1 Department of Medicine, Mayo Clinic Rochester, Rochester, MN 55905, USA
2 Department of Immunology, Mayo Clinic Rochester, Rochester, MN 55905,
USA
3 Department of Dermatology, University of Utah, Salt Lake City, UT 84132,
USA
4 Department of Physiology, University of Minnesota, St Paul, MN 55108,
USA
5 Department of Animal Science, University of Minnesota, St Paul, MN 55108,
USA
* Author for correspondence (e-mail: bankers.jennifer{at}mayo.edu)
Accepted 22 April 2003
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Summary |
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Key words: Membrane potential, PKC, superoxide, H+ channel, NADPH oxidase
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Introduction |
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Membrane depolarization after NADPH oxidase activation has been reported in
neutrophils (Seligmann and Gallin,
1980; Whitin et al.,
1980
; Henderson et al.,
1987
; Demaurex et al.,
1993
; Åhlin et al.,
1995
; Susták et al., 1997;
Jankowski and Grinstein, 1999
)
and appears to be produced in part by NADPH oxidase activity itself. Electron
transfer through NADPH oxidase may depolarize the cell, and concomitant proton
efflux attenuates this depolarization. Neutrophils from patients with chronic
granulotomatous disease (CGD), in which NADPH oxidase is not functional, do
not depolarize in response to various stimuli
(Seligmann and Gallin, 1980
;
Cohen et al., 1981
;
Castranova et al., 1981
;
Åhlin et al., 1995
).
Similarly, neutrophils treated with the NADPH oxidase electron transport
inhibitor diphenyleneiodonium (DPI) have attenuated depolarization
(Henderson et al., 1987
;
Jankowski and Grinstein,
1999
). When proton efflux is inhibited during oxidase activation,
eosinophils (Bánfi et al.,
1999
) and neutrophils
(Henderson et al., 1987
;
Suszták et al., 1997
)
depolarize to a substantially greater degree. Likewise, depolarization is
attenuated when NADPH oxidase is activated under conditions that favor
increased proton efflux (Henderson et al.,
1987
). Thus, activation of the electrogenic NADPH oxidase assures
coordination of proton efflux that is required for continued superoxide
production.
Jankowski and Grinstein (Jankowski and
Grinstein, 1999) have confirmed that neutrophils depolarize
sufficiently during activation to promote the efflux of protons through the
NADPH oxidase-associated proton channel. After phorbol ester stimulation,
neutrophils depolarized from a resting membrane potential of 58 mV to
+58.6±6 mV, a magnitude consistent with a physiological role for the
proton conductance during NADPH oxidase activity. Eosinophil membrane
potential has been measured in both resting and activated states. Roberts and
Gallin (Roberts and Gallin,
1985
) used di-O-C5(3), a membrane potential-sensitive
carboxycyanine dye, to document relative eosinophil depolarization after
phorbol 12-myristate 13-acetate (PMA) stimulation; however, when loaded with
the same dye, eosinophils depolarized less than neutrophils. Subsequently,
current clamp recordings on eosinophils suggested an absolute resting membrane
potential of approximately 60 to 80 mV
(Gordienko, 1996
;
Tare, 1998
), and an increase
of 30-40 mV following NADPH oxidase activation with NADPH and GTP
S
(Bánfi et al., 1999
).
Therefore, on the basis of current data describing eosinophil plasma membrane
potential, the NADPH oxidase-associated proton conductance is unlikely to be
activated in eosinophils. This is in contrast to what has been described for
neutrophils and to circumstantial evidence for proton efflux after NADPH
oxidase stimulation in eosinophils.
We have previously reported that PMA-stimulated human eosinophils produce
superoxide and activate the NADPH oxidase-associated proton conductance
(Bankers-Fulbright, 2001).
Here, we describe the effects of PMA on the membrane potential of eosinophils
using a cell-permeable oxonol membrane potential indicator, bis-barbituric
acid oxonol [diBAC4(3)], to determine the extent of depolarization
(Krasznai et al., 1995
). Like
neutrophils, eosinophils depolarize to a membrane potential sufficient to
allow activation of the NADPH oxidase-associated proton channel under
conditions where superoxide is produced. Additionally, protein kinase C
(PKC
appears to be crucial for PMA-induced depolarization.
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Materials and Methods |
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Eosinophil isolation
Peripheral blood human eosinophils were isolated as described
(Hansel et al., 1991;
Ide et al., 1994
). Briefly,
heparinized blood was collected from atopic and nonatopic volunteers, an equal
volume of 1
PIPES was added (25 mM PIPES, 50 mM NaCl, 5 mM KCl, 25 mM
NaOH, 5.4 mM glucose, pH 7.4), and the diluted blood was layered onto Percoll
(density 1.085 g/ml). After centrifugation at 1000 g for 30
minutes at 4°C, the plasma and Percoll layers were removed by aspiration.
Tubes were wiped to remove contaminating leukocytes, and red cells were lysed
by osmotic shock. The remaining pellet, containing neutrophils and
eosinophils, was incubated with an equal volume of anti-CD16 magnetic beads
(Miltenyi Biotec, Auburn, CA) on ice for 30 minutes. After incubation the cell
mixture was diluted with 1
PIPES + 1%
calf serum (
CS)
(HyClone Laboratories, Logan, UT) and eluted through a steel wool column
suspended in a strong magnet. Column eluate (14 ml) was collected and the
number of eosinophils was determined by staining with Randolph's stain.
Eosinophil purity was always greater than 93% and the major contaminating
cells were neutrophils.
Transmembrane potential measurements
Eosinophil transmembrane potential was determined using
diBAC4(3), a ratiometric fluorescent membrane potential-sensitive
dye (Molecular Probes, Eugene, OR) based on a protocol by Krasznai et al.
(Krasznai et al., 1995). Cells
were resuspended at 0.5
106 cells/ml in HybriCare media (no
bicarbonate buffer, with 25 mM HEPES) (ATCC, Rockville, MD) in HSA-blocked
FACS tubes (Falcon 2052, Fisher Scientific, Lincoln Park, NJ). For the
standard curve, cells were resuspended at room temperature with varying
concentrations of diBAC4(3) (5000 nM to 300 nM). Cells to be
stimulated were resuspended with 600 nM diBAC4(3). Additionally,
control cells were fixed for 1 hour on ice with ice-cold 2% formaldehyde,
washed with HybriCare and incubated for 1 hour at room temperature with 600 nM
diBAC4(3). All samples were run on a FACScan flow cytometer (Becton
Dickinson, Lincoln Park, NJ) at room temperature. Cells were stimulated with
800 nM PMA and fluorescence was measured at time 0 and every minute thereafter
for 10 minutes. Inhibitors were incubated with cells at least 10 minutes
before PMA stