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J. Biol. Chem., Vol. 277, Issue 51, 49965-49975, December 20, 2002
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From the a Department of Environmental and Occupational Health, j Department of Pharmacology, and k Pittsburgh Cancer Institute, University of Pittsburgh, Pittsburgh, Pennsylvania 15260, the b Magee-Womens Research Institute, Pittsburgh, Pennsylvania 15213, the g Institute of Evolutionary Physiology and Biochemistry, Russian Academy of Sciences, St. Petersburg 194223, Russia, the d Department of Medical Chemistry and Biochemistry, Palacký University, 77126 Olomouc, Czech Republic, the e Department of Cell Biology, Physiology, and Immunology, University of Córdoba, 14071 Córdoba, Spain, and the h Institute of Environmental Medicine, Karolinska Institutet, 17177 Stockholm, Sweden
Received for publication, May 8, 2002, and in revised form, September 27, 2002
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ABSTRACT |
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 TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Resolution of inflammation requires clearance of
activated neutrophils by phagocytes in a manner that protects adjacent
tissues from injury. Mechanisms governing apoptosis and clearance of
activated neutrophils from inflamed areas are still poorly understood.
We used dimethylsulfoxide-differentiated HL-60 cells showing inducible oxidase activity to study NADPH oxidase-induced apoptosis pathways typical of neutrophils. Activation of the NADPH oxidase by phorbol myristate acetate caused oxidative stress as shown by production of
superoxide and hydrogen peroxide, depletion of intracellular glutathione, and peroxidation of all three major classes of membrane phospholipids, phosphatidylcholine, phosphatidylethanolamine, and
phosphatidylserine. In addition, phorbol myristate acetate stimulation
of the NADPH oxidase caused apoptosis, as evidenced by
apoptosis-specific phosphatidylserine externalization, increased caspase-3 activity, chromatin condensation, and nuclear fragmentation. Furthermore, phorbol myristate acetate stimulation of the NADPH oxidase
caused recognition and ingestion of dimethylsulfoxide-differentiated HL-60 cells by J774A.1 macrophages. To reveal the apoptosis-related component of oxidative stress in the phorbol myristate acetate-induced response, we pretreated cells with a pancaspase inhibitor,
benzyloxycarbonyl-Val-Ala-Asp-fluoromethyl ketone (z-VAD-fmk), and
found that it caused partial inhibition of hydrogen peroxide formation
as well as selective protection of only phosphatidylserine, whereas
more abundant phospholipids, phosphatidylcholine and
phosphatidylethanolamine, were oxidized to the same extent in the
absence or presence of z-VAD-fmk. In contrast, inhibitors of NADPH
oxidase activity, diphenylene iodonium and staurosporine, as well as
antioxidant enzymes, superoxide dismutase/catalase, completely
protected all phospholipids against peroxidation, inhibited expression
of apoptotic biomarkers and externalization of phosphatidylserine, and
reduced phagocytosis of differentiated HL-60 cells by J774A.1
macrophages. Similarly, zymosan-induced activation of the NADPH oxidase
resulted in the production of superoxide and oxidation of different
classes of phospholipids of which only phosphatidylserine was protected
by z-VAD-fmk. Accordingly, zymosan caused apoptosis in differentiated HL-60 cells, as evidenced by caspase-3 activation and
phosphatidylserine externalization. Finally, zymosan triggered
caspase-3 activation and extensive SOD/catalase-inhibitable
phosphatidylserine exposure in human neutrophils. Overall, our results
indicate that NADPH oxidase-induced oxidative stress in neutrophil-like
cells triggers apoptosis and subsequent recognition and removal of
these cells through pathways dependent on oxidation and externalization
of phosphatidylserine.
Neutrophils aid host defense by killing invading microorganisms
through production of highly reactive oxygen species
(ROS)1 generated by
activation of the NADPH oxidase complex. When released inappropriately
into the extracellular milieu, these ROS can cause persistent
inflammation and considerable damage to the surrounding, healthy
tissues. To prevent calamitous release of ROS, macrophages remove
excess activated neutrophils from an inflammatory site in a regulated
way, through processes that ensure swift resolution of inflammation yet
make provision for neutrophils to fulfill their microbicidal function.
Phagocytic cells carry out this clearance by recognizing apoptotic
neutrophils through a mechanism that involves the exposure of
phosphatidylserine (PS) on the neutrophil cell surface (1-6).
Neutrophils are short lived; in the absence of inflammation, resting
neutrophils undergo apoptosis in the circulation after 6-9 h (7).
Conversely, when neutrophils reach a site of inflammation, apoptosis is
delayed by inflammatory cytokines in the tissues, providing additional
time for completion of the neutrophil's microbicidal function (8).
However, neutrophils also become activated when they migrate from the
blood; their subsequent production of ROS generates oxidative stress
that instigates apoptosis (9, 10). Redox-dependent
mechanisms for apoptosis include the intracellular production of
superoxide (O Although the involvement of the NADPH oxidase in neutrophil apoptosis
has been demonstrated (9, 10), specific signaling pathways through
which oxidative stress participates in recognition and clearance of
apoptotic neutrophils have not been elucidated. We have previously
shown that specific oxidation and externalization of PS was
characteristic of oxidant-induced apoptosis in several different cell
lines (12). We further hypothesized that NADPH oxidase-induced
oxidative stress plays a specific role in recognition and clearance, a
role realized through selective oxidation of PS, associated with PS
externalization on the neutrophilic cell surface, and subsequent
recognition of apoptotic cells by macrophages. In the present work, we
used Me2SO-differentiated HL-60 cells as a model to
study NADPH oxidase-induced apoptosis pathways typical of neutrophils.
In a separate series of experiments, we also used human neutrophils to
corroborate our findings.
Me2SO-differentiated HL-60 cells, in contrast to their
nondifferentiated parental cells, possess a complete NADPH oxidase system (13) that can be activated by various agents (phorbol esters,
chemoattractant peptides, and phagocytosable particles such as
opsonized zymosan, calcium ionophores, etc.). Our results suggest the
following sequence of events in the apoptotic execution program of
Me2SO-differentiated HL-60 cells stimulated with phorbol 12-myristate 13-acetate (PMA) or opsonized zymosan: (i) activation of
NADPH oxidase-dependent O Chemicals
Fetal bovine serum (FBS), Me2SO, PMA, SOD, catalase,
DPI, staurosporine, cytochrome c from horse heart, GSH,
Hoechst 33342, proteinase K, Tris-acetate-EDTA buffer, RNase A,
3-amino-1,2,4-triazole, guaiacol, fluorescamine, and zymosan were
purchased from Sigma. HPLC solvents (methanol, chloroform, hexane, and
water) were obtained from Aldrich. Cetyltrimethylammonium bromide was
purchased from Acros Organics (Pittsburgh, PA).
7-Amino-4-methylcoumarin and acetyl-Asp-Glu-Val-Asp-amino-4-methylcoumarin were purchased
from Peptides International (Louisville, KY). ThioGlo-1TM maleimide reagent was from Covalent Associates Inc. (Woburn, MA).
2-Methyl-6-(4-methoxyphenyl)-3,7-dihydroimidazol[1,2-a]pyrazin-3-one, hydrochloride (MCLA); Amplex Red hydrogen peroxide assay kit; and
cis-parinaric acid (PnA) (Z-9, E-11, E-13,
Z-15-octadecatetraenoic acid) dihydrorhodomine 123 were purchased
from Molecular Probes Inc. (Eugene, OR). Pancaspase inhibitor,
z-VAD-fmk were purchased from Calbiochem, La Jolla, CA. Dulbecco's
modified Eagle's medium and RPMI 1640 medium, agarose, and 100-bp DNA
ladder standards were obtained from Invitrogen Invitrogen (Carlsbad,
CA). The purity of PnA was determined by UV spectrophotometry at
304 nm in ethanol ( Cell Cultures and Treatments
HL-60 human promyelocytic leukemia cells were maintained in RPMI
1640 medium supplemented with 12.5% heat-inactivated FBS at 37 °C
and in a humidified atmosphere (5% CO2 plus 95% air). Cells were seeded at a density of 5 × 105/ml and
grown for 6 days in the presence of 1.25% Me2SO to induce differentiation to the neutrophilic lineage. Fresh medium with Me2SO was added on the third day of culture to prevent cell
overgrowth and depletion of nutrients (14). Several different criteria were utilized to characterize the effectiveness of differentiation (15): (i) an increase of NADPH oxidase activity from <0.05 in nondifferentiated HL-60 cells to 1.25 nmol of
superoxide/min/106 cells in Me2SO-treated
PMA-stimulated HL-60 cells; (ii) a 12-fold decrease of MPO activity
(from 55.25 ± 9.16 to 4.62 ± 1.30 nmol of guaiacol
reduced/min/106 cells before and after Me2SO
treatment, respectively); (iii) the fact that over 95% of
Me2SO-treated cells were viable and had a significantly
smaller size than parental nondifferentiated cells (14); and (iv)
FACScan analysis to determine the number of
Me2SO-differentiated HL-60 cells after stimulation with PMA utilizing dihydrorhodamine 123 staining for the NADPH oxidase. After
PMA stimulation, 97 ± 1% of Me2SO-differentiated
HL-60 cells were responsive to dihydrorhodamine 123 (5 µg/ml)
staining, whereas less than 1% of nondifferentiated cells appeared to
be dihydrorhodamine 123-positive (16). After 6 days, cells were
collected by centrifugation at 1000 × g for 5 min, washed in
prewarmed PBS buffer (pH 7.4), and resuspended in PBS buffer containing
0.5 mM CaCl2, 1 mM
MgCl2, and 30 mM glucose (PBS+). Cells were
either immediately used for assays or kept on ice for no longer than
3 h. For some experiments (those involving assays for GSH,
caspase-3 activity, aminophospholipid externalization, annexin V
binding assay, nuclear morphology, and DNA laddering), cells were
washed and resuspended in FBS-free RPMI 1640 medium without phenol red.
After treatment of cells (2 × 106 cells/ml) with PMA,
an equal volume of phenol red-free RPMI 1640 medium supplemented with
25% FBS (to yield a final concentration of 12.5% FBS) was added to
cells, and incubation was extended for another 2 or 4 h. At the
end of this period, cells were recovered and washed once with
serum-free RPMI 1640 medium. When tested, inhibitors (20 µM DPI, 0.1 µg/ml staurosporine, 50 units/ml SOD plus
50 units/ml catalase (1000 units/ml for neutrophils), and 50 µM z-VAD-fmk) were included in the incubation mixture
5-30 min prior to PMA treatment.
Both Me2SO-differentiated HL-60 cells and neutrophils were
stimulated by 50 µg/ml zymosan (opsonized in human or autologous serum, for neutrophil experiments, at 37 °C for 20-30 min) in RPMI
medium for 2-6 h. Neutrophils were also stimulated with 0.1 µg/ml
PMA. In all experiments, cell viability after treatments was greater
than 90%, as assayed by the trypan blue dye exclusion method.
Macrophages (J774A.1; ATCC cell line) were grown in Dulbecco's
modified Eagle's medium supplemented with 10% heat-inactivated FBS,
100 units/ml penicillin, 100 µg/ml streptomycin, and 50 µg/ml gentamycin sulfate and incubated in a humidified atmosphere (5% CO2 plus 95% air) at 37 °C.
Neutrophils were isolated from blood obtained from adult blood donors
as described by Fadeel et al. (17). Cells (1.0 × 106 cells/ml) were maintained in RPMI 1640 medium
supplemented with 10% FBS and penicillin plus streptomycin. Following
PMA treatment, neutrophils were recovered by trypsinization according
to standard procedures; for zymosan-treated cells, no trypsinization
was necessary.
