, resulting in a PKC α distribution similar to that of resting cells (Fig. 3, lanes 1–3 vs 7–9). PMA stimulated PKC βI translocation to the insoluble fraction in the presence, but not the absence, of Ca2+. In the presence of Ca2+, PKC βI in the cytosol and membrane fractions decreased by 9-fold (Fig. 3, lanes 1 and 2 vs 4 and 5) and the insoluble fraction increased >40-fold (lane 3 vs 6). In the absence of Ca2+, the distribution of PKC βI was virtually identical in PMA-treated and control cells (Fig. 3, lanes 1–3 vs 7–9). This confirms that the cells were indeed Ca depleted, and that Ca depletion inhibited the PMA-induced translocation of cPKCs α and βI.

In contrast, Ca2+ depletion did not block the translocation of nPKC isoforms to the particulate (membrane + insoluble) fractions. PMA treatment decreased cytosolic PKC δ (∼2-fold) and increased its levels in the particulate fractions (∼3-fold), as shown in Fig. 3. This pattern occurred in both Ca2+-containing and Ca2+-depleted cells (Fig. 3, lanes 4–6 are similar to lanes 7–9). In cells treated with PMA, PKC δ was decreased in the cytosol (>3-fold) and increased in the particulate fractions (3- to 4-fold). Similar translocation occurred in Ca2+-containing and Ca2+-depleted cells (Fig. 3, lanes 4–6 are similar to lanes 7–9). The translocation of nPKCs δ and ε in Ca2+-depleted cells was predicted, as these isoforms lack the functional Ca2+ binding domain found in cPKC isoforms (7).

Phagocytosis in human monocytes is increased by PMA or DAG, consistent with a role for PKC activation in phagocytosis (5, 8). Likewise, phagocytosis in RAW 264.7 cells was also PMA/DAG sensitive, increasing >60% when the cells were treated with either of these PKC activators (Fig. 4). Conversely, treatment with the PKC inhibitors calphostin C or staurosporine decreased phagocytosis in a dose-dependent fashion (Fig. 5, A and B). Thus, similar to our results in primary monocytes and MonoMac 6 cells (5, 6), phagocytosis in RAW 264.7 cells requires a PKC that is both Ca2+ independent and PMA/DAG sensitive, matching the characteristics of the nPKC isoforms.

FIGURE 4.

PMA and DAG increase phagocytosis in RAW 264.7 cells. Cells were pretreated with the indicated concentrations of PMA (7 min) or DAG (15 min), then incubated with EIgG targets (60 min, 37°C) and phagocytosis was assayed as described in Materials and Methods, except that for the DAG experiments the cells were not serum deprived. Control (100%) is defined as the phagocytosis in cells treated with carrier (DMSO) only. Data are expressed as mean ± SEM, n = 3. ∗, p < 0.05.

FIGURE 4.

PMA and DAG increase phagocytosis in RAW 264.7 cells. Cells were pretreated with the indicated concentrations of PMA (7 min) or DAG (15 min), then incubated with EIgG targets (60 min, 37°C) and phagocytosis was assayed as described in Materials and Methods, except that for the DAG experiments the cells were not serum deprived. Control (100%) is defined as the phagocytosis in cells treated with carrier (DMSO) only. Data are expressed as mean ± SEM, n = 3. ∗, p < 0.05.

Close modal

Unlike particle uptake via complement receptors or endocytosis, IgG-mediated phagocytosis is accompanied by a respiratory burst (1, 21), which requires translocation and assembly of NADPH oxidase components at the membrane, and results in production of O2⨪ and H2O2 (18, 22). NADPH oxidase activation has been shown to require an increase in intracellular Ca2+ and PKC activity (22, 23). As predicted, calphostin C and staurosporine decreased H2O2 production by RAW 264.7 cells >75% at doses similar to those that affect phagocytosis (Fig. 5, A and B). Additionally, depletion of intracellular Ca2+ abolished H2O2 production (Fig. 6). As PKCs α and βI did not translocate to the membrane in Ca2+-depleted cells (Fig. 3), these results are consistent with the hypothesis that cPKC activation is required for IgG-mediated respiratory burst in RAW 264.7 cells.

