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Departments of
*
Biochemistry and
Medicine, Division of Infectious Diseases, Wake Forest University School of Medicine, Winston-Salem, NC 27157
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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(29%)
and small amounts of PKC-
(0.6%), but not PKC-ßII, were present
in cytoskeletal fractions. Upon stimulation with the PKC agonist PMA,
the levels of PKC-, PKC-ßII, and PKC- increased in the
cytoskeletal fraction, concomitant with a decrease in the
noncytoskeletal (Triton-soluble) fractions. PKC- maximally
associated with cytoskeletal fractions at 160 nM PMA and then declined,
while PKC- and PKC-ßII plateaued at 300 nM PMA. Translocation of
PKC- was maximal by 2 min and sustained for at least 10 min.
Translocation of PKC- and PKC-ßII was biphasic, plateauing at 2–3
min and then increasing up to 10 min. Under maximal stimulation
conditions, PKC isoforms were entirely cytoskeletal associated.
Translocation of the NADPH oxidase component
p47phox to the cytoskeletal fraction correlated
with translocation of PKC- and PKC-ßII, but not with translocation
of PKC-. Oxidase activity in cytoskeletal fractions paralleled
translocation of PKC-, PKC-ßII, and
p47phox. Stimulation with
1,2-dioctanoylglycerol resulted in little translocation of PKC isoforms
or p47phox, and in minimal oxidase activity. We
conclude that conventional PKC isoforms (PKC- and/or PKC-ßII) may
regulate PMA-stimulated cytoskeletal association and activation of
NADPH oxidase. PKC- may modulate other PMN responses that involve
cytoskeletal components. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Our primary goal is to understand the roles that both the cytoskeleton and protein kinases play in signaling events associated with the functional responses of PMN. In particular, our focus is the activation of the respiratory burst enzyme, NADPH oxidase. This enzyme is a multicomponent complex capable of converting O2 to O2-, which can be further metabolized to other toxic oxygen species to aid in the destruction of host pathogens (7). NADPH oxidase consists of a flavocytochrome b558 heterodimer (p22phox and gp91phox) and cytosolic components (p47phox, p67phox, p40phox, and Rac 1 or 2) (7, 8, 9). The cytosolic proteins translocate to form a complex with the membrane-associated cytochrome during enzyme activation (10). The signaling mechanisms that regulate the assembly and activation of NADPH oxidase are unclear.
Recent findings have implicated a role for cytoskeletal elements in the regulation of NADPH oxidase activation. In a cell-free system that examines oxidase activation, the addition of G-actin and its subsequent polymerization to F-actin enhances oxidase activity (11). This suggests that polymerized actin may provide a scaffolding to align oxidase components, which allows increased oxidase activity. How this observation relates to in vivo NADPH oxidase activation is unclear, but whole cell studies of the oxidase have demonstrated relationships with the cytoskeleton. Quinn et al. (12) and Mukherjee et al. (13) have demonstrated cosedimentation of NADPH oxidase components and activity with actin- and fodrin-rich membrane subdomains from PMA- and FMLP-stimulated PMN, respectively. Therefore, oxidase components may be localizing to the submembranous cytoskeleton during PMN stimulation. Grogan et al. (14) demonstrated the existence of a p40phox/p67phox/coronin complex in PMN and implicated a role for the oxidase in regulating changes in the cytoskeleton. Coronin is an actin-binding protein identified in Dictyostelium discoideum thought to be involved in cell motility (15). Each of the NADPH oxidase components has been shown to be associated with cytoskeletal (Triton-insoluble) fractions in PMA-stimulated PMN (16, 17, 18, 19). Woodman et al. (18) reported that approximately >95% of measurable oxidase activity was present in cytoskeletal fractions of PMA-stimulated cells. These observations strongly suggest that cytoskeletal elements play a role in NADPH oxidase activation. However, the mechanisms involved are unclear. It is important to identify the interaction between signaling pathways, cytoskeletal elements, and NADPH oxidase components to fully understand the complicated mechanism of oxidase activation and its regulation.
Activation of NADPH oxidase is mediated, at least in part, by
phosphorylation (20). PMA directly binds to and activates
protein kinase C (PKC) isoforms, implying that the
cytoskeletal-associated activation of NADPH oxidase by PMA could be
mediated by PKC. The PKC family of serine/threonine protein kinases is
divided into three classes, based on differential dependencies on
Ca2+ and lipids for activation
(21, 22, 23). The conventional class (cPKC, PKC-, PKC-ßI,
PKC-ßII, and PKC-
) requires Ca2+,
phosphatidylserine (PS), and diacylglycerol (DAG). The novel class
(nPKC, PKC-, PKC-
, PKC-
, PKC-
, and PKC-µ), like the cPKC
class, requires PS and DAG, but is Ca2+
independent. The atypical class (atypical PKC, PKC-
, and PKC-
) is
Ca2+ and DAG independent, but requires PS for
activation. PMA, a DAG analogue, is a potent activator of conventional
and novel PKC isoforms. Five PKC isoforms have been identified in human
PMN. These include: PKC-, PKC-ßI, PKC-ßII (cPKCs); PKC- (a
nPKC); and PKC- (an atypical PKC) (24, 25, 26, 27). Thus,
PKC-, PKC-ß, or PKC- are prime candidates for mediating
PMA-induced responses, as well as responses to receptors that trigger
the liberation of second messengers by phospholipases.
