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J. Biol. Chem., Vol. 275, Issue 42, 32566-32571, October 20, 2000
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From the
Received for publication, June 21, 2000, and in revised form, August 4, 2000
In activated neutrophils NADPH oxidase is
regulated through various signaling intermediates, including
heterotrimeric G proteins, kinases, GTPases, and phospholipases.
ADP-ribosylation factor (ARF) describes a family of GTPases associated
with phospholipase D (PLD) activation. PLD is implicated in NADPH
oxidase activation, although it is unclear whether activation of PLD by
ARF is linked to receptor-mediated oxidase activation. We explored
whether ARF participates in NADPH oxidase activation by
formyl-methionine-leucine-phenylalanine (fMLP) and whether this
involves PLD. Using multicolor forward angle light scattering analyses
to measure superoxide production in differentiated neutrophil-like
PLB-985 cells, we tested enhanced green fluorescent fusion proteins of
wild-type ARF1 or ARF6, or their mutant counterparts. The ARF6(Q67L)
mutant defective in GTP hydrolysis caused increased superoxide
production, whereas the ARF6(T27N) mutant defective in GTP binding
caused diminished responses to fMLP. The ARF1 mutants had no effect on
fMLP responses, and none of the ARF proteins affected phorbol
12-myristate 13-acetate-elicited oxidase activity. PLD inhibitors
1-butanol and 2,3-diphosphoglycerate, or the ARF6(N48R) mutant assumed
to be defective in PLD activation, blocked fMLP-elicited oxidase
activity in transfected cells. The data suggest that ARF6 but not ARF1
modulates receptor-mediated NADPH oxidase activation in a
PLD-dependent mechanism. Because PMA-elicited NADPH oxidase
activation also appears to be PLD-dependent, but
ARF-independent, ARF6 and protein kinase C may act through distinct
pathways, both involving PLD.
The phagocyte NADPH oxidase is an important innate defense system
against bacterial and fungal infections. Inherited deficiencies of this
enzyme result in chronic granulomatous disease, which is characterized
by enhanced susceptibility to microbial infection and dysregulated
inflammatory responses. Although the components of this
superoxide-generating system have been the subject of intensive
investigation, the signaling mechanisms responsible for oxidase
activation (the respiratory burst) are complex and not clearly defined
(1). Many studies indicate that several GTPases of the Ras superfamily
are involved at various levels of regulation of this inflammatory
process (2). Rac was identified as a third cytosolic component required
for activation of the NADPH oxidase, with Rac1 as the active component
in guinea pig macrophages (3) and Rac2 in human neutrophils (4). In
addition, Rap1A, which localizes to the plasma membrane and granule
membranes in human neutrophils, was shown to associate with cytochrome
b558 of the oxidase (5). Mutant Rap1A inhibits
oxidase activity in transfected B cells (6), although its role in the
system is not entirely clear.
The ADP-ribosylation factor
(ARF)1subfamily of
Ras-related proteins consists of six mammalian GTPases (ARF1-ARF6),
five of which are detected in man (ARF1, 3-6)) (7). Originally
identified as cofactors required for cholera toxin-catalyzed
ADP-ribosylation of G All human ARF mRNA species have been detected in HL-60 cells (20),
and several of these ARF isoforms (ARF 1, 5, and 6) appear to activate
rat brain PLD (21). The best characterized ARF protein, ARF1, is
localized to the Golgi complex and is critical for vesicular transport
along secretory pathways (9). Unlike ARF1, ARF6, which is the least
conserved of the human ARF proteins, localizes at the cell periphery
and cycles between the plasma membrane and endosomal compartments in a
guanine nucleotide-dependent manner (22, 23). ARF6 was
characterized as a regulator of membrane trafficking (22, 23) and
remodeling of the plasma membrane and the underlying cytoskeleton
(24-26). ARF6 has been linked functionally to PLD activation. In
several mammalian cells ARF6(T27N) colocalizes with hPLD1a and hPLD1b
(27), whereas in chromaffin cells ARF6 appears to activate PLD in
vivo during exocytosis (28). Both ARF6 and PLD have been
implicated in cells undergoing phagocytosis. ARF6 mutants defective in
GTP binding (T27N) or GTP hydrolysis (Q67L) inhibit Fc In light of the growing body of evidence linking ARF activation to
receptor stimulation of phagocytes, and findings linking PLD activation
to ARF, as well as to receptor-mediated oxidative responses, we
explored the possible involvement of ARF1 and ARF6 in NADPH oxidase
activation and whether this involves participation of PLD. For this
purpose, we used the PLB-985 cell line induced to differentiate into a
neutrophil-like phenotype following treatment with dibutyryl cAMP
(Bt2cAMP). In previous work (30) we demonstrated that these cells are readily transfected while exhibiting phenotypic traits of differentiated phagocytes. Using this model, we explored possible roles of ARF1 and ARF6 through transfection of mutated forms
produced as fusions with enhanced green fluorescent protein (EGFP), and
we demonstrated involvement of both ARF6 and PLD in formyl-methionine-leucine-phenylalanine (fMLP) receptor-mediated activation of the respiratory burst. In contrast, phorbol 12-myristate 13-acetate (PMA) activation of the oxidase appears to be
PLD-dependent, but ARF independent, suggesting that ARF and
protein kinase C act through different signaling pathways
leading to oxidase activation, both apparently involving activation of
PLD.
Cell Culture--
PLB-985 cells were grown in stationary
suspension cultures in RPMI 1640 medium containing 10% bovine serum
(Hyclone Laboratories, Inc., Logan, UT), 2 mM
L-glutamine, 100 units/ml penicillin, 100 µg/ml
streptomycin, and 12.5 units/ml nystatin at 37 °C, in a humidified
atmosphere of 5% CO2. Cell number and viability were determined by trypan blue exclusion.
Transfection of PLB-985 Cells--
To induce a granulocytic
phenotype, PLB-985 cells were pretreated with 0.3 mM
Bt2cAMP (Sigma) for 3 days. The cells were then transfected 24 h after Bt2cAMP induction and grown in
RPMI 1640 medium containing 20% bovine serum in the presence of 0.3 mM Bt2cAMP for an additional 2 days. For
transient transfections, PLB-985 cells (1 × 107 cells
washed in 0.3 ml of cold Ca2+- and Mg2+-free
PBS) were electroporated (99 µs, three pulses, 0.9 volt, using a BTX-
Electro square porator T 820) and 20 µg each of plasmid DNA. 48 h after electroporation (3 days postinduction), the living PLB-985
cells were obtained from Ficoll density gradients (30) and used for
flow cytometric analysis. The ARF cDNAs (ARF6-WT, ARF6(Q67L),
ARF6(T27N), ARF1-WT, ARF1(Q71L), ARF1(T31N)) were cloned into the
vector pEGFP-N1 (CLONTECH) and fused to the EGFP of
Aequorea victoria (provided by Dr. Guillermo Romero,
University of Pittsburgh (31)).
Cell Labeling and NADPH Oxidase Activity Measurements by Flow
Cytometry--
Flow cytometric assays of NADPH oxidase activity
involved the method from Model et al. (32) with slight
modifications. After Ficoll gradient separation, the PLB-985 cells
(107 cells/ml) were labeled in Ca2+- and
Mg2+-free PBS containing 15 µM
4-carboxydihydrotetramethylrosamine succinimidyl ester (Ros-SE;
OxyBURST Orange-RE, Molecular probes, Eugene, OR), incubated on ice,
and shaken in the dark for 30 min. After labeling, the cells were
washed twice with cold Ca2+- and Mg2+-free PBS,
resuspended at a concentration of 107 cells/ml in cold PBS,
and kept on ice protected from light. The assay mixture for oxidase
activity contained 100 µl of ice-cold cell suspension
(106 cells), 900 µl of Hanks' balanced saline solution
with Ca2+ and Mg2+ and 100 units/ml
horseradish peroxidase (Sigma). The cells were prewarmed for 5 min in a
37 °C shaking water bath followed by the addition of activators (1 µM fMLP or 100 ng/ml PMA). Cells stimulated with fMLP
were incubated for an additional 3 min; cells stimulated with PMA were
incubated for an additional 15 min. After incubation the reactions were
stopped by placing the tubes on ice and analyzed by flow cytometry
within 1-2 h.
Flow Cytometry--
Flow cytometric analysis was performed on
0.5 × 106 cells using a Becton Dickinson FACStar Plus
flow cytometer (San Jose, CA), equipped with an argon laser operating
at 488 nm and spectra physics 2020 argon laser pumping a 355 dye laser
head operating at 590 nm. Relative fluorescence signals were collected
in four-decade log 10 amplifiers and reported as linear geometric mean
fluorescence value. To detect oxidized rosamine fluorescence the cells
were excited at 488 nm, and the emission was detected with a 575/26 nm
bandpass filter. EGFP fluorescence was detected with the same excitation wavelength using 512/20 nm bandpass filter. Because EGFP-transfected cells exhibited a broad range of EGFP expression levels, data collected from these cells were subdivided further into
three regions based on fluorescence intensities: region 1 (R1, EGFP
fluorescence = 3 × 101-102); region
2 (R2, EGFP fluorescence = 102-103), and
region 3 (R3, EGFP fluorescence = 103-104).
Detection of gp91phox by FACS Analysis--
Transfected cells
(106) were washed twice, resuspended in 100 µl of
Ca2+- and Mg2+-free PBS, and incubated for 30 min at room temperature with mouse anti-gp91phox antibody
(7D5), which reacts with an extracellular epitope (33). The cells were
then washed twice with PBS and incubated with goat anti-mouse IgG (H+L)
CY5-conjugated F(ab')2 fragment (Jackson Immunoresearch
Laboratories, Inc., West Grove, PA) for 30 min at room temperature in
the dark. After washing, the cells were analyzed on the flow cytometer,
and gp91phox was detected by excitation at 590 nm using a
668/14 bandpass filter. Transfected cells were detected independently
by EGFP fluorescence, as described above. Histograms were constructed
based on analysis of 0.5 × 106 cells.
Chemiluminescence Assay of NADPH Oxidase
Activity--
Superoxide production by untransfected PLB-985 cells was
assayed by chemiluminescence using a superoxide-specific, enhanced luminol-based substrate (DIOGENES, National Diagnostics), as described previously (34). The reactions were monitored for 15 min at 37 °C
following stimulation with 100 ng/ml PMA, using a Luminoskan plate-reading luminometer (Labsystem, Helsinki, Finland). Conditions for dose- and time-dependent inhibition of oxidase activity
were explored with the PLD inhibitors 1-butanol and
2,3-diphosphogylcerate (2,3-DPG) (as well as 3-butanol, control) to
determine the most effective treatment for inhibiting superoxide
generation without reducing cell viability below 98%. Optimal
concentrations were chosen at 0.5% 1-butanol, 0.5% 3-butanol, or 5 mM 2,3-DPG, which were added to the cells 10 min prior to activation.
Translocation and Immunoblot
Analysis--
Bt2cAMP-differentiated PLB-985 cells were
stimulated for 3 min with 1 µM fMLP or for 5 min with 100 ng/ml PMA at 37 °C in Hanks' balanced saline solution with
Ca2+ and Mg2+. Cell membranes were prepared
following sonication by methods described previously (30). 50 µg of
membrane proteins were separated by electrophoresis on 12%
polyacrylamide SDS gels and blotted to nitrocellulose. ARF1 and ARF6
were detected by using rabbit anti-ARF1 (provided by Dr. Richard A. Kahn, Emory University, Atlanta, GA (35)) and rabbit anti-ARF6
(provided by Dr. Julie G. Donaldson, NIH, Bethesda, MD (26)) according
to standard protocols (30).
In previous studies we demonstrated the PLB-985 cell line to be a
useful model system that is capable of developing a differentiated myeloid phenotype while being amenable to gene transfection protocols (30). Because of the relatively low transfection efficiency of these
cells and the requirements for rapid transfection protocols, we
developed a multicolor FACS analysis to identify and characterize small
subpopulations of transiently transfected cells within mixed cell
populations. Transfected cells were identified by detection of
recombinant EGFP, expressed either alone or fused with various ARF
proteins of interest. Following treatments with Bt2cAMP for 3 days to induce a neutrophil-like phenotype, the cells were analyzed for forward angle light scattering (FALS) and 900 side
light scattering properties. Two distinct populations were observed
based on distinct light scattering properties (Fig.
1A): a low FALS population,
indicated as gate G1, and a high FALS population, indicated as gate G2
or G3 (corresponding to untransfected or transfected cell cultures,
respectively). The high FALS population from transfected cell cultures
(G3) exhibited higher 900 light scatter compared with
control untransfected cells (G2), consistent with a greater granularity
that was associated with either the electroporation or sedimentation
protocols employed with these cultures. Fig. 1B (right
panels) shows that the high FALS population from a transfected
culture (G3) contained the majority of EGFP-positive transfected cells,
whereas the low FALS cell population (G1) was not readily transfected,
as indicted by the low EGFP fluorescence readings observed with this
gated population. All subsequent fluorescence-based functional studies focused on the function of the G3 gated population exhibiting the
highest level of EGFP fluorescence (emission wavelength = 512 nm)
as an indication of the highest levels of recombinant ARF expression
(EGFP fluorescence greater than 103).
Flow cytometric methods were also used to monitor superoxide production
by single cells by measuring changes in fluorescence (emitted at 575 nm) upon oxidation of rosamine conjugated to the cell surface. For this
purpose, PLB-985 cells were prelabeled with Ros-SE just prior to
stimulation. Surface-conjugated Ros-SE is oxidized directly by hydrogen
peroxide in activated PLB-985 cells; this oxidation is apparently
dependent on peroxidase activity because reactions that lacked
exogenously added horseradish peroxidase showed no change in Ros-SE
fluorescence (data not shown). As shown in Fig. 1C,
activation of Bt2cAMP-differentiated PLB-985 cells with 1 µM fMLP or with 100 ng/ml PMA caused significant
increases in Ros-SE fluorescence compared with unstimulated cells. In
further support of the notion that Ros-SE oxidation was an indirect
assay of superoxide production, changes in Ros-SE fluorescence observed in cells stimulated by 1 µM fMLP were completely
inhibited by diphenyleneiodonium (8 µM), a known
flavoprotein inhibitor of NADPH oxidase activity (Fig. 1C)
(36). Taken together, these results indicated that the oxidation of
surface-bound Ros-SE reflected NADPH oxidase activation and that the
accumulation of fluorescence under these conditions was not caused by
auto-oxidation of the probe.
Having established compatible assays for both transfected EGFP
expression and oxidase activation in single cells, we explored the
possible involvement of transfected ARF proteins in NADPH oxidase
activation. Bt2cAMP-differentiated PLB-985 cells were transiently transfected with WT forms of recombinant ARF1 or ARF6 produced as fusion proteins with EGFP (ARF1-WT, ARF6-WT) or with fusion
proteins of two mutants of each ARF. ARF6(T27N) and ARF1(T31N) mutants
represent putative dominant-negative mutants with reduced affinity for
GTP, whereas the ARF6(Q67L) and ARF1Q71L) mutants represent putative
active forms with reduced GTPase activity. Earlier work has shown that
the fusion of these proteins at their C terminus with EGFP does not
interfere with GTP-dependent cycling between Golgi membrane
and cytoplasmic compartments (ARF1) or signaling through PLD in
response to receptor stimulation (ARF1, ARF6) in whole transfected
cells (37, 38). We compared activation of NADPH oxidase in
Bt2cAMP-differentiated PLB-985 cells, PLB-985 cells
transfected with empty EGFP-N1 vector (control), and cells transfected
with the various EGFP/ARF protein constructs in response to 1 µM fMLP or 100 ng/ml PMA.
Fig. 2A presents results from
a representative double fluorescence FACS analysis of transfected,
fMLP-activated PLB-985 cells. Cells expressing the highest levels of
recombinant EGFP/ARF proteins in R3 exhibited the most dramatic results
(Figs. 2B and 3). Superoxide production observed in differentiated PLB-985 cells transfected with
ARF1-WT, the two ARF1 mutants, or ARF6-WT was similar to activity
observed in the differentiated, untransfected parental line or
differentiated cells transfected with empty EGFP-N1 vector. In
contrast, cells transfected with ARF6 mutants showed dramatic alterations in oxidase activity. The PLB-985 cells transiently transfected with ARF6(Q67L) showed a significant elevation
(p = 0.03) in oxidase activity, whereas PLB-985 cells
transfected with ARF6(T27N) did not generate any detectable superoxide
in response to fMLP. These effects were only observed when using fMLP
as an agonist; transfected cells stimulated with PMA, a
nonphysiological activator presumed to bypass early receptor-activated
signaling intermediates, showed no effects on oxidative output with the expression of the same recombinant ARF fusion proteins (data not shown). Superoxide production in response to fMLP was similar in all
cell populations that exhibited low EGFP fluorescence (i.e. EGFP Fl < 30, R0) regardless of the ARF construct transfected (data not shown), indicating that all of the cultures analyzed had the
same oxidative potential, and therefore the differences in
fMLP-elicited oxidase activity observed were limited to cells expressing high levels of ARF6 mutants.
A Regulatory Role for ADP-ribosylation Factor 6 (ARF6) in
Activation of the Phagocyte NADPH Oxidase*
,¶
Laboratory of Host Defenses and the
§ Flow Cytometry Section, NIAID, National Institutes of
Health, Bethesda, Maryland 20892
ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
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INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES
s (8), the ARFs have been
shown to play critical roles in vesicular transport (9). Evidence
supporting a role for ARF in granulocyte functions came from studies in
neutrophils and HL-60 cells, where ARF1 and ARF3 were identified as
cytosolic regulators of phospholipase D (PLD) (10, 11). However, these studies were conducted with cell-free reconstitution assays using recombinant ARF1 or cytosol from bovine brain, and therefore the identity of the endogenous ARF(s) participating in PLD activation and
subsequent phagocyte functions is not known. PLD activity and its
product phosphatidic acid have been implicated in a variety of
responses by stimulated phagocytes, including secretion (12, 13),
phagocytosis (14-16), and activation of NADPH oxidase (17). Several
other studies suggest a role for ARF in receptor-dependent signaling in phagocytes (18, 19).
receptor
(FcR)-mediated phagocytosis in the RAW 264.7 macrophage cell line
(29). Phagocytosis of complement-opsonized particles activates PLD in
macrophages. Using Mycobacterium tuberculosis, Kusner
and colleagues (14) correlated inhibition of phagocytosis with
diminished PLD activity and demonstrated that phagocytosis could be
restored by exogenous PLD (14). Finally, studies in neutrophils (15)
and monocytic U937 cells (16) have demonstrated that stimulation of
FcR is tightly linked to PLD activation.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES
RESULTS AND DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES
View larger version (33K):
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Fig. 1.
FACS analysis of PLB-985 cells.
A, representative FALS and side (90o) light
scatter analysis of PLB-985 cells treated for 3 days with
Bt2cAMP to induce development of a neutrophil-like
phenotype. Untransfected PLB-985 cells (left panel) or cells
transfected by electroporation 1 day after induction (right
panel) were subjected to Ficoll gradient sedimentation on day 3 and analyzed as described under "Experimental Procedures." In both
cases two populations (G1 and G2 or G3) were observed which differed in
size and granularity. B, representative EGFP FACS analysis
of untransfected (left panel) and transfected (right
panel) PLB-985 cells. Only the high FALS population of transfected
cells (G3) exhibited significant EGFP fluorescence. The percentage of
transfected cells (indicated in left corners of right
panels) was calculated from EGFP fluorescence cell counts observed
above 30, corrected for background counts observed in untransfected
cells. C, detection of the respiratory burst in
differentiated PLB-985 cells using cell surface-conjugated Ros-SE
oxidase activity (F1/F0) is expressed as
oxidized Ros-SE fluorescence observed in activated cells
(F1), relative to the fluorescence observed without
activation (F0). PLB-985 cells were differentiated,
labeled, and stimulated, as described under "Experimental
Procedures." Inhibition by diphenyleneiodonium (DPI + fMLP) involved
a pretreatment with 8 µM DPI for 5 min prior to the
addition of fMLP. Data represent the means ± S.E.
(n = 4).
View larger version (20K):
[in a new window]
Fig. 2.
Comparison of the respiratory burst activity
in PLB-985 cells transfected with various EGFP/ARF fusion
proteins. A, representative double dot plot obtained
from FACS analysis of respiratory burst activity of EGFP-transfected
PLB-985 cells. R1-R3 represent areas of increasing EGFP fluorescence
within the upper right (double positive) quadrant.
B, fMLP-elicited (1 µM) NADPH oxidase activity
deduced from oxidized Ros-SE fluorescence values
(F1/F0) of transfected cell G3 populations that
exhibited the highest levels of EGFP/ARF protein expression (R3). Data
represent the means ± S.E. (n = 5-7).
Differences between oxidase activity with either ARF6(Q67L) or
ARF6(T27N) and control were statistically significant,
p = 0.03 and p = 0.00,1 respectively.
PLB, untransfected, differentiated PLB-985 cells; EGFP-N1,
differentiated PLB-985 cells transfected with empty EGFP-N1 vector; WT,
wild-type. Various mutant EGFP/-ARF vectors are described under
"Experimental Procedures."
View larger version (16K):
[in a new window]
Fig. 3.
Correlation between mutant ARF6 protein
production and the enhancement (Arf6(Q67L)) or inhibition (ARF6(T27N))
of respiratory burst activity in transfected PLB-985 cells stimulated
with fMLP. Oxidase activities were compared within the three
regions of dot plots illustrated in Fig. 2A. Data represent
the means ± S.E. (n = 4).
To confirm the relationship between ARF6 expression and oxidative output in fMLP-activated cells, we compared superoxide production in cells analyzed within several regions based on EGFP fluorescence intensities. Fig. 3 compares oxidase activity between PLB-985 cells transfected with ARF6 protein constructs (T27N, or Q67L, or WT) and control EGFP-N1 vector. The results extend those obtained in Fig. 2B by showing that transfection with ARF6-WT and EGFP-N1 had no effect on oxidase function in all regions analyzed, while confirming that the alterations in oxidase activity by transfected ARF6(T27N) or ARF6(Q67L) correlated closely with the amount of these proteins produced. A similar analysis of PLB-985 cells transfected with ARF1 protein constructs showed no effect on oxidase activity (data not shown).
To address concerns of whether differences in oxidase activity could be
explained simply by differences in differentiation or expression of
essential oxidase components, we examined the cell surface expression
of gp91phox in these cultures by FACS analysis using a
monoclonal antibody directed against an extracellular epitope of this
protein (33). Fig. 4 shows that all
transfected cell populations detected in R3 expressed comparable
amounts of cell surface gp91phox, as indicated by peak
levels of secondary anti-mouse antibody detected. These results showed
that the alterations in oxidase activity observed with ARF6(T27N) or
ARF6(Q67L) expression were not caused by differences in
gp91phox expression and confirmed that all transfected
cultures differentiated to a comparable extent, consistent with
findings that demonstrated comparable oxidative output in all
populations exhibiting low EGFP fluorescence. These findings provide
further support to the conclusion that ARF6, but not ARF1, has a
specific signaling role in fMLP-mediated activation of the respiratory
burst.
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fMLP-mediated oxidase activation in neutrophils and HL-60 cells is
inhibited by primary alcohols (39), suggesting that fMLP-stimulated oxidase activation through ARF6 is also PLD-dependent in
PLB-985 cells. To test this hypothesis, we used inhibitors of PLD,
1-butanol (39) and 2,3-DPG (14, 40), substrates that inhibit formation of phosphatidic acid, and compared fMLP-mediated activation of NADPH
oxidase in Bt2cAMP-differentiated PLB-985 cells transfected with ARF6-WT, ARF6(Q67L), or empty EGFP-N1 vector (Fig.
5). The PLD inhibitors 1-butanol (0.5%)
and 2,3-DPG (5 mM) blocked superoxide production in PLB-985
cells transfected with empty EGFP-N1 or ARF6-WT. Furthermore, both
inhibitors significantly blocked the enhanced superoxide production
observed in ARF6(Q67L) transfected cells (2,3-DPG versus
control, p = 0.007; 1-butanol versus
control, p = 0.001), whereas 3-butanol (0.5%), which
is not a substrate for PLD, had little effect on the oxidative response
of these cells.
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To explore further the notion that fMLP receptor-mediated activation of
the respiratory burst involves ARF6 signaling through activation of
PLD, we also examined mutant forms of ARF1 and ARF6 which are thought
to be defective in their activation of PLD. A recent report (13)
identified regions within the crystallographic structure of ARF1
important for hPLD1 activation. The substitution of asparagine to
arginine at position 52 (N52R) completely abolished the ability of ARF1
to activate hPLD1 in vitro, while not affecting another
ARF1-associated response, the recruitment of coatamer to membranes.
Based on this observation, we mutated asparagine 52 to arginine (N52R)
in ARF1/EGFP, as well as the corresponding site within ARF6(N48R)/EGFP.
As shown in Fig. 6, PLB-985 cells transfected with ARF1-WT or ARF1(N52R), as well as ARF6-WT, showed no
effect on superoxide production, whereas cells transfected with
ARF6(N48R) exhibited significant inhibition (p = 0.01)
of superoxide production. Although the effects of this mutation on PLD
activation were not examined directly, these findings provide additional support to the notion that ARF6 participates in
receptor-mediated oxidase activation and suggest that ARF6 acts through
activation of PLD, consistent with results obtained with the
pharmacological agents shown in Fig. 5.
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Phorbol esters are also effective stimuli of PLD activity in a wide
range of intact cells (41). In neutrophils and HL-60 cells, most
agonists that activate the respiratory burst also activate protein
kinase C and PLD (39). To clarify further whether protein kinase
C-dependent stimulation of the oxidase involves PLD in
Bt2cAMP-differentiated PLB-985 cells, we tested the effect of the same PLD inhibitors on PMA-elicited superoxide production in
these cells (Fig. 7). PLD inhibitors,
0.5% 1-butanol or 5 mM 2,3-DPG, caused significant
inhibition (p = 0.005 and p = 0.001, respectively) of superoxide production in response to PMA, whereas the
control compound 3-butanol (0.5%) had no effect on superoxide. These
observations, together with the absence of any demonstrable effects of
dominant negative mutants of ARF1 or ARF6 on PMA-stimulated oxidase
activity, suggest that protein kinase C activation of the oxidase is
PLD-dependent but ARF-independent.
|
As another correlate to ARF involvement in oxidase activation in
differentiated PLB-985 cells, we examined membrane translocation of
ARF1 and ARF6 following stimulation with either 1 µM fMLP
or 100 ng/ml PMA. Fig. 8 shows that
stimulation by either of these agonists caused enhanced membrane
binding of both the ARF1 and ARF6 isoforms. These observations are
consistent with previous reports demonstrating that both fMLP and PMA
stimulate translocation of ARF to the plasma membrane in HL-60 cells
and neutrophils, although these studies did not distinguish between the
two isoforms (18, 19). Thus, PMA stimulates ARF1 and ARF6 activation
and translocation to membranes in PLB-985 cells but has other direct or
overriding effects on oxidase activation which appear to involve PLD
but are insensitive to the effects of the dominant ARF1 or ARF6
isoforms.
|
The present study provides novel evidence for unique involvement of ARF6 in N-formyl peptide receptor-stimulated activation of NADPH oxidase because expression of the GTP-binding deficient mutant (ARF6(T27N)) inhibited superoxide production, whereas the GTPase-deficient mutant (ARF6(Q67L)) enhanced superoxide production. Furthermore, the effects of these ARF6 mutants on superoxide production were dose-dependent. In contrast, these dominant-negative and positive effects on superoxide production were not observed when the corresponding ARF1 proteins were expressed, indicating that this response is specific for ARF6. Furthermore, the inhibitory effects of the ARF6(N48R) mutant, as well as those of the PLD inhibitors 1-butanol and 2,3-DPG, provide support for a model in which ARF6 regulates the fMLP receptor-activated respiratory burst through a PLD-dependent mechanism. This model is consistent with longstanding observations correlating neutrophil receptor-mediated oxidative responses to elevations in phosphatidic acid levels (17, 39). Recent reconstitution studies in HL60 cells showed that ARF1 activates hPLD1 and that both ARF1 and hPLD1 are involved in secretion of lysosomal granules (13). These observations, together with our findings, indicate that both ARF1 and ARF6 become activated in stimulated myeloid cells but that the two proteins are involved in different functions which may relate to their segregation into different cellular compartments (23).