This hypothesis predicts that the cPKC selective inhibitors CGP 41251 and Gö 6976 will block the IgG-stimulated burst, but not affect phagocytosis. Fig. 7 (A and B) demonstrates that CGP 41251 and Gö 6976 caused a dose-dependent decrease in H2O2 production, but not phagocytosis. Interestingly, the nonselective PKC inhibitor GF109203X also blocked respiratory burst but not phagocytosis (Fig. 7 C). These results are further evidence for a role for cPKC α and/or βI in the respiratory burst in RAW 264.7 cells. Phagocytosis was not affected, implying that cPKCs are not involved in FcγR-mediated ingestion and that these drugs are not otherwise interfering with FcγR signaling.

cPKC and nPKC isoforms associate with membranes by binding DAG and PS (7). We reasoned that those PKC isoforms involved in FcγR-initiated signaling would translocate to the membrane fraction during IgG-mediated phagocytosis, so we measured membrane-bound cPKC α, cPKC βI, nPKC δ, and nPKC ε during synchronized phagocytosis of IgG-opsonized glass beads. Time course studies showed that phagocytosis took at least 7.5 min (Fig. 8 A). Few completely internalized targets were observed before 7.5 min. Internalized targets were present in most cells at 7.5 min, and phagosomes continued to form and close by the 10- and 15-min time points. Therefore, fractions were prepared from cells at time points between 0 and 15 min to study translocation of PKC isoforms during phagosome formation (0–5 min) and formation/closure (7.5–15 min).

FIGURE 8.

A, Time-dependent internalization of IgG-opsonized glass beads. Cells were grown on glass coverslips and phagocytosis was determined as described in Materials and Methods. Data shown are average ± range, n = 2. B, Translocation of PKC isoforms during BIgG phagocytosis. Western blots (described in Materials and Methods) show PKC isoforms α, βI, δ, and ε in the membrane fractions of RAW 264.7 cells undergoing phagocytosis of IgG-opsonized glass beads for the indicated times. Cells were Ca replete (+Ca) or Ca depleted (−Ca) as described in Materials and Methods. Mouse brain (+) was included as a positive control. Blots are representative of a minimum of three experiments. C, Quantitation of membrane-associated nPKC isoforms during phagocytosis in Ca-depleted cells. Densities were normalized for cell DNA before comparison by ANOVA. ∗, Significantly greater than 0 time point, p < 0.05. PKC δ, n = 3; PKC ε, n = 4.

FIGURE 8.

A, Time-dependent internalization of IgG-opsonized glass beads. Cells were grown on glass coverslips and phagocytosis was determined as described in Materials and Methods. Data shown are average ± range, n = 2. B, Translocation of PKC isoforms during BIgG phagocytosis. Western blots (described in Materials and Methods) show PKC isoforms α, βI, δ, and ε in the membrane fractions of RAW 264.7 cells undergoing phagocytosis of IgG-opsonized glass beads for the indicated times. Cells were Ca replete (+Ca) or Ca depleted (−Ca) as described in Materials and Methods. Mouse brain (+) was included as a positive control. Blots are representative of a minimum of three experiments. C, Quantitation of membrane-associated nPKC isoforms during phagocytosis in Ca-depleted cells. Densities were normalized for cell DNA before comparison by ANOVA. ∗, Significantly greater than 0 time point, p < 0.05. PKC δ, n = 3; PKC ε, n = 4.