Activation of PKC isoforms is generally associated with translocation
from one site in the cell to another (21, 23).
Associations between PKC isoforms and cytoskeletal proteins have been
demonstrated in some cell types. PKC- colocalizes with MARCKS
(myristoylated alanine-rich C kinase substrate) in macrophages
(28). Blobe et al. (29) demonstrated a
specific association of PKC-ßII, but not PKC-ßI, with actin, which
altered the substrate preferences of the ßII isoform. PKC-
colocalizes with, and is thought to phosphorylate, vimentin, an
intermediate filament, in differentiated HL-60 cells
(30).
Very little is known about the propensity of human neutrophil PKC
isoforms to localize to Triton-insoluble fractions or the relationship
this localization has to NADPH oxidase activation. Redistribution of
p47phox to Triton-insoluble fractions in
response to PMA stimulation was diminished when PMN were pretreated
with the serine/threonine protein kinase inhibitor H-7
(17), implying that the assembly of NADPH oxidase at the
cytoskeleton is mediated by phosphorylation. Curnutte et al.
(17) demonstrated translocation of PKC-ß (using a
general PKC-ß Ab that recognizes both PKC-ßI and PKC-ßII) to
Triton-insoluble fractions. However, other isoforms have not been
studied. In this study, we examined the correlation between the
translocation of three PMA-responsive PKC isoforms (PKC-, PKC-ßII,
and PKC-) in human PMN, the translocation of
p47phox, and the appearance of NADPH oxidase
activity in Triton-insoluble fractions. We used two known PKC
activators, PMA and 1,2-dioctanoyl-sn-glycerol (diC8).
Results indicate that the translocation of cPKC isoforms PKC- and
PKC-ßII correlated most closely with the appearance of
p47phox and NADPH oxidase activity in
cytoskeletal fractions.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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PMA, Triton X-100, PMSF, aprotinin, leupeptin, pepstatin,
n-tosyl phenylalanine chloromethylketone, benzamidine,
iodoacetamide, ferricytochrome c, and
diisopropylfluorophosphate (DFP) were obtained from Sigma (St. Louis,
MO). HBSS and PBS were from Life Technologies (Gaithersburg, MD).
Dextran T-500 was purchased from Pharmacia Biotech (Piscataway, NJ).
Nitrocellulose was obtained from Schleicher & Schuell (Keene, NH).
Superoxide dismutase (SOD) was purchased from DDI Pharmaceuticals
(Mountain View, CA). NADPH was from Boehringer Mannheim (Indianapolis,
IN). The sources of Abs were as follows: anti-PKC-, a mAb
directed against the catalytic domain, was from Upstate Biotechnology
(Lake Placid, NY) (31); anti-PKC-ßII (human
sequence, amino acids 661–673) (32) was from Oxford
Biomedical Research (Oxford, MI); anti-PKC- (human sequence,
amino acids 658–676) was a gift from David J. Burns, formerly of
Sphinx Pharmaceuticals (Durham, NC) (26, 33); polyclonal
Abs raised in goats, anti-p47phox, and
anti-p67phox were gifts from Thomas L. Leto
of the National Institutes of Health (34). HRP-conjugated
secondary Abs were obtained from Transduction Laboratories (Lexington,
KY; goat anti-mouse IgG and goat anti-rabbit IgG) and Organon
Technika (Durham, NC; rabbit anti-goat IgG). A PKC- antigenic
peptide was purchased from Oxford Biomedical Research. PKC- and
PKC- standards were gifts from David J. Burns, formerly of
Sphinx Pharmaceuticals. Rabbit brain cytosol was used as the PKC-ßII
standard and was generated as described under Materials and
Methods for PMN cytosol (35). diC8 was from Serdary
Research Laboratories (London, Ontario, Canada).
PMN isolation
PMN were isolated from heparinized venous blood obtained from healthy human adult donors by dextran sedimentation, Isolymph (Gallard-Schlesinger Industries, Carle Place, NY) centrifugation, and hypotonic lysis of RBC (36). PMN were resuspended at 5 x 107 cells/ml in HBSS supplemented with 4.2 mM sodium bicarbonate and 10 mM HEPES, pH 7.4.
Preparation of Triton-soluble and Triton-insoluble fractions
PMN at 5 x 107 cells/ml were treated with DFP for 5 min (37, 38) and were prewarmed for 5 min at 37°C. At 90 s into the prewarm, NaN3 was added to a final concentration of 1 mM (36). DMSO or ethanol was added as a vehicle control to unstimulated cells. Stimulated cells were treated for varying times at varying concentrations of PMA or diC8. Stimulation reactions were quenched with 45 ml of ice-cold HBSS, followed by centrifugation (250 x g, 10 min, 4°C). Fractions were obtained according to the method of Curnutte et al. (17) with modifications. Briefly, PMN were resuspended in 0.75% Triton X-100, 60 mM PIPES, 25 mM HEPES, pH 7.4, 10 mM EGTA, 2 mM MgCl2, 8 µg/ml aprotinin, 156 µg/ml benzamidine, 1 mM iodoacetamide, 20 µg/ml leupeptin, 100 ng/ml pepstatin, 100 µM n-tosyl phenylalanine chloromethylketone, and 1 mM PMSF, and incubated on ice for 20 min. The cell lysate was layered on the same buffer containing 6% sucrose (1:3, v/v, cell lysate:6% sucrose buffer) and centrifuged at 150,000 x g for 30 min (SW55 rotor; Beckman Instruments, Palo Alto, CA). The top fraction was collected as the noncytoskeletal (Triton-soluble, TS) fraction. The pellet was resuspended by sonication (2 x 10 s) in the same buffer without Triton X-100 and iodoacetamide, and termed the cytoskeletal (Triton-insoluble, TI) fraction. All fractions were stored in 10% glycerol at -70°C. Protein concentrations were determined using the BCA protein assay (Pierce, Rockford, IL) (39) and BSA as standard.
Preparation of cytosol and membrane fractions
PMN at 5 x 107 cells/ml were treated
for varying times at varying concentrations of diC8, with ethanol added
as a vehicle control to unstimulated cells. Reactions were quenched
with a 10-fold excess of cold modified HBSS. Cells were treated with
DFP for 5 min (37, 38) and resuspended at 1 x
108 cells/ml in 50 mM Tris, pH 7.5, 2 mM EGTA, 10
µM benzamidine, 1 µg/ml leupeptin, 10 µM pepstatin, 0.2 µg/ml
aprotinin, 50 mM 2-ME, and 1 mM PMSF. PMN were sonicated to 
90%
breakage on ice and centrifuged at 800 x g for 10 min
to remove nuclei and unbroken cells. The postnuclear supernatant was
layered on a 15%/40% discontinuous sucrose gradient (35)
and centrifuged at 150,000 x g for 30 min at 4°C
(SW55 rotor; Beckman Instruments). The top fraction was collected as
the cytosolic fraction and centrifuged again at 150,000 x
g for 60 min (Ti50 rotor; Beckman Instruments, Palo Alto,
CA). The 15%/40% interface was collected as the membrane fraction.
Protein concentrations were determined using the Pierce Coomassie Blue
Plus protein assay and BSA as a standard.
Lipid phosphorus assay
Lipids were extracted from PMN fractions by modified Bligh and Dyer (40). Lipid phosphorus was determined as described by Rouser et al. (41) with modifications. Briefly, unknown samples were dried under N2 to remove organic solvents, and were oxidized at 180–190°C for 1 h after addition of 150 µl 70% perchloric acid. After cooling, the tubes were rinsed with 900 µl dH2O. Then 167 µl each of 2.5% ammonium molybdate (w/v) and of 10% ascorbic acid (w/v) were added and the tubes were incubated in a 50°C water bath for 15 min. Absorbance was read at 820 nm using Na2HPO4 dilutions as standards.
SDS-PAGE and Western blotting
Samples were prepared for SDS-PAGE according to total protein by
addition of Laemmli sample buffer, boiled for 5 min, and loaded on 7%
(PKC) or 9% (p47 and p67) SDS polyacrylamide gels (42).
After separation, the proteins were transferred (43) to
nitrocellulose overnight. Blots were stained with Ponceau-S (0.2%
Ponceau-S, 3% TCA, and 3% sulfosalicylic acid) (44) to
visualize m.w. markers and were destained with deionized water,
followed by Tris-buffered saline with 0.1% Tween-20 (TBS-T). The blots
were blocked with TBS-T containing 5% nonfat dry milk for 60 min,
washed, and incubated with the primary Ab diluted in 3% BSA/0.02%
NaN3 in PBS, pH 7.3, for 2 h
(anti-PKC-, 1:500; anti-PKC-ßII, 1:100; anti-PKC-,
1:1500; anti-p47phox, 1:1000;
anti-p67phox, 1:1000). Then the blots were
washed and incubated with HRP-conjugated secondary Ab for 60 min. The
blots were washed again, then incubated with enhanced chemiluminescence
(ECL) reagent (Pierce, Rockford, IL) for 1 min, and exposed to film for
visualization. Autoradiograms were analyzed and quantified by
densitometry (PDI, Huntington Station, NY).