Quantification of NADPH Oxidase Activity
After stimulation of cells with PMA, NADPH oxidase
activity was determined as the superoxide (O MCLA-enhanced
Chemiluminescence--
Me2SO-differentiated and naive
HL-60 cells (2 × 106 cells/ml) in prewarmed PBS+ were
monitored continuously for 1 min in Luminescent Analyzer 633 (Coral
Biomedical Inc., San Diego, CA), set at 37 °C and continuous mixing,
in the presence of 4 µM MCLA. After 1 min, a stimulant
(0.125 µg/ml PMA or 50 µg/ml zymosan) was added by automated
injection, and continuous readings were taken for another 10-30 min.
Assays were performed in the absence and in the presence of various
inhibitors (SOD/catalase, z-VAD-fmk, staurosporine, or DPI) added 5-30
min prior to the addition of MCLA. Total O Cytochrome c Reduction-coupled Reaction--
Cells, at a density
of 2 × 106 cells/ml of prewarmed PBS+ (1-ml final
volume) containing 100 µM cytochrome c, were
incubated at 37 °C in the absence and in the presence of 50 units/ml
SOD. At 5-min intervals after stimulation of cells with 0.125 µg/ml PMA, 50-µl aliquots were taken; reactions were stopped by the addition of an excess of SOD, and aliquots were mixed on a vortex and
put on ice. Cells were pelleted, and clear supernatants were monitored
in a SpectroMate microspectrophotometer (World Precision Instruments
Inc., Sarasota, FL). Superoxide-dependent reduction of
cytochrome c was calculated by subtracting the absorbance
values at 550 nm ( Assay of Extracellular H2O2
H2O2 levels were determined using Amplex
Red (10-acetyl-3,7-dihydroxyphenoxazine) reagent, the oxidation of
which yields a fluorescent product in the presence of hydrogen peroxide
and horseradish peroxidase (18). Briefly, Amplex Red (50 µM) and 1 unit/ml horseradish peroxidase were added to
cells (105/100 µl) incubated in PBS; the reaction was
initiated by PMA in the presence or absence of DPI, SOD/catalase, or
z-VAD-fmk for 30 min at 37 °C. After incubation, the cells were
centrifuged (1000 × g for 5 min), and the measurements
were carried out in the supernatants at 530/590 nm
( Amplex Red assay of H2O2 determines only
extracellular H2O2. The source of this
extracellular H2O2 could be (i) extracellular nonenzymatic dismutation of superoxide radicals generated by PMA stimulation and (ii) intracellularly produced
H2O2 (in particular, generated by disrupted
mitochondrial electron transport in the course of apoptosis). This
intracellularly produced H2O2 could be
catalytically decomposed, at least in part, before it was excreted from
cells into extracellular environments. As a result, part of
intracellularly generated H2O2 was not
detectable in our experiments with the Amplex Red assay. To minimize
potential effect of adventitious transition metals on the results of
the assay, our measurements of H2O2 in cell
suspensions in PBS were performed after pretreatment of the buffer with
a chelating resin (Chelex 100 resin; Bio-Rad).
Determination of Intracellular GSH and Protein Sulfhydryl
Contents
Glutathione content in the cells was determined
fluorometrically using ThioGlo-1TM as previously described (19).
Briefly, cells treated with PMA and/or a variety of inhibitors
(SOD/catalase, staurosporine, and DPI) were incubated for 30 min,
harvested, resuspended in PBS, and lysed by freezing and thawing once.
Immediately after the addition of 10 µM ThioGlo-1TM to
the cell lysates, fluorescence was measured in a CytoFluor 2350 (Millipore Corp.) fluorescence microplate reader using excitation at
360 ± 40 nm and emission at 530 ± 25 nm. Total protein
sulfhydryls relative to controls were determined as an additional
fluorescence response at the same wavelength 1 h after the
addition of 3.3 mM SDS to the ThioGlo-1TM-treated lysates
kept at room temperature in the dark.
Determination of Phospholipid Peroxidation
PnA was metabolically incorporated into
Me2SO-differentiated HL-60 cell phospholipids (2 × 106 cells/ml) by the addition of PnA-human serum albumin
complex to give a final concentration of 1 µg of PnA/106
cells in FBS-free RPMI 1640 medium without phenol red; cells were
incubated for 2 h at 37 °C. PnA-labeled cells were incubated with 0.125 µg/ml PMA or 50 µg/ml opsonized zymosan in PBS+ buffer for 30 min at 37 °C in the absence and in the presence of various inhibitors including SOD/catalase, DPI, staurosporine, and z-VAD-fmk. At the end of the incubation period, phospholipid oxidation was determined according to previously described methods (20).
High Performance Thin Layer Chromatography (HPTLC) of
Phospholipids
The phospholipid classes in the lipid extract (50 µg of total
phospholipids) were separated by two-dimensional HPTLC on silica G
plates (5 × 5 cm; Whatman) according to methods previously
described (21).
Fluorescamine Labeling of Externalized
Aminophospholipids
Labeling of externalized aminophospholipids, PS and
phosphatidylethanolamine (PE), with fluorescamine (a nonpermeating
probe for visualizing lipids that contain primary amino groups) was carried out by a slight modification of methods previously described by
our laboratory (22). Briefly, HL-60 cells (3 × 107)
treated with PMA from 0.5 to 2.5 h were suspended in labeling buffer (150 mM NaCl, 5 mM KCl, 1 mM
MgCl2, 2 mM CaCl2, 5 mM
NaHCO3, 5 mM glucose, and 20 mM
HEPES; pH 8.0). Cells were gently agitated in the presence of
fluorescamine (200 µM) for 15 s. The reaction was
stopped by the addition of 40 mM Tris-HCl, pH 7.4. Cells
were collected by centrifugation, and lipids were extracted by the Folch procedure (23) and analyzed by HPTLC. Fluorescamine-modified PS
and PE were localized by exposure of HPTLC plates to UV light by using
a Fluor-STM MultiImager (Bio-Rad) imaging system. Unmodified phospholipids were visualized under visible light in a Fluor-STM MultiImager (Bio-Rad) imaging system after exposure of HPTLC
plates to iodine vapor. The phosphorus content of phospholipids was
determined according to Bottcher et al. (24) after scraping
representative spots from the plate. The amounts of modified PS and PE
were expressed as percentages of the total PS and PE (unmodified plus
modified) recovered from the plate on the basis of phosphorus content assay.