Close modal

Fig. 8 B shows Western blots of PKC isoforms in the membrane fractions from +Ca and −Ca cells. In +Ca cells, PKCs α and βI were present in the membrane at all time points. In contrast, PKCs α and βI were not detectable in the membrane fractions in cells depleted of intracellular Ca2+, although they could be readily detected in mouse brain lysate included as a positive control on the blots. As phagocytosis proceeded normally in Ca2+-depleted cells, these results indicate that membrane localization of PKCs α and βI is not required for phagocytosis. In the +Ca experiments, PKC βI did not increase in the membranes during phagocytosis. PKC α always increased in the +Ca membrane during phagocytosis, but the magnitude of the increase at the specific time points was variable. To test for an effect of phagocytosis on PKC α distribution, ANOVA indicated that levels of membrane-bound PKC α increased during phagocytosis (p < 0.05).

In contrast to PKCs α and βI, membrane-bound PKCs δ and ε were present during phagocytosis in both +Ca and −Ca cells (Fig. 8,B). ANOVA indicated that phagocytosis increased PKCs δ and ε in the membrane in both +Ca and −Ca cells (p < 0.05). To examine the pattern of nPKC translocation during phagosome formation, the levels of membrane PKCs δ and ε in Ca2+-depleted cells were quantified (Fig. 8 C). PKC δ increased gradually and was maximally elevated after 7.5 min, corresponding to the appearance of closed phagosomes. PKC ε was significantly increased in the membrane fraction by 5 min, suggesting that this isoform translocates during the formation of phagosomes.

Although the amount of membrane-bound PKC increased during phagocytosis, the cytosolic levels did not change (data not shown). Unlike PKC activation during PMA stimulation, the amount of PKC associated with the membranes during phagocytosis was small compared with that in the cytosol. This is consistent with phagocytosis being a localized membrane event, with a relatively small proportion of the PKC mass targeted to membrane sites involved in phagocytosis.

The detection of PKCs α and ε in membrane fractions during phagocytosis is consistent with their localization to phagosomes. To identify the membranes to which these isoforms move, we transfected GPF-conjugated PKCs α and ε into RAW 264.7 cells and examined their location during phagocytosis. In the presence of Ca2+, GFP PKC α concentrated in membrane regions associated with targets at 2.5 and 7.5 min, but at 15 min, when phagosome closure was complete, little localization remains (Fig. 9, a–f). In contrast, GFP PKC α was cytosolic at all times in Ca2+-depleted cells; no concentration of the GFP signal was detected (Fig. 9, g–l). Analysis of topographic images confirmed the lack of GFP PKC α concentration in Ca2+-depleted cells (data not shown).

FIGURE 9.

GFP PKC α and GFP PKC ε localize to targets during IgG-mediated phagocytosis. RAW cells were transfected with GFP PKC α (a–l) or GFP PKC ε (m–r) and incubated with Alexa 568 BIgG in the presence (a–f) or absence (g–r) of calcium as described in Materials and Methods. Cells were fixed at the indicated times and imaged by confocal microscopy. Images are presented as pairs with the upper image reflecting the merge of the GFP and Alexa 568 signals and the bottom representing the GFP signal alone. A concentration of GFP signal can be seen in regions corresponding to targets at 2.5 and 7.5 min for GFP PKC α in the presence (a–f), but not the absence (g–l), of calcium and for GFP PKC ε in the absence of calcium (m–r); images for GFP PKC ε in the presence of calcium were similar (data not shown). Note that by 15 min, when most phagosomes are closed, little colocalization of GFP and Alexa 568 are apparent. Analysis of topographic images confirm that there is no accumulation of GFP PKC α in calcium-depleted cells. All images were taken at a magnification of ×60 and are representative of six to eight experiments.

FIGURE 9.

GFP PKC α and GFP PKC ε localize to targets during IgG-mediated phagocytosis. RAW cells were transfected with GFP PKC α (a–l) or GFP PKC ε (m–r) and incubated with Alexa 568 BIgG in the presence (a–f) or absence (g–r) of calcium as described in Materials and Methods. Cells were fixed at the indicated times and imaged by confocal microscopy. Images are presented as pairs with the upper image reflecting the merge of the GFP and Alexa 568 signals and the bottom representing the GFP signal alone. A concentration of GFP signal can be seen in regions corresponding to targets at 2.5 and 7.5 min for GFP PKC α in the presence (a–f), but not the absence (g–l), of calcium and for GFP PKC ε in the absence of calcium (m–r); images for GFP PKC ε in the presence of calcium were similar (data not shown). Note that by 15 min, when most phagosomes are closed, little colocalization of GFP and Alexa 568 are apparent. Analysis of topographic images confirm that there is no accumulation of GFP PKC α in calcium-depleted cells. All images were taken at a magnification of ×60 and are representative of six to eight experiments.