NADPH oxidase assay
Cytoskeletal fractions from stimulated PMN were assayed for
NADPH oxidase activity as the SOD-inhibitable reduction of
ferricytochrome c, as described by Caldwell et al.
(35) with modifications. Briefly, reaction tubes contained
48 mM KPO4 buffer (pH 7), 10 µM flavin adenine
dinucleotide, 76 µM ferricytochrome c, 1 mM EGTA, 7.6 mM
MgCl2, 1 µM GTPS, 0.15 mM SDS, and 60–120
µg Triton-insoluble protein in a final volume of 237.5 µl. SOD
(final concentration, 50 µg/ml) was added to one-half the reaction
mixture. Addition of NADPH (final concentration, 0.2 mM) started the
reaction, which was monitored at 550 nm on a UV-2401PC Shimadzu
(Columbia, MD). Superoxide produced was calculated from the linear
slopes, using an extinction coefficient of 21
mM-1cm-1
(45), and normalized to Triton-insoluble protein.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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We first characterized the Triton-soluble and Triton-insoluble
fractions for their protein and phospholipid content. Total protein was
divided equally, 2.16 ± 0.06 and 2.17 ± 0.09 mg
protein/108 cell equivalents (n =
11) in Triton-soluble and Triton-insoluble fractions, respectively.
This protein distribution was comparable with detergent fractionations
previously reported in PMN (46). Samples from several
experiments were subjected to SDS-PAGE and stained with Coomassie
blue. The pattern of proteins in each fraction was consistent between
experiments and similar to that reported by Woodman et al.
(18) (data not shown). Additionally, the prominent protein
bands did not change with cell stimulation. The Triton-soluble
fraction, as expected, contained the majority of the phospholipid
(190 nmol phospholipid/mg protein), while the Triton-insoluble
fraction contained 30 nmol phospholipid/mg protein. These values did
not change with cell stimulation. This phospholipid distribution was
similar to that for detergent fractionations previously reported in PMN
(47) and other cells (48).
Translocation of PKC and p47phox and Triton-insoluble NADPH oxidase activation are dependent on the concentrations of PMA during stimulation
An assessment of PKC isoform translocation as a measure of
activation in Triton-soluble and Triton-insoluble fractions was chosen
for the following reasons. First, measurement of PKC activity in
Triton-containing fractions is problematic, due to the inhibitor effect
of the detergent (data not shown). Second, activity measurements do not
differentiate between the activities of individual PKC isoforms within
a class. Finally, PKC translocation generally is indicative of PKC
activation (21, 23). Fig. 1
A shows the concentration
dependence of PKC translocation from Triton-soluble to Triton-insoluble
fractions. Isolated human PMN were stimulated with concentrations of
PMA ranging from 0–1600 nM for 5 min and separated into Triton-soluble
and Triton-insoluble fractions. Decreases in the level of each PKC
isoform in the Triton-soluble fractions, concomitant with increases in
the Triton-insoluble fractions, were observed with increasing PMA
concentrations. There was an almost complete loss of PKC in the
Triton-soluble fractions with cell stimulation, as only low percentages
of the total present in unstimulated cells were present after treatment
with 1600 nM PMA (PKC-, 5.5%; PKC-ßII, 2.7%; PKC-, 5.4%;
based on densitometric analysis). PKC-ßII (33% total recovery at
1600 nM PMA, compared with unstimulated cells) appears to be more
sensitive to degradation than PKC- or PKC- (67.4% and 39.3%
recoveries, respectively).
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B). Translocation of
p47phox to Triton-insoluble fractions occurred
in response to PMA stimulation, in a concentration-dependent manner.
Densitometric analysis revealed that 43% of
p47phox remained Triton soluble at the highest
concentration of PMA used, and the recovery of this protein was 99.3%.
Conversely, p67phox was entirely Triton
insoluble under all conditions, even in unstimulated PMN. These results
confirm and extend previous observations (17, 18) of the
localization of these proteins before and after PMA stimulation,
supporting the validity of our method. As a result of its unchanging
nature, p67phox served as a good marker for
protein integrity in the Triton-insoluble fractions under the various
conditions of cell stimulation.
Fig. 2A shows the percentage
of the total of each PKC isoform and of p47phox
in the Triton-insoluble fractions summarized from several experiments,
as assessed by densitometry. Two patterns of PKC isoform translocation
are observed. Translocation of PKC- was apparent at 30 nM PMA,
reached a maximum at 160 nM PMA and then declined. In contrast, the
concentration curve for translocation of cPKC- and cPKC-ßII was
shifted to the right and did not decline. The concentration curves for
translocation of p47phox (Fig. 2A)
and the appearance of NADPH oxidase activity in the Triton-insoluble
fraction (Fig. 2B) were similar to the curves for
translocation of the cPKCs. However, the translocation of
p47phox plateaued at 160 nM, instead of 300 nM
PMA. The similarities in concentration dependence of translocation of
cPKCs and p47phox and appearance of
Triton-insoluble NADPH oxidase activity, coupled with the different
concentration dependence of translocation of nPKC-, suggest the
involvement of cPKC in NADPH oxidase regulation.
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We determined the time course for PMA-induced translocation of PKC
isoforms and p47phox, shown in Fig. 3, A and B,
respectively. Human PMN were stimulated with 300 nM PMA for 0–10 min,
and were separated into Triton-soluble and Triton-insoluble fractions,
as described in Materials and Methods. A control for cell
exposure to the stimulus during preparation of the Triton-soluble and
Triton-insoluble fractions was included, in which either the vehicle
(DMSO, 0v) or the stimulus (PMA, 0p) was added at the time the
stimulation reactions were terminated. Slight effects on the
translocation of PKC and p47phox were apparent
in the 0p control, indicating that PMA was not completely inert during
the processing period. However, the effects of PMA were more dramatic
during the time course conducted at 37°C. A rapid decrease in the
level of each PKC isoform in Triton-soluble fractions was observed,
with most of each isoform disappearing by 3 min (Fig. 3A).
Concomitant increases in the Triton-insoluble fraction occurred. In
some experiments, PKC-ßII appeared to return to the Triton-soluble
fraction at the 10-min time point, but this finding was inconsistent.
PMA also induced a time-dependent decrease in the level of
p47phox in Triton-soluble fractions, with a
concomitant increase in Triton-insoluble fractions (Fig. 3B). However, in contrast to the PKC isoforms, 38% of
p47phox remained in the Triton-soluble fraction
at the 10-min time point. Similar to the findings in Fig. 1B, p67phox was entirely associated
with the Triton-insoluble fraction, and the level of this protein did
not change with time of PMA stimulation.
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A and less evident in Fig. 1A, PKC-
appears to be undergoing a gel shift as a result of PMA stimulation.
Using the antigenic peptide for the PKC- Ab, we confirmed that both
bands visualized on the blots were PKC- (data not shown). This shift
in mobility may be the result of phosphorylation of PKC- (see
Discussion). For purposes of quantitation by densitometry
(see Figs. 2, 4, and 5), the two PKC- bands were added together for
any given condition.
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A summarizes the
appearance of PKC isoforms and p47phox in the
Triton-insoluble fractions during the PMA time course, using
densitometric analysis. As in the concentration curve experiments, two
patterns were observed. The redistribution of cPKCs and
p47phox to the Triton-insoluble fractions has a
biphasic appearance, with an initial increase and a plateau at 2–3
min, then a further increase and a second plateau between 5 and 10 min.
The curves for PKC- and PKC-ßII are similar, although the level of
PKC-ßII (as percentage of total PKC-ßII in unstimulated PMN) was
lower due to less recovery of this isoform. The appearance of
p47phox in the Triton-insoluble fraction
slightly preceded the appearance of PKC- and PKC-ßII. In contrast
to the pattern with the cPKCs and p47phox, the
level of PKC- in the Triton-insoluble fraction did not change after
the maximum increase was reached at 2 min. Like PKC-, PKC-ßII, and
p47phox, the time course for the appearance of
Triton-insoluble NADPH oxidase activity has a biphasic appearance (Fig. 4B). After an initial 30-s lag, oxidase activity
dramatically increased up to 2 min. There was no change in activity
from 2–5 min, but there was a further increase at the 10-min time
point. Thus, appearance of oxidase activity in the Triton-insoluble
fraction lags behind the appearance of p47phox
and PKC isoforms and resembles the pattern of translocation of the
cPKCs, but not that of nPKC-. This suggests that translocation and
phosphorylation of oxidase components by a cPKC isoform must occur
before superoxide production is observed. Comparison between DAG and PMA for inducing translocation of PKC and p47phox and appearance of Triton-insoluble NADPH oxidase activity
The actions of PMA on neutrophils can be mimicked, to a limited
extent, by more physiological PKC activators, such as cell-permeable
DAGs. We examined whether diC8, a cell-permeable DAG, affected the
distribution of PKC isoforms and p47phox in
Triton-soluble and Triton-insoluble fractions, as well as appearance of
NADPH oxidase activity in Triton-insoluble fractions. PMN were treated
with either solvent (unstimulated), 70 µM diC8 for 3 min, or 300 nM
PMA for either 3 or 10 min. Preliminary experiments determined that
these conditions of diC8 stimulation resulted in maximum effects.
Triton-soluble and Triton-insoluble fractions were prepared and the
summarized levels of each protein and the level of NADPH oxidase
activity in the Triton-insoluble fractions are shown in Fig. 5. In unstimulated PMN, 29% of the total
PKC- was Triton insoluble. In contrast, only 0.1% of PKC- and no
PKC-ßII were Triton insoluble in unstimulated PMN. Similarly, only
4.2% of p47phox and virtually no NADPH oxidase
activity were present in Triton-insoluble fractions from unstimulated
cells.