In conclusion, we have developed a unique assay using molecular
approaches to study the signaling cascade leading from
N-formyl peptide receptor stimulation to NADPH oxidase
activation in intact neutrophil-like cells. Using this assay we
demonstrated that ARF6 has a direct role in NADPH oxidase regulation
and suggest that this physiological function of ARF6 is mediated
through PLD. ARF6 and protein kinase C appear to activate the oxidase
in parallel pathways. PLD apparently participates in both pathways and
is located downstream of either protein kinase C or ARF6. Future work
should address the identity of the PLD isozyme that links ARF6
activation to the oxidase, as well as upstream signaling intermediates
responsible for ARF6 activation.
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ACKNOWLEDGEMENTS |
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We are grateful to Dr. G. Romero for providing the EGFP/ARF constructs and Dr. J. M. Donaldson, Dr. R. A. Kahn, and Dr. M. Nakamura for providing anti-ARF6, anti-ARF1, and anti-gp91phox, respectively. We also thank S. J. Chanock for providing the PLB-985 cells and D. Stephany for skillful support in the FACS analysis.
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FOOTNOTES |
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* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
¶ To whom correspondence should be addressed: National Institutes of Health, Bldg. 10, Rm. 11N106, Bethesda, MD 20892. Tel.: 301-402-5120; Fax: 301-402-4369; E-mail: tleto@niaid.nih.gov.
Published, JBC Papers in Press, August 7, 2000, DOI 10.1074/jbc.M005406200
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ABBREVIATIONS |
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The abbreviations used are: ARF, ADP-ribosylation factor; PLD, phospholipase D; Bt2cAMP, dibutyryl cAMP; EGFP, enhanced green fluorescent protein; fMLP, formyl-methionine-leucine-phenylalanine; PMA, phorbol 12-myristate 13-acetate; PBS, phosphate-buffered saline; WT, wild-type; Ros-SE, 4-carboxydihydrotetramethylrosamine succinimidyl ester; R1-R3, regions 1-3; G1-G3, gates 1-3; FACS, fluorescence-activated cell sorting; FALS, forward angle light scattering; 2, 3-DPG, 2,3-diphosphoglycerate.
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REFERENCES |
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TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES
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| 1. | Leto, T. L. (1999) in Inflammation: Basic Principles and Clinical Correlates (Gallin, J. I. , and Snyderman, R., eds) , pp. 769-786, Lippincott Williams and Wilkins, Philadelphia |
| 2. | Bokoch, G. M. (1995) Blood 86, 1649-1660 |
| 3. | Abo, A., Pick, E., Hall, A., Totty, N., Teahan, C. G., and Segal, A. W. (1991) Nature 353, 668-670 |
| 4. | Knaus, U. G., Heyworth, P. G., Evans, T., Curnutte, J. T., and Bokoch, G. M. (1993) Science 254, 1512-1515 |
| 5. | Quinn, M. T., Parkos, C. A., Walker, L., Orkin, S. H., Dinauer, M. C., and Jesaitis, A. J. (1989) Nature 342, 198-200 |
| 6. | Maly, F. E., Quilliam, L. A., Dorseuil, O., Der, C. J., and Bokoch, G. M. (1994) J. Biol. Chem. 269, 18743-18746 |
| 7. | Moss, J., and Vaughan, M. (1998) J. Biol. Chem. 273, 21431-21434 |
| 8. | Kahn, R. A., and Gilman, A. G. (1986) J. Biol. Chem. 261, 7906-7911 |
| 9. | Donaldson, J. G., and Klausner, R. D. (1994) Curr. Opin. Cell Biol. 6, 527-532 |
| 10. | Brown, A. H., Gutowski, S., Moomaw, C. R., Slaughter, C., and Sternweis, P. C. (1993) Cell 75, 1137-1144 |