Close modal

PKC ε also translocated to forming phagosomes (Fig. 9, m–r). The localization of GFP PKC ε was similar in the presence (data not shown) and absence (Fig. 9, m–r) of Ca2+. That is, GFP PKC ε localized to targets at early time points (2.5 min, 7.5 min; Fig. 9, m, n, p, and q) but this localization was largely lost by 15 min. These images demonstrate that PKC ε is associated with the phagocytic cup. The results confirm the biochemical data (Fig. 8) demonstrating translocation of PKC ε to cell membranes during phagocytosis in Ca2+-depleted cells. The confocal data extend these findings by localizing the PKC ε to the target-associated membranes, consistent with our hypothesis that nPKC is involved in IgG-mediated phagocytosis.

IgG-mediated phagocytosis is a key function of the immune system for identifying and destroying pathogens. Ligation and cross-linking of phagocyte IgG receptors (FcγR) results in particle internalization, but unlike the phagocytosis of complement-opsonized particles or apoptotic cells, FcγR ligation induces macrophages to produce a respiratory burst (1, 21, 24). In monocyte/macrophages, phagocytosis and respiratory burst are increased by PMA and reduced by PKC inhibitors, suggesting a requirement for PKC (5, 6, 8, 15, 18, 25).

Different PKC isoforms are likely to be involved in the multiple signaling pathways from FcγR, and research has focused on identifying required isoforms to study their functions in these pathways. Increased Ca2+-dependent PKC activity has been reported during IgG-mediated phagocytosis in monocytes (8). When monocytes were stimulated by cross-linking FcγR, Ca2+-dependent PKC activity increased and both cPKC β and nPKC ε translocated to the membrane fraction (9). Importantly, PKC α has been observed in nascent and fully formed phagosomes by immunostaining (10). Although these reports provide evidence that cPKCs are activated upon FcγR ligation, they are not consistent with repeated observations that phagocytosis proceeds at exceedingly low [Ca2+]i in monocyte/macrophages and related cell lines (6, 11, 12). The [Ca2+]i measured in these studies (≤2 nM) is well below that required for cPKC isoforms to bind to membranes (0.8–1 μM) or to be fully activated (1.5–38 μM) (26). Thus, although cPKCs may be activated during phagocytosis, they are apparently not required for particle ingestion.

Phagocytosis in RAW 264.7 cells was Ca2+ independent (Fig. 2), increased by PMA and DAG (Fig. 4), and decreased by staurosporine and calphostin C (Fig. 5), indicating that PKC is critical for phagocytosis in these cells. As calphostin C acts as a competitive inhibitor for the DAG binding site on cPKC and nPKC isoforms (27, 28), inhibition of phagocytosis by calphostin C is further evidence that a DAG-requiring PKC isoform is involved. Thus, phagocytosis responded to conditions consistent with activation of a DAG-dependent, Ca2+-independent nPKC isoform.

We hypothesized that PKC activated during phagocytosis would translocate to, and be detected in, the membrane fraction. In this study, PKC α increased in the membrane fraction during phagocytosis, consistent with its activation and previous reports documenting PKC α and Ca2+-dependent PKC activity in phagosomes (8, 9, 10). However, Ca2+ depletion, which did not inhibit phagocytosis, completely prevented membrane association of PKC α and βI (Figs. 3, 8 B, and 9). Both PKCs δ and ε increased in the membrane fraction with a time course consistent with their participation in phagocytosis. PKC δ was present at time points corresponding to the presence of fully formed phagosomes. Importantly, membrane-associated PKC ε increased before and during particle internalization. This agrees with previous reports that PKC activity increases after FcγR ligation but before phagosome closure (8, 9) and that membrane-associated PKC ε is increased by 2–5 min after FcγR cross-linking in monocytes (9). Thus participation of nPKC δ and/or ε in phagocytic signaling is supported by their rapid translocation to membranes upon initiation of IgG-mediated phagocytosis as well as by their activation characteristics.