DiC8 treatment induced only small changes in the distribution of PKC
isoforms and p47phox and in the level of NADPH
oxidase activity. Of the PKC isoforms, only PKC- levels showed an
increase (10% change in distribution) in Triton-insoluble fractions in
response to diC8. This was accompanied by a similar small increase
(10.8% change in distribution) in the level of
p47phox, and a small increase in NADPH oxidase
activity (unstimulated: 0.001 nmol O2-/min/mg;
diC8-stimulated: 0.03 nmol O2-/min/mg). As
already noted, PMA induced substantial translocation of PKC isoforms
and p47phox, as well as in appearance of NADPH
oxidase activity. The two patterns observed in Fig. 4 were also
apparent in this experiment, in that there were increases between 3 and
10 min in Triton-insoluble levels of PKC-, PKC-ßII, and
p47phox, and in oxidase activity, but not in the
level of PKC-. Taken together, these data with different agonists
add support to the concept of a closer relationship between cPKC
isoforms and NADPH oxidase activation, than between nPKC- and the
oxidase.
Although diC8 induced little (PKC-) or no (PKC-ßII, PKC-)
measurable translocation of PKC isoforms to Triton-insoluble fractions,
effects of this agonist on levels of PKC isoforms in Triton-soluble
fractions were noted (data not shown). PKC- levels were reduced in
the Triton-soluble fraction from 99.9% in unstimulated PMN to 94.4%
with diC8 treatment, consistent with the appearance of 10% of
PKC- in the Triton-insoluble fraction. The level of PKC-ßII in
Triton-soluble fractions declined from 100% in fractions from
unstimulated PMN to 89.4% in fractions from diC8-treated PMN and that
of PKC- declined from 70.8% to 58.4%. These decreases in the
Triton-soluble fraction without corresponding increases in the
Triton-insoluble fraction imply that diC8 does activate PKC-ßII and
PKC- and increases their sensitivity to proteolysis.
DiC8 induces translocation of PKC-ßII and PKC- from cytosol to
membrane fractions
To confirm the ability of diC8 to activate PKC-ßII and PKC-
in PMN, we examined whether this agonist stimulated translocation of
these isoforms to membrane fractions. Previous studies using
oleoyl-acetyl-glycerol had shown that this DAG could induce
translocation of PKC activity from cytosol to membrane fractions
(49). As shown in Fig. 6,
treatment of PMN with diC8 for 30 s resulted in a
concentration-dependent translocation of PKC-ßII and PKC- from
cytosol to membrane fractions, which began to plateau at 70 µM diC8.
Higher concentrations of diC8 were not tested, because they appeared to
partially solubilize the PMN. Virtually all of the cytosolic PKC-
disappeared at 25 and 70 µM diC8, with recovery of 15–20% of the
isoform in the membrane fraction. Thus, 80% of the PKC- present
in membrane and cytosolic fractions from unstimulated cells was not
recovered in either fraction after maximal diC8 treatment, suggesting
compartmentalization and/or proteolysis of this isoform. Translocation
of some PKC- to fractions other than the membrane also is implied by
the observation that the total recovery of PKC- in Triton-soluble
and Triton-insoluble fractions was higher (60%, see above). In
contrast, the effects of diC8 on the translocation of PKC-ßII were
less dramatic. The level of PKC-ßII in cytosolic fractions decreased
by 20–25% in response to 25–70 µM diC8, accompanied by an
increase in membrane fractions from 0% in unstimulated cells to 3–5%
with diC8 stimulation. Thus, total recovery of PKC-ßII after maximal
diC8 treatment was 80–85%, suggesting less compartmentalization or
proteolysis of this isoform compared with PKC-.
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to membrane fractions was also
dependent on the time of diC8 treatment (Fig. 7). Translocation in response to diC8
occurred rapidly, with close to maximal effects at 15 s and
maximal effects at 30 s for both isoforms. There was a slight
tendency for both isoforms to partially return from the membrane to the
cytosol at 60 and 120 s, but the changes were small. Overall, the
results in
Figs. 5–7 demonstrate that stimulation of PMN with diC8
induces translocation of PKC-ßII and PKC- to membrane fractions,
but not to Triton-insoluble fractions.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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,
PKC-ßII, and PKC- isoforms to Triton-insoluble fractions.