Flow Cytometric Analysis of PS Externalization
Annexin V binding to cells was determined using a commercially
available staining kit (Oncogene Research Products, Boston, MA) and
flow cytometry as previously described (22). PMA-stimulated cells were
washed once with PBS. Cells were incubated with fluorescein isothiocyanate-conjugated annexin V (0.5 µg/ml) for 15 min and then
were collected by centrifugation and washed with binding buffer.
Propidium iodide (0.6 µg/ml) was added, and cells were immediately
analyzed with a FACScan flow cytometer (Becton Dickinson, San Jose, CA)
with simultaneous monitoring of green fluorescence (530 nm, 30-nm band
pass filter) for annexin V-fluorescein isothiocyanate and red
fluorescence (long pass emission filter that transmits light >650 nm)
associated with propidium iodide. A time course for PS externalization
was carried out at 0, 0.5, 2.5, and 4.5 h after the addition of
PMA or opsonized zymosan to cells. For neutrophils, PS externalization
was assessed at 3 and 6 h after the addition of PMA and zymosan, respectively.
Determination of Caspase-3 Activity
The activity of caspase-3 was determined as described previously
(25). Briefly, at the indicated times (30 min, 2.5 h, and 4.5 h) after stimulation with PMA or opsonized zymosan, cells were
collected, washed in PBS, and lysed for 20 min on ice in lysis buffer
containing 10 mM HEPES-KOH (pH 7.4), 2 mM EDTA,
0.1% CHAPS, 1 mM phenylmethylsulfonyl fluoride, and 5 mM dithiothreitol. The suspensions were centrifuged at
4 °C, and the supernatants were collected as lysates. For
measurement of caspase activity, 10 µg of lysate diluted to 20 µl
with lysis buffer was mixed with 20 µl of 2× ICE buffer (40 mM HEPES-KOH (pH 7.4), 20% (v/v) glycerol, 1 mM phenylmethylsulfonyl fluoride, and 4 mM
dithiothreitol) containing 40 µM
acetyl-Asp-Glu-Val-Asp-amino-4-methylcoumarin (a fluorogenic peptide substrate) and incubated for 60 min at 37 °C. After 60 min,
460 µl of distilled water was added, and the fluorescence was
measured in a CytoFluor 2350 (Millipore) fluorescence microplate reader
using excitation at 360 ± 40 nm and emission at 460 ± 40 nm. One unit of caspase activity was defined as the amount of enzyme
required to release 1 pmol of 7-amino-4-methylcoumarin/min. The protein
concentration of 10 µg of cell lysates was measured by the method of
Bradford (26). For neutrophil experiments, Asp-Glu-Val-Asp-7-amino-4-methylcoumarin data are presented as pmol of
7-amino-4-methylcoumarin released per 106 cells, as
previously described (17).
Determination of Apoptotic Nuclear Morphology
At specified time intervals, commensurate aliquots of
PMA-stimulated HL-60 cell suspension were taken, and cells were washed and resuspended in PBS. Hoechst 33342 (5 µg/ml) was added, and cells
were examined under fluorescence microscopy. Results were expressed as
the percentage of the cells showing characteristic nuclear
morphological features of apoptosis (nuclear condensation and
fragmentation) relative to the total number of counted cells ( Determination of Apoptosis by DNA Fragmentation
At 30 min and 4.5 h after the addition of PMA, an aliquot
of cells (2 × 106 cells/ml) was taken and washed and
resuspended in 20 µl of lysis buffer (pH 8.0), which contained 10 mM EDTA, 0.5% sarkosyl, and 50 mM Tris. After
incubation for 15 min at 4 °C, proteinase K (20 µg) was added, and
samples were incubated at 50 °C overnight. Samples were centrifuged
at 5800 × g for 10 min. RNase A (500 µg/ml) was then
added to the pellet, and the suspension was incubated at 37 °C for
1 h. Samples were loaded into an agarose gel; a 100-bp DNA ladder
was used as standard. Electrophoresis was run for 2 h at 50 V in
Tris-acetate-EDTA buffer. The gel was then stained with ethidium
bromide (0.5 µg/ml), and the image was analyzed using the Bio-Rad
Multi-Analyst Software.
Phagocytosis of Me2SO-differentiated HL-60 Cells by
J774A.1 Macrophages
Macrophage J774A.1 cells were used for phagocytosis assays.
Before adding target (naive or Me2SO-differentiated HL-60)
cells, macrophages were seeded into an eight-well chamber slide (5 × 104 cells/well) and cultured overnight.
PMA-stimulated cells were washed with serum-free RPMI medium without
phenol red and fluorescently labeled with Hoechst 33342 (1 µg/ml, 10 min at 37 °C) and subsequently washed again (three times) with PBS buffer.
Fluorescently labeled cells (5 × 105 cells/well) were
added to macrophages, and the mixture was incubated for 1 h at
37 °C. After incubation, unbound target cells were washed three
times with RPMI medium and three times with PBS; well contents were fixed with solution of 2% formaldehyde in PBS for 30 min at room temperature. The cells were examined under a Nikon ECLIPSE TE 200 fluorescence microscope (Tokyo, Japan) equipped with a digital Hamamatsu CCD camera (C4742-95-12NBR) and analyzed using the
MetaImaging SeriesTM software version 4.6 (Universal Imaging Corp.,
Downingtown, PA). A minimum of 300 macrophages were analyzed per
experimental condition. Results were expressed as the percentage of the
phagocytosis-positive macrophages.
For the phagocytosis assay, macrophages that had side-by-side
connection with target cells (binding) and/or internalized target cells
(engulfment) were considered phagocytosis-positive. To avoid errors due
to projections in engulfment assessments, we controlled for counted
phagocytosed cells by changing the focusing distance. Use of
fluorescent labeling (Hoechst 33342 for target cells) along with bright
field analysis of typical macrophage and
Me2SO-differentiated HL-60 cell morphologies assured our
counting only the macrophages with bound or engulfed HL-60 cells as
phagocytosis-positive.