PKC activity has been implicated in regulation of respiratory burst (14, 25, 34, 35), with cPKC isoforms identified as the primary mediators (13, 14, 17, 23). In our studies, CGP 41251 and Gö 6976 inhibited respiratory burst and not phagocytosis, thus separating these pathways at the level of PKC activation (Fig. 7, A and B). Because PKC α was the only cPKC that translocated to membranes during phagocytosis, this suggests that respiratory burst is mediated by PKC α in RAW 264.7 cells, consistent with results in primary human monocytes (17). A recent report that overexpression of a dominant-negative PKC α in RAW 264.7 cells decreased their ability to kill the intracellular pathogen Leishmania donovani, but had no effect on their phagocytosis (36), provides additional evidence that PKC α is activated, but not required, during phagocytosis.

Our results that the cPKC inhibitor Gö 6976 reduced, but did not eliminate, respiratory burst in neutrophils are in agreement with those reported by Pongracz and Lord (13). In their studies, the magnitude of the reduction (50–60%) was similar to that seen in our experiments (Fig. 7) using the same cPKC inhibitors. Similarly, the down-regulation of PKC β in differentiated HL-60 cells using antisense oligonucleotides produced a partial inhibition of superoxide release (14). In contrast, nonselective PKC inhibitors (Fig. 5 and 7 C) gave a much more complete inhibition of respiratory burst. Although the absence of a respiratory burst in the Ca2+-depleted cells may seem to contradict the partial inhibition of respiratory burst achieved with pharmacological inhibition of cPKC, the lack of respiratory burst in the Ca2+-depleted cells may also be due to the inhibition of other Ca2+-dependent enzymes, notably cPLA2, also required for superoxide production (37). Taken together, these data suggest that the respiratory burst is regulated by multiple signaling pathways, with part of the burst mediated by cPKC and part by other PKC isoforms.

GF109203X is used as a nonisoform-selective inhibitor of PKC (38, 39). Unexpectedly, GF109203X inhibited respiratory burst but not phagocytosis in the same manner as the cPKC-selective drugs (Fig. 7). There are several possible explanations for this discrepancy. One is that the concentrations of GF109203X used in our study were sufficient to inhibit the cPKC but not the nPKC isoforms. The IC50 values given for GF109203X vary with the different isoforms and are lower for PKC α than for PKC δ or ε (39). However, our dose curve (1–10 μM) is at and above the levels previously shown to effect PKC activity and function in vivo (38, 39, 40, 41), so this interpretation seems unlikely.

Alternatively, phagocytosis may require PKC translocation but not kinase activity. GF109203X is a competitive inhibitor of the PKC ATP-binding site (38, 39), and as such should not inhibit the translocation of the PKC isoforms. Calphostin C inhibits the DAG binding site and translocation of PKC ε (27) and presumably other PKC isoforms. This suggests that the translocation and presence of the PKC enzyme, but not its kinase activity, may be required during phagocytic signaling. Precedence for such a mechanism comes from studies on the regulation of neurite outgrowth by PKC ε. Zeidman and coworkers have shown that the catalytic domain of PKC ε is not required to induce neurite outgrowth, and the effect cannot be inhibited by GF109203X, indicating that kinase activity is not required (42). In a similar manner, PKC α has been shown to activate phospholipase D even in the absence of its kinase activity (43). Our results are consistent with the hypothesis that the role of PKC ε in phagocytosis is independent of its kinase activity.