Detergent-insoluble cellular fractions have been used by many
investigators to represent the cytoskeleton (16, 17, 18, 19, 46, 47, 50, 51, 52, 53); thus, we speculate that these PKC isoforms translocate
to cytoskeletal elements in response to PMA. This suggests that at
least part of the function of these isoforms in PMN is conducted by
regulation of cytoskeletal-mediated events. Furthermore, different
patterns of translocation were observed for the cPKC isoforms, compared
with nPKC-, suggesting that these classes of PKC have different
translocation mechanisms, different localizations within the
Triton-insoluble fractions, and/or different functional roles. Indeed,
we found that the pattern of p47phox
translocation and NADPH oxidase activation more closely resembled the
pattern of cPKC translocation. Thus, our data suggest that the
conventional class of PKC isoforms regulates the activation and/or
cytoskeletal localization of NADPH oxidase.
Other reports have suggested that cPKCs participate in NADPH oxidase
activation. Majumdar et al. (54) demonstrated that
p47phox was a better substrate for partially
purified neutrophil PKC-ß than for an unidentified
Ca2+-independent PKC. Additionally, a
pseudosubstrate peptide that inhibited PKC-ß, but not the
Ca2+-independent PKC, reduced
O2- release in
electropermeabilized PMN. Duyster and coworkers (55)
implicated a role for cPKC-ß over nPKC- in
O2- release by rat liver
macrophages. Recently, Korchak et al. (56) used PKC-ß
antisense oligonucleotides to reduce PKC-ß levels in HL-60 cells,
differentiated to a PMN-like phenotype. The greatest effect on
O2- release was seen in
FMLP-stimulated PMN, with modest effects on
O2- release induced by PMA or a
phagocytic agonist. A similar study (57), performed in
normal human monocytes, demonstrated marked inhibition of
O2- release induced by
opsonized zymosan in cells treated with PKC-, but not PKC-ß
antisense. Our results support the possibility that cPKC-
participates in the regulation of NADPH oxidase, but we cannot exclude
a role for cPKC-ß.
The translocation of PKC- and PKC-ßII to the Triton-insoluble
fraction preceded NADPH oxidase activation and occurred at similar
concentrations of PMA. Thus, cPKC isoforms could either regulate the
interaction of NADPH oxidase components with cytoskeletal elements
and/or regulate NADPH oxidase activity after the enzyme complex becomes
cytoskeletal associated. The inhibition of the translocation of
p47phox to cytoskeletal fractions by
nonselective protein kinase inhibitors (17) suggests that
PKC and/or other protein kinases may regulate this process. However,
comparison of the patterns of translocation of
p47phox and the cPKCs shows that translocation
of p47phox occurred at slightly earlier time
points and at similar (cPKC-) or lower (cPKC-ßII) concentrations
of PMA. This suggests that NADPH oxidase components, in particular
p47phox, could regulate association of cPKC
isoforms with the cytoskeleton. Curnutte et al. (17)
reported marked reduction of PMA-stimulated translocation of cPKC-ß
in neutrophils from patients with a form of chronic granulomatous
disease, who are missing p47phox, lending
support to this possibility. Thus, the relationships between PKC,
cytoskeletal elements, and the oxidase are likely to be complex.
Most of the components of NADPH oxidase are phosphorylated during stimulation of intact PMN (58, 59, 60, 61, 62), so the cPKCs may directly phosphorylate one or more components. Indeed, a mixture of the cPKCs purified from rat brain, as well as partially purified cPKC from human PMN, phosphorylate p47phox in vitro (54, 63). In a cell-free system developed by Babior and colleagues (64), the cPKC-mediated phosphorylation of p47phox is a necessary step for NADPH oxidase activation to occur. However, other protein kinases can phosphorylate p47phox in vitro (63, 65, 66), and other oxidase components are in vitro substrates for protein kinases besides PKC (62, 67). Additionally, the regulation by PKC could be at the level of intermediate cytoskeletal-associated proteins. Many cytoskeletal-associated proteins are substrates for PKC (28, 30, 68, 69, 70), and cytoskeletal reorganization is regulated by phosphorylation (6, 71). Thus, it is likely that multiple phosphorylation-dependent mechanisms contribute to the regulation of NADPH oxidase.
To further address the relationship between PKC and NADPH oxidase, it
is important to identify the cytoskeletal elements with which PKC and
oxidase components interact. Previous results implicate interactions
between neutrophil NADPH oxidase components and the actin filament
system (11, 12, 13, 14). Similarly, cPKC- and cPKC-ß are
reported to bind, phosphorylate, or colocalize with actin or
actin-binding proteins in macrophages and other cell types (28, 29, 69, 72, 73, 74, 75). Such studies have not been performed in PMN.
PMN stimulated with PMA exhibit an increase in F-actin content
(76) and membrane ruffling (77, 78). Thus,
the actin-based cytoskeleton is likely to participate in the
interactions between cPKC and NADPH oxidase components. However,
actin-filament disrupting agents (e.g., cytochalasin B) do not affect
superoxide release stimulated by PMA (36), possibly
because the submembranous actin network is not disrupted
(13). NADPH oxidase components and activity previously
have been localized to the submembranous cytoskeleton (12, 13). Thus, it will be necessary to more precisely localize the
cytoskeletal proteins interacting with the cPKCs and NADPH oxidase
components, as well as to determine whether PKC isoforms and oxidase
components colocalize.