Statistical Evaluations
Data are expressed as mean ± S.E. Changes in variables for
different assays were analyzed either by Student's t test
(single comparisons) or by one-way analysis of variance for multiple
comparisons. If any analysis of variance revealed significant changes
among samples, multiple unpaired Student's t tests were
performed. Differences were considered to be significant at
p < 0.05.
Superoxide and Hydrogen Peroxide Production after Activation of the
NADPH Oxidase in Me2SO-differentiated HL 60 Cells
Results of the online assay of MCLA-enhanced chemiluminescence
measuring the rate of O
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
= 80 mM
1
cm1). All other chemicals used were of analytical grade.
= 29.5 mM1
cm1) of samples without SOD from those with SOD.
ex/em) by using a CytoFluor model
2350 fluorescence microplate reader (Millipore Corp., Bedford, MA).
Fluorescence read-outs were converted into H2O2
concentrations (µM) by using a calibration curve and
presented as nmol of H2O2/106 cells.
200
cells/time point).
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
Superoxide production induced by PMA.
A, MCLA-enhanced chemiluminescence was monitored after the
addition of 0.125 µg/ml PMA (arrow) to nondifferentiated
HL-60 cells (1) and to Me2SO-differentiated
HL-60 cells (2-4) in the absence (4) and in the
presence of either 0.1 µg/ml staurosporine (2) or 20 µM DPI (3). Curves are the mean of 3-7
experiments. Bars represent S.D. Values were obtained by
subtracting values in the absence of SOD from those in its presence
(A, inset, representative curves for
PMA-stimulated Me2SO-differentiated HL-60 cells).
B, cytochrome c reduction was measured after the
addition of 0.125 µg/ml PMA in Me2SO-differentiated ( 
)
and nondifferentiated HL-60 cells (
); values represent mean ± S.E.; n = 4-5.
The cytochrome c reduction assay documents the total amount
of O
Amplex Red determination of H2O2 production in
Me2SO-differentiated HL-60 cells showed its significant
stimulation by PMA to yield ~3.6 nmol of
H2O2/106 cells during 30 min of
incubation (after subtraction of the background level) (Fig.
2). Comparison with the data on
O
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NADPH Oxidase-induced GSH Depletion
Activation of the NADPH oxidase by PMA in
Me2SO-differentiated HL-60 cells yielded a statistically
significant (20%) decrease in GSH content (Fig.
3A). This depletion was
completely reversed when NADPH oxidase activation was inhibited by
pretreatment with either DPI or staurosporine. Significant protection
was also afforded by SOD/catalase (~10% GSH-oxidized), implying that
SOD and catalase at these concentrations provide partial protection
against oxidative stress induced by PMA (Fig. 3A). Analysis
of the sulfhydryl groups associated with proteins revealed no changes
under any of the assay conditions tested (Fig. 3B). No
changes in GSH content were observed in nondifferentiated HL-60 cells
treated with PMA, supporting the link between GSH depletion and NADPH
oxidase activation (not shown).
|
NADPH Oxidase-induced Phospholipid Peroxidation without Alteration of Phospholipid Composition in Cells
All three major membrane phospholipids (phosphatidylcholine (PC),
PE, and PS) underwent substantial PMA-induced peroxidation in
Me2SO-differentiated HL-60 cells (Fig.
4). To establish whether oxidation of any
particular class of phospholipids was associated with the execution of
the apoptotic program, we used a pancaspase inhibitor, z-VAD-fmk.
Initially, we determined whether z-VAD-fmk in fact inhibited
PMA-induced caspase-3 activation. To this end, we measured the activity
of caspase-3 and found that it decreased to background level when
Me2SO-differentiated HL-60 cells were stimulated by PMA in
the presence of 50 µM z-VAD-fmk (Fig.
5). Importantly, only one phospholipid,
PS, was protected by z-VAD-fmk against peroxidation (to ~96% of
PnA-PS content in control cells). This indicates that oxidation of PS
was most likely specifically associated with PMA-induced apoptosis,
whereas oxidation of other phospholipids (PC and PE) was probably due
to PMA- dependent activation of the NADPH oxidase and
subsequent nonspecific oxidative stress. Phospholipid oxidation was not
detected when cells were pretreated either with DPI or with
staurosporine at concentrations that inhibited NADPH oxidase activity.
Furthermore, when cells were incubated with SOD/catalase, no
phospholipid oxidation was observed after PMA stimulation. PMA did not
induce any phospholipid oxidation in nondifferentiated HL-60 cells (not
shown).
|
|
It should be noted that PnA-labeled phospholipids represent a small fraction (1-3%) of the total amount of phospholipids present in the cell. As a result, massive oxidation of PnA-labeled PS represented a relatively low level of oxidation of total PS; this was also the case for other phospholipids. Stimulation of Me2SO-differentiated HL-60 cells by PMA did not alter the composition of their phospholipids either relative to control (nonstimulated) cells or relative to time after PMA stimulation (Table I).
|
Externalization of PS Induced by Activation of the NADPH Oxidase
A selective redistribution of PS in plasma membrane from cells
treated with PMA was observed as an increased proportion of fluorescamine-reactive PS on the cell surface, which escalated over
time (Fig. 6A). The concurrent
(nonstimulated) control group did not show any change from the
background level of externalized PS; after 30 min of PMA treatment,
cells exhibited an increase from 2.6 to 11.5% of externalized PS (Fig.