| 11. | Cockcroft, S., Geraint, T. M. H., Fensome, A., Geny, B., Cunningham, E., Gout, I., Hiles, I., Totty, N. F., Truong, O., and Hsuan, J. J. (1994) Science 263, 523-526 |
| 12. | Fensome, A., Cunningham, E., Prosser, S., Tan, S. K., Swigart, P., Thomas, G., Hsuan, J., and Cockcroft, S. (1996) Curr. Biol. 6, 730-738 |
| 13. | Jones, D. H., Bax, B., Fensome, A., and Cockcroft, S. (1999) Biochem. J. 341, 185-192 |
| 14. | Kusner, D. J., Hall, C. F., and Schlesinger, L. S. (1996) J. Exp. Med. 184, 585-595 |
| 15. | Serrander, L., Fallman, M., and Stendahl, O. (1996) Inflammation 20, 439-450 |
| 16. | Melendez, A., Floto, R. A., Gillooly, D. J., Harnett, M. M., and Allen, J. M. (1998) J. Biol. Chem. 273, 9393-9402 |
| 17. | McPhail, L. C., Qualliotine-Mann, D., Agwu, D. E., and McCall, C. E. (1993) Eur. J. Haematol. 51, 294-300 |
| 18. | Houle, M. G., Kahn, R. A., Naccache, P. H., and Bourgoin, S. (1995) J. Biol. Chem. 270, 22795-22800 |
| 19. | Fensome, A., Whatmore, J., Morgan, C., Jones, D., and Cockcroft, S. (1998) J. Biol. Chem. 273, 13157-13164 |
| 20. | Tsuchiya, M., Price, S. R., Tsai, S.-C., Moss, J., and Vaughan, M. (1991) J. Biol. Chem. 266, 2772-2777 |
| 21. | Massenburg, D., Han, J.-S., Liyanage, M., Patton, W. A., Rhee, S. G., Moss, J., and Vaughan, M. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 11718-11722 |
| 22. | D'Souza-Schorey, C., Li, G., Colombo, M. I., and Stahl, P. D. (1995) Science 267, 1175-1178 |
| 23. | Peters, P. J., Hsu, V. W., Ooi, C. E., Finazzi, D., Teal, S. B., Oorschot, V., Donaldson, J. G., and Klausner, R. D. (1995) J. Cell Biol. 128, 1003-1017 |
| 24. | Radhakrishna, H., Klausner, R. D., and Donaldson, J. G. (1996) J. Cell Biol. 134, 935-947 |
| 25. | D'Souza-Schorey, C., Boshans, R. L., McDonough, M., Stahl, P. D., and Aelst, L. V. (1997) EMBO J. 16, 5445-5454 |
| 26. | Song, J., Khachikian, Z., Radhakrishna, H., and Donaldson, J. G. (1998) J. Cell Sci. 111, 2257-2267 |
| 27. | Toda, K., Nogami, M., Murakami, K., Kanaho, Y., and Nakayama, K. (1999) FEBS Lett. 442, 221-225 |
| 28. | Caumont, A.-S., Galas, M.-C., Vitale, N., Aunis, D., and Bader, M.-F. (1998) J. Biol. Chem. 273, 1373-1379 |
| 29. | Zhang, Q., Cox, D., Tseng, C.-C., Donaldson, J. G., and Greenberg, S. (1998) J. Biol. Chem. 273, 19977-19981 |
| 30. | Dana, R., Leto, T. L., Malech, H. L., and Levy, R. (1998) J. Biol. Chem. 273, 441-445 |
| 31. | Shome, K., Nie, Y., and Romero, G. (1998) J. Biol. Chem. 273, 30836-30841 |
| 32. | Model, M. A., KuKuruga, M. A., and Todd, R. F., III (1997) J. Immunol. Methods 202, 105-111 |
| 33. | Kuribayashi, F., Kumatori, A., Suzuki, S., Nakamura, M., Matsumoto, T., and Tsuji, Y. (1995) Biochem. Biophys. Res. Commun. 209, 146-152 |
| 34. | de Mendez, I., and Leto, T. L. (1995) Blood 85, 1104-1110 |
| 35. | Cavenagh, M. M., Whitney, J. A., Carroll, K., Zhang, C-J., Boman, A. L., Rosenwald, A. G., Mellman, I., and Kahn, R. A. (1996) J. Biol. Chem. 271, 21767-21774 |
| 36. | Cross, A. R., Henderson, L., Jones, O. T., Delpiano, M. A., Hentschel, J., and Acker, H. (1990) Biochem. J. 272, 743-747 |
| 37. | Vasudevan, C., Han, W., Tan, Y., Nie, Y., Li, D., Shome, K., Watkins, S. C., Levitan, E. S., and Romero, G. (1999) J. Cell Sci. 111, 1277-1285 |
| 38. | Shome, K., Rizzo, M. A., Vasudevan, C., Andresen, B., and Romero, G. (2000) Endocrinology 141, 2200-2208 |
| 39. | Olson, S. C., and Lambeth, J. D. (1996) Chem. Phys. Lipids 80, 3-19 |
| 40. | Kanaho, Y., Nakai, Y., Katoh, M., and Nozawa, Y. (1993) J. Biol. Chem. 268, 12492-12497 |
| 41. | Exton, J. H. (1999) Biochim. Biophys. Acta 1439, 121-133 |
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