In conclusion, we have separated the signaling pathways for IgG-mediated phagocytosis and respiratory burst at the level of PKC activation. We have demonstrated that cPKC inhibition or Ca2+ depletion inhibited respiratory burst but not phagocytosis. IgG-mediated phagocytosis was Ca2+ independent and DAG sensitive, implying activation of the nPKC isoforms. Both nPKCs δ and ε increased in cell membranes during phagocytosis with a time course consistent with particle ingestion. Our results suggest that PKCs δ and ε translocate and are likely candidates for signal transduction during IgG-mediated phagocytosis in RAW 264.7 cells.

We thank Luiz Rodriguez and Michael Raley for their assistance with the H2O2 assay and Allison Berrier for her critical review of the manuscript. We also acknowledge the Strategic Research Initiative of the Albany Medical Center for its support of the Albany Medical Center Imaging Core.

1

This work was supported by Biomedical Science (to M.R.L.) and Postdoctoral Fellowship (to E.C.L.) grants from the National Arthritis Foundation and by National Institutes of Health Grant S10RR12894 (to J.E.M.).

3

Abbreviations used in this paper: PKC, protein kinase C; DAG, diacylglycerol; GFP, green fluorescent protein; [Ca2+]i, intracellular calcium concentration; PS, phosphatidylserine; nPKC, novel PKC; cPKC, classic PKC; Mg/EGTA, 2 mM MgCl2 and 1 mM EGTA in HBSS; BIgG, IgG opsonized glass beads.