6A). Whereas the level of PS exposed on the cell surface of
PMA-treated cells reached ~16.6% 2 h later, the control group
showed only a slight increase to 6.1% (Fig. 6A). This
demonstrates that PS was externalized and became accessible to
fluorescamine in the extracellular leaflet of plasma membrane after
exposure to PMA. Some amount of PE is exposed on the surface of normal
cells and hence should be available for fluorescamine. Indeed, we found
that ~6.3% of total PE was reactive toward fluorescamine in normal
Me2SO-differentiated HL-60 cells (Fig. 6A,
inset). The amount of fluorescamine-reactive PE was
increased almost 3-fold after 0.5 h of incubation in serum-free RPMI medium as well as 2.5 h of incubation. Most importantly, PMA
stimulation did not significantly affect PE externalization detectable
by this assay as compared with its amounts available for fluorescamine
in nonstimulated cells. Thus, no NADPH oxidase-dependent PE
externalization was detectable in Me2SO-differentiated
HL-60 cells.
|
Although the fluorescamine-based assay allows quantification of the
total amount of PS externalized in the cell suspensions, it does not
provide any information about how these PS molecules are distributed in
the cell population. The use of annexin V, a molecule that specifically
binds to PS, together with flow cytometry permitted us to address this
question. We observed a time-dependent increase in the
proportion of cells with externalized PS (Fig. 6B). At
4 h after PMA stimulation, 70% of cells externalized PS on the
outer leaflet of plasma membrane. Approximately 49% of these
PS-externalizing cells were annexin V+/PI;
51% of the cells were annexin V+/PI+. At an
earlier time point (30 min of PMA stimulation), 40% of cells were
either annexin V+/PI or annexin
V+/PI+. Most of the cells (~62%) with
externalized PS retained an intact membrane (annexin
V+/PI). Pretreatment of cells with DPI or
staurosporine almost completely abrogated PS externalization after
activation of the NADPH oxidase by PMA (Fig. 5C). The
combination of SOD/catalase partially inhibited such externalization of
PS (Fig. 5C). These results clearly support an association
between activation of the NADPH oxidase and PS externalization on the
surface of Me2SO-differentiated HL-60 cells as well as the
involvement of ROS in this process.
NADPH Oxidase-induced Expression of Biomarkers of Apoptosis
To provide evidence for NADPH oxidase-induced apoptosis upon PMA stimulation of Me2SO-differentiated HL-60 cells, several biomarkers of apoptosis were assayed.
Activation of Caspase-3-- After stimulation of the cells with PMA, activation of caspase-3 was observed for as long as 4.5 h (Fig. 5), showing significant activation 2.5 h after PMA exposure (Fig. 5, inset). Furthermore, treatment of cells with SOD/catalase significantly inhibited PMA-induced activation of caspase-3. Expectedly, caspase-3 activation was sensitive to z-VAD-fmk (50 µM). Inhibition of the NADPH oxidase by either DPI or staurosporine almost completely blocked caspase-3 activity (Fig. 5). Nondifferentiated HL-60 cells showed no activation of caspase-3 upon stimulation by PMA (not shown).
Chromatin Condensation and Nuclear
Fragmentation--
Microscopic examination of nuclear morphology
showed that, after exposure to PMA, an increasing percentage of
Me2SO-differentiated HL-60 cells exhibited nuclear
condensation and fragmentation, typical characteristics of apoptosis
(Fig. 7). Inhibition of the NADPH oxidase
by either staurosporine or DPI and scavenging of O
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Activation of the NADPH Oxidase Stimulates Phagocytosis of
Me2SO-differentiated HL-60 Cells by J774A.1
Macrophages--
Exposure of PS on the surface of the plasma membrane
acts as a distinctive signal that allows macrophages to recognize
PS-externalized cells and remove them from surrounding tissues (1-6).
The percentage of macrophages recognizing and/or phagocytizing
PMA-stimulated Me2SO-differentiated HL-60 cells was
~23%, a significant increase when compared with the 10% of
phagocytosis-positive macrophages observed after co-culture with
nonstimulated Me2SO-differentiated HL-60 cells (Fig.
8). When the NADPH oxidase was inhibited
by either DPI or staurosporine, the percentage of recognizing and/or phagocytizing macrophages decreased to values observed when
non-PMA-stimulated Me2SO-differentiated HL-60 cells were
added to macrophage cultures (Fig. 8). Treatment of
Me2SO-differentiated HL-60 cells with SOD/catalase showed a
significant inhibitory effect on phagocytosis (Fig. 8). After exposure
to PMA, nondifferentiated HL-60 cells were not phagocytized by J774A.1
macrophages (not shown).
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Zymosan Stimulation of the NADPH Oxidase and Oxidative Stress in Me2SO-differentiated HL-60 Cells
We further used opsonized zymosan to determine whether similar
PS-dependent responses could be observed in
Me2SO-differentiated HL-60 cells with physiologically more
relevant phagocytosable stimuli. We found that serum-opsonized zymosan
caused activation of caspase-3 in Me2SO-differentiated
HL-60 cells that was completely blocked by a pancaspase inhibitor,
z-VAD-fmk (Fig. 9A). Zymosan induced a pronounced production of superoxide in HL-60 cells (Fig. 9B). Whereas zymosan-induced oxidative burst was ~3-fold
less than that induced by PMA, it was also partially inhibitable by z-VAD-fmk and completely blocked by SOD/catalase as well as by DPI
(data not shown). Again, similar to PMA, opsonized zymosan caused a
significant oxidation of different classes of phospholipids (Fig.
9C). Remarkably, only PS oxidation was significantly
inhibited by z-VAD-fmk (from 31.0 ± 3.9% of PS oxidized in the
absence of z-VAD-fmk to 16.0 ± 3.7% of PS in the presence of
z-VAD-fmk), whereas oxidation of other major phospholipids such as PC
and PE was not significantly changed by the pancaspase inhibitor. We
also observed that zymosan caused PS externalization in
Me2SO-differentiated HL-60 cells (~44% of cells were
annexin V+/PI). PS externalization was
partially suppressed by SOD/catalase and to a lesser extent by
z-VAD-fmk (data not shown). Thus, both PMA and opsonized zymosan caused
oxidative stress and PS oxidation in Me2SO-differentiated
HL-60 cells associated with the execution of the apoptotic program as
revealed by the sensitivity toward z-VAD-fmk.
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PMA- and Zymosan-induced NADPH Oxidase Activation and PS Externalization in Neutrophils
Finally, we performed an additional series of experiments with
human neutrophils stimulated with PMA or serum-opsonized zymosan. We
found that no caspase-3 activation was induced in PMA-treated cells in
line with previous observations (17). Zymosan treatment, on the other
hand, markedly enhanced caspase-3 activation when compared with
constitutive, background levels (Fig.
10A). This effect was not
pronounced at 3 h (data not shown), but was seen clearly at later
time points (6 h). Caspase-3 activation was sensitive to DPI
(p < 0.001) in zymosan-induced neutrophils.
SOD/catalase caused a 32% decrease (which did not reach the level of
significance); as expected, z-VAD-fmk completely blocked the activation
of caspases under these conditions. Furthermore, both PMA and zymosan
triggered extensive PS exposure in neutrophils (Fig. 10B).
PMA-induced PS externalization was significantly inhibited by DPI
(p < 0.001). No effect of z-VAD-fmk on PS exposure was
seen in PMA-treated cells. Zymosan-induced PS externalization was
inhibited by DPI (p < 0.001) as well as by
SOD/catalase (p < 0.05) and was only slightly (by
16%) suppressed by z-VAD-fmk. These data show that zymosan triggers
NADPH oxidase-dependent activation of caspases and PS
externalization in human neutrophils. Importantly, the sensitivity of
zymosan-dependent PS externalization toward DPI and
SOD/catalase demonstrates that apoptosis-specific signaling pathways are engaged as neutrophils encounter a phagocytosable stimulus.
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DISCUSSION |
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TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Neutrophils possess powerful ROS-generating machinery, the NADPH oxidase system, to kill invading microorganisms. Uncontrolled activation of neutrophils in anomalous inflammatory processes promotes different oxidation-associated injuries in several acute conditions, including acute respiratory distress syndrome and reperfusion injury, and in chronic diseases such as emphysema, atherosclerosis, and rheumatoid arthritis (for a review, see Ref. 28). Preeclampsia, a human pregnancy-related disorder, has been also associated with neutrophil activation (29, 30) and with delay of neutrophil apoptosis (31) as sources of the high level of oxidative stress.
Activated neutrophils should be removed from inflamed areas once invading microorganisms have been eliminated and the infection has been resolved. This removal takes place through recognition of apoptotic neutrophils by phagocytes via a pathway that involves exposure of PS on neutrophil cell surfaces (1-6). Yet the mechanisms underlying apoptosis in activated neutrophils, particularly those involving PS externalization, are not fully characterized.
NADPH Oxidase-induced Oxidative Stress and Apoptosis in Me2SO-differentiated HL-60 Cells-- In the present work, we have demonstrated that when the NADPH oxidase was activated by PMA or opsonized zymosan in neutrophil-like Me2SO-differentiated HL-60 cells, apoptosis was increased. Furthermore, the increase was correlated with enhanced phagocytosis by macrophages. Our data further show that activation of the NADPH oxidase complex in neutrophil-like cells triggers the mechanisms necessary for phagocytic clearance of neutrophils from inflamed areas by mechanisms involving oxidative stress.
Several authors have observed no induction of apoptosis in Me2SO-differentiated HL-60 cells after stimulation with PMA and have attributed this observation to the lack of specific granules and, therefore, of intracellular production of H2O2 in the cells (9). Neutrophils, however, release oxidants extracellularly despite ~95% of the NADPH oxidase being intracellular (for a review, see Ref. 32). Although it has been reported that neutrophils become apoptotic through the effects of intracellular H2O2 generated as a secondary product during activation of the granule pool of the NADPH oxidase (9), the mechanisms governing clearance of apoptotic neutrophils still remain elusive.
Here, we have presented contrasting positive evidence that Me2SO-differentiated HL-60 cells undergo apoptosis when the NADPH oxidase is activated by PMA. The major difference between our work and that of Lundqvist and Bengtsson (9) is the culture conditions used for the apoptosis experiments. When we tried to reproduce the serum-free culture conditions used in Lundqvist and Bengtsson's experiments (9), we also noted an increased percentage (~25-30%) of apoptotic cells in the control group after 2-4 h of incubation in PBS+ buffer (data not shown). Therefore, the cells for our experiments were transferred into a serum-containing medium after treatments, in order to avoid added stress to the cells. Thus, we were able to observe differences in apoptosis resulting from activation of the NADPH oxidase (Fig. 7) that would otherwise have been masked by a high control background.
To provide evidence for the involvement of NADPH oxidase-dependent oxidative stress in apoptosis induced by PMA or zymosan in Me2SO-differentiated HL-60 cells and human neutrophils, we used two different inhibitors of the enzyme, DPI and staurosporine, as well as antioxidants, SOD/catalase. DPI, a flavoprotein inhibitor, is commonly used as an inhibitor of the NADPH oxidase (33). Staurosporine exerts multiple effects, and, as a protein kinase C inhibitor, it acts as a potent suppressor of the NADPH oxidase (since protein kinase C is involved in activation of the NADPH oxidase). In fact, we observed that superoxide production was completely blocked by staurosporine in Me2SO-differentiated HL-60 cells stimulated by PMA. These findings concur with the previous observation that staurosporine prevents PMA-induced cytotoxicity in neutrophils (34). Thus, staurosporine is able to prevent NADPH oxidase-dependent oxidative stress and subsequent oxidant-induced apoptosis in Me2SO-differentiated HL-60 cells. This effect of staurosporine was similar to that of DPI. Not surprisingly, as both inhibitors prevented oxidant-induced triggering of apoptosis, they also completely blocked expression of all biomarkers of apoptosis. There are several reports demonstrating that staurosporine is able to induce apoptosis in different cell lines predominantly via mitochondrial permeability transition pathways (35). Further studies are necessary to determine to what extent this propensity of staurosporine may be realized in Me2SO-differentiated HL-60 cells and neutrophils with high levels of NADPH oxidase activity under the experimental conditions of this study.
To facilitate removal of ROS during PMA-induced oxidative stress and
apoptosis, we chose to use a combination of SOD and catalase. Since
both enzymes remain in the extracellular compartments, they can only
eliminate extracellular O
Oxidation of PS and Phospholipid Signaling in Me2SO-differentiated HL-60 Cells-- Here we propose a new role for oxidative stress in phospholipid signaling through oxidation of PS and its further externalization. We observed that all three major membrane phospholipids, PC, PE, and PS, underwent significant PMA-induced peroxidation in Me2SO-differentiated HL-60 cells. In these cells, PMA induces oxidation through two different mechanisms: (i) via stimulation of the NADPH oxidase that generates ROS and, hence, catalyzes oxidation of different cellular constituents, including phospholipids, and (ii) due to triggering and execution of the apoptotic program. One may assume that the former pathway will be associated with nonspecific oxidation of all major classes of phospholipids, whereas the latter may be specific to those involved in oxidative signaling pathways of apoptosis. To differentiate between these two mechanisms, we used a pancaspase inhibitor, z-VAD-fmk, that blocked PMA-induced caspase-3 activity in Me2SO-differentiated HL-60 cells. We found that, indeed, only PS was protected by z-VAD-fmk against PMA-induced oxidation, whereas two more abundant phospholipids, PC and PE, were oxidized to the same extent in the absence or presence of z-VAD-fmk. In contrast, inhibitors of NADPH oxidase activity, DPI and staurosporine, blocked oxidation of all phospholipids to the same degree. Similarly, antioxidant enzymes, SOD/catalase, protected all three phospholipids against peroxidation. Taken together, these results strongly suggest that PS oxidation is likely to represent an apoptosis-specific event. This is further supported by partial inhibition of PMA-induced H2O2 production by z-VAD-fmk in Me2SO-differentiated HL-60 cells.
We further used opsonized zymosan to determine whether similar PS-dependent responses could be observed in Me2SO-differentiated HL-60 cells with physiologically more relevant phagocytosable stimuli. Opsonized zymosan activated the NADPH oxidase and induced apoptosis, as evidenced by superoxide anion production, PS externalization, and caspase-3 activation. Furthermore, we observed the same phospholipid oxidation pattern with zymosan as with PMA. Thus, both PMA and opsonized zymosan caused oxidative stress in Me2SO-differentiated HL-60 cells, part of which was associated with the execution of apoptotic program, as revealed by the sensitivity toward z-VAD-fmk.
Non-oxidant-induced apoptosis may be a preferable model for the elucidation of phospholipid oxidative signaling mechanisms. Therefore, in a separate study, we utilized a model of staurosporine-induced apoptosis in HL-60 cells in which treatment with staurosporine caused marked oxidation of PS while PC remained unoxidized (37). In addition, employing another model of non-oxidant-induced apoptosis (death receptor triggering using agonistic anti-Fas antibodies), we have recently reported production of both superoxide (detected by MCLA chemiluminescence response) as well as H2O2 (detected by Amplex Red assay) in Jurkat T cells (38). Both superoxide generation and accumulation of H2O2 were substantially attenuated by z-VAD-fmk. Most notably, PS was preferentially oxidized upon Fas ligation (38).
Oxidized epitopes on the surface of apoptotic cells are known to be important signals for the recognition of target cells by macrophages (39-40). To verify the importance of oxidized PS in phagocytosis of apoptotic cells, we used nonapoptotic HL-60 cells with PS or with PS plus oxidized PS integrated in the outer surface of their plasma membrane (38). We found that a mixture of PS plus oxidized PS yielded an almost 2-fold enhancement of PS-dependent phagocytosis of HL-60 cells by J774A.1 macrophages. Conversely, pretreatment of macrophages with liposomes containing a mixture of PS plus oxidized PS inhibited phagocytosis of apoptotic HL-60 cells more effectively than pretreatment with liposomes loaded with nonoxidized PS alone.
In the current study, phospholipid peroxidation did not affect overall phospholipid composition in Me2SO-differentiated HL-60 cell membranes. Rather, NADPH oxidase-dependent oxidation of PS altered the usual distribution of PS in resting, nonstimulated cells and resulted in externalization of PS on the cell surface (9-10% more externalized PS than in the control group). The most commonly used assay for assessment of PS externalization, based on the interactions of PS with annexin V on the cell surface, is not absolutely specific. Other negatively charged moieties including peroxidation products of aminophospholipids can bind annexin V as has been recently reported by Balasubramanian et al. (41). It should be kept in mind, however, that these interactions occur at relatively high concentrations of aminophospholipid peroxidation products, hardly achievable in living cells (after exposure of cells to ~5 mM malonyl dialdehyde), including cells executing the apoptotic program. Our PnA-based assay of phospholipid peroxidation indicates that the total concentration of oxidatively modified phospholipids did not exceed 3-5 µM (i.e. was about 3 orders of magnitude lower than that used by Balasubramanian et al. (41) for cell treatments). The amount of oxidatively modified aminophospholipids on the cell surface is likely to be even smaller. Therefore, binding of annexin V to aminophospholipid peroxidation products on the surface of apoptotic cells does not seem to substantially affect measurements of PS externalization.
Oxidation of PS may stimulate externalization of both PS and oxidized PS. There are different models explaining specific mechanisms through which oxidation of PS may facilitate externalization of PS or oxidized PS. According to one such model, oxidized PS poisons the aminophospholipid translocase, the enzymatic activity that is responsible for the maintenance of plasma membrane phospholipid asymmetry (42), in which case both PS and oxidized PS would be expected to appear on the cell surface. Another scenario is that the aminophospholipid translocase fails to recognize oxidized PS, resulting in selective externalization of oxidized PS. In the latter case, inhibition of PS oxidation should result in complete inhibition of PS externalization. In the former case, inhibition of PS oxidation may be consistent with the presence of PS (but not oxidized PS) on the cell surface. However, we do not have any information on the amounts of oxidized PS on the surface of apoptotic cells. Oxidation of phospholipids may also occur in the outer leaflet of the membrane by extracellular oxidants such
),
staurosporine (
), or DPI (
) had no additions. (T
represents the beginning of treatment, and S represents the
addition of serum-containing medium). Values are mean ± S.E.
(n = 3). a, p < 0.01 versus control; b, p < 0.05 versus PMA plus SOD/catalase.