Translocation of PKC isoforms to the Triton-insoluble fraction occurs at a slower rate than that observed for cytosol to membrane translocation, which is complete by less than 1 min (Ref. 26 , and data not shown). PKC may first associate with the membrane (where lipid activators are present) and then dock with cytoskeletal (Triton-insoluble) proteins that serve as substrates or anchor PKC in proximity with its substrate. In fact, many PKC-binding proteins bind to PKC in a PS-dependent manner (72, 79), suggesting that, in intact cells, membrane association of PKC may be required before binding of PKC to proteins can occur.
It is interesting that, unlike PMA, diC8 induced low levels of only
cPKC- translocation and NADPH oxidase activation in the
Triton-insoluble fraction. DAGs are less potent than PMA for PKC
activation and activation of NADPH oxidase in whole cells, although the
optimal rates of O2- release
are comparable (49, 80). However, effects of exogenously
added DAGs on PKC and functional responses are transient, while those
to PMA are more sustained (81, 82), most likely due to
rapid metabolism of the DAG (81, 83). Therefore, the low
levels of Triton-insoluble NADPH oxidase activity induced by diC8 may
be caused by the rapidly reversible nature of the activation process,
similar to results obtained with FMLP (36). The low
ability of diC8 to induce translocation of PKC isoforms to
Triton-insoluble fractions also may be related to the transient nature
of PKC activation. Alternatively, recent evidence in living cells
suggests that PKC may localize to different membrane domains, when PMA
and diC8 are compared as agonists (84).
Although the translocation of the cPKCs to the Triton-insoluble
fraction is linked to activation of the respiratory burst, the role of
cytoskeletal-associated nPKC- in PMN is not known. About 30% of
nPKC- was Triton insoluble in unstimulated PMN (Fig. 5), implying a
constitutive cytoskeletal association, perhaps with the intermediate
filament system (30). PMA induced rapid translocation of
the remainder of the isoform from the Triton-soluble to the
Triton-insoluble fraction. This response was clearly more sensitive
than the response of the cPKCs, occurring at lower concentrations of
PMA and at a faster rate. Translocation also was accompanied by a shift
in mobility to slower migrating species, possibly as a result of
tyrosine phosphorylation by Src family members (85, 86, 87).
Members of the Src family of protein tyrosine kinases localize to
Triton-insoluble fractions of human PMN upon PMA stimulation (50, 51) and, thus, could phosphorylate nPKC-. Further studies are
needed to test this possibility and to determine the effects of these
putative phosphorylation events on nPKC- function in PMN.
The association of PKC isoforms with the cytoskeleton in PMN has additional functional implications for the regulation of PKC directly. Triton-insoluble proteins may serve as an alternate mechanism for PKC activation. In MOLT-4 cells, Blobe et al. (29) demonstrated that PKC-ßII was fully activated in the presence of Mg2+ when bound to actin in the absence of Ca2+ and lipids. Also, Triton-insoluble association may represent a means of PKC down-regulation, in which binding to proteins could make the protease-sensitive hinge region more accessible to proteolysis. This latter possibility is supported by our observation that PMA stimulation of PMN decreased the total recovery of each PKC isoform, in conjunction with translocation to the Triton-insoluble fraction.
In conclusion, we report the translocation of the PMA-responsive
isoforms of PKC to the cytoskeleton (detergent-insoluble fractions) in
PMN. The pattern of translocation was PKC class specific, and a close
correlation between cytoskeletal association of cPKC isoforms and NADPH
oxidase activity was established. The results strengthen the concept
that PKC regulates functional responses in PMN through
cytoskeletal-based interactions. To further understand the relationship
between PKC, the cytoskeleton, and the activation of NADPH oxidase,
studies are currently underway in our laboratory to determine whether
oxidase components colocalize with PKC- or PKC-ßII in the
Triton-insoluble fractions after PMN stimulation, and to identify other
proteins in that fraction with which PKC isoforms associate.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Linda C. McPhail, Department of Biochemistry, Wake Forest University School of Medicine, Medical Center Boulevard, Winston-Salem, NC 27157-1016. E-mail address:
3 Abbreviations used in this paper: PMN, polymorphonuclear leukocyte or neutrophil; PKC, protein kinase C; cPKC, conventional PKC; DAG, diacylglycerol; DFP, iisopropylfluorophosphate; diC8, 1,2-dioctanoyl-sn-glycerol; ECL, enhanced chemiluminescence; nPKC, novel PKC; phox, phagocyte oxidase; PS, phosphatidylserine; SOD, superoxide dismutase.
Received for publication December 31, 1998. Accepted for publication August 5, 1999.
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