1
Greenberg, S..
1995
. Signal transduction for phagocytosis.
Trends Cell Biol.
5
:
93
2
Karimi, K., M. R. Lennartz.
1998
. Mitogen-activated protein kinase is activated during IgG-mediated phagocytosis, but is not required for target ingestion.
Inflammation
22
:
67
3
Kwiatkowska, K., A. Sobota.
1999
. Signaling pathways in phagocytosis.
Bioessays
21
:
422
4
Lennartz, M. R..
1999
. Phospholipases and phagocytosis: the role of phospholipid-derived second messengers in phagocytosis.
Int. J. Biochem. Cell Biol.
31
:
415
5
Karimi, K., M. R. Lennartz.
1995
. Protein kinase C activation precedes arachidonic acid release during IgG-mediated phagocytosis.
J. Immunol.
155
:
5786
6
Karimi, K., T. R. Gemmill, M. R. Lennartz.
1999
. Protein kinase C and a calcium-independent phospholipase are required for IgG-mediated phagocytosis by Mono-Mac-6 cells.
J. Leukocyte Biol.
65
:
854
7
Jaken, S..
1996
. Protein kinase C isozymes and substrates.
Curr. Opin. Cell Biol.
8
:
168
8
Zheleznyak, A., E. J. Brown.
1992
. Immunoglobulin-mediated phagocytosis by human monocytes requires protein kinase C activation: evidence for protein kinase C translocation to phagosomes.
J. Biol. Chem.
267
:
12042
9
Zheng, L., T. P. Zomerdijk, C. Aarnoudse, R. van Furth, P. H. Nibbering.
1995
. Role of protein kinase C isozymes in Fc γ receptor-mediated intracellular killing of Staphylococcus aureus by human monocytes.
J. Immunol.
155
:
776
10
Allen, L. A., A. Aderem.
1996
. Mechanisms of phagocytosis.
Curr. Opin. Immunol.
8
:
36
11
Di Virgilio, F., B. C. Meyer, S. Greenberg, S. C. Silverstein.
1988
. Fc receptor-mediated phagocytosis occurs in macrophages at exceedingly low cytosolic Ca2+ levels.
J. Cell Biol.
106
:
657
12
Lennartz, M. R., J. B. Lefkowith, F. A. Bromley, E. J. Brown.
1993
. Immunoglobulin G-mediated phagocytosis activates a calcium-independent, phosphatidylethanolamine-specific phospholipase.
J. Leukocyte Biol.
54
:
389
13
Pongracz, J., J. M. Lord.
1998
. Superoxide production in human neutrophils: evidence for signal redundancy and the involvement of more than one PKC isoenzyme class.
Biochem. Biophys. Res. Commun.
247
:
624
14
Korchak, H. M., M. W. Rossi, L. E. Kilpatrick.
1998
. Selective role for β-protein kinase C in signaling for O2− generation but not degranulation or adherence in differentiated HL60 cells.
J. Biol. Chem.
273
:
27292
15
Sakata, A., E. Ida, M. Tominaga, K. Onoue.
1987
. Arachidonic acid acts as an intracellular activator of NADPH-oxidase in Fc γ receptor-mediated superoxide generation in macrophages.
J. Immunol.
138
:
4353
16
Muid, R. E., M. M. Dale, P. D. Davis, L. H. Elliott, C. H. Hill, H. Kumar, G. Lawton, B. M. Twomey, J. Wadsworth, S. E. Wilkinson, et al
1991
. A novel conformationally restricted protein kinase C inhibitor, Ro 31–8425, inhibits human neutrophil superoxide generation by soluble, particulate and post-receptor stimuli.
FEBS Lett.
293
:
169
17
Li, Q., V. Subbulakshmi, A. P. Fields, N. R. Murray, M. K. Cathcart.
1999
. Protein kinase C α regulates human monocyte O2− production and low density lipoprotein lipid oxidation.
J. Biol. Chem.
274
:
3764
18
Schwacha, M. G., P. W. Gudewicz, J. A. Snyder, D. J. Loegering.
1993
. Depression of macrophage respiratory burst capacity and arachidonic acid release after Fc receptor-mediated phagocytosis.
J. Immunol.
150
:
236
19
Labarca, C., K. Paigen.
1980
. A simple, rapid, and sensitive DNA assay procedure.
Anal. Biochem.
102
:
344
20
Shirai, Y., K. Kashiwagi, K. Yagi, N. Sakai, N. Saito.
1998
. Distinct effects of fatty acids on translocation of γ- and ε- subspecies of protein kinase C.
J. Cell Biol.
143
:
511
21
Wright, S. D., S. C. Silverstein.
1983
. Receptors for C3b and C3bi promote phagocytosis but not the release of toxic oxygen from human phagocytes.
J. Exp. Med.
158
:
2016
22
Chanock, S. J., J. el Benna, R. M. Smith, B. M. Babior.
1994
. The respiratory burst oxidase.
J. Biol. Chem.
269
:
24519
23
Kim-Park, W. K., M. A. Moore, Z. W. Hakki, M. J. Kowolik.
1997
. Activation of the neutrophil respiratory burst requires both intracellular and extracellular calcium.
Ann. NY Acad. Sci.
832
:
394
24
Savill, J..
1997
. Apoptosis in resolution of inflammation.
J. Leukocyte Biol.
61
:
375
25
Li, Q., M. K. Cathcart.
1994
. Protein kinase C activity is required for lipid oxidation of low density lipoprotein by activated human monocytes.
J. Biol. Chem.
269
:
17508
26
Keranen, L. M., A. C. Newton.
1997
. Ca2+ differentially regulates conventional protein kinase C’s membrane interaction and activation.
J. Biol. Chem.
272
:
25959
27
Mayne, G. C., A. W. Murray.
1998
. Evidence that protein kinase C ε mediates phorbol ester inhibition of calphostin C- and tumor necrosis factor-α-induced apoptosis in U937 histiocytic lymphoma cells.
J. Biol. Chem.
273
:
24115
28
Kobayashi, E., H. Nakano, M. Morimoto, T. Tamaoki.
1989
. Calphostin C (UCN-1028C), a novel microbial compound, is a highly potent and specific inhibitor of protein kinase C.
Biochem. Biophys. Res. Commun.
159
:
548
29
Germano, P., J. Gomez, M. G. Kazanietz, P. M. Blumberg, J. Rivera.
1994
. Phosphorylation of the γ chain of the high affinity receptor for immunoglobulin E by receptor-associated protein kinase C-δ.
J. Biol. Chem.
269
:
23102
30
Uberall, F., S. Giselbrecht, K. Hellbert, F. Fresser, B. Bauer, M Gschwendt, H. H. Grunicke, G. Baier.
1997
. Conventional PKC-α, novel PKC-ε and PKC-θ, but not atypical PKC-λ are MARCKS kinases in intact NIH 3T3 fibroblasts.
J. Biol. Chem.
272
:
4072
31
Matsuoka, Y., X. Li, V. Bennett.
1998
. Adducin is an in vivo substrate for protein kinase C: phosphorylation in the MARCKS-related domain inhibits activity in promoting spectrin-actin complexes and occurs in many cells, including dendritic spines of neurons.
J. Cell Biol.
142
:
485
32
Watters, D., B. Garrone, G. Gobert, S. Williams, R. Gardiner, M. Lavin.
1996
. Bistratene A causes phosphorylation of talin and redistribution of actin microfilaments in fibroblasts: possible role for PKC-δ.
Exp. Cell Res.
29
:
327
33
Prekeris, R., R. M. Hernandez, M. W. Mayhew, M. K. White, D. M. Terrian.
1998
. Molecular analysis of the interactions between protein kinase C-ε and filamentous actin.
J. Biol. Chem.
273
:
26790
34
Kadri-Hassani, N., C. L. Leger, B. Descomps.
1995
. The fatty acid bimodal action on superoxide anion production by human adherent monocytes under phorbol 12-myristate 13-acetate or diacylglycerol activation can be explained by the modulation of protein kinase C and p47phox translocation.
J. Biol. Chem.
270
:
15111
35
Sergeant, S., L. C. McPhail.
1997
. Opsonized zymosan stimulates the redistribution of protein kinase C isoforms in human neutrophils.
J. Immunol.
159
:
2877
36
St.-Denis, A., V. Caouras, F. Gervais, A. Descoteaux.
1999
. Role of protein kinase C-α in the control of infection by intracellular pathogens in macrophages.
J. Immunol.
163
:
5505
37
Li, Q., M. K. Cathcart.
1997
. Selective inhibition of cytosolic phospholipase A2 in activated human monocytes: regulation of superoxide anion production and low density lipoprotein oxidation.
J. Biol. Chem.
272
:
2404
38
Toullec, D., P. Pianetti, H. Coste, P. Bellevergue, T. Grand-Perret, M. Ajakane, V. Baudet, P. Boissin, E. Boursier, F. Loriolle, et al
1991
. The bisindolylmaleimide GF 109203X is a potent and selective inhibitor of protein kinase C.
J. Biol. Chem.
266
:
15771
39
Martiny-Baron, G., M. G. Kazanietz, H. Mischak, P. M. Blumberg, G. Kochs, H. Hug, D. Marme, C. Schachtele.
1993
. Selective inhibition of protein kinase C isozymes by the indolocarbazole Go 6976.
J. Biol. Chem.
268
:
9194
40
Hundle, B., T. McMahon, J. Dadgar, R. O. Messing.
1995
. Overexpression of ε-protein kinase C enhances nerve growth factor-induced phosphorylation of mitogen-activated protein kinases and neurite outgrowth.
J. Biol. Chem.
270
:
30134
41
Meldrum, D. R., X. Meng, B. C. Sheridan, R. C. McIntyre, Jr, A. H. Harken, A. Banerjee.
1998
. Tissue-specific protein kinase C isoforms differentially mediate macrophage TNF α and IL-1β production.
Shock
9
:
256
42
Zeidman, R., B. Lofgren, S. Pahlman, C. Larsson.
1999
. PKC ε, via its regulatory domain and independently of its catalytic domain, induces neurite-like processes in neuroblastoma cells.
J. Cell Biol.
145
:
713
43
Singer, W. D., H. A. Brown, X. Jiang, P. C. Sternweis.
1996
. Regulation of phospholipase D by protein kinase C is synergistic with ADP-ribosylation factor and independent of protein kinase activity.
J. Biol. Chem.
271
:
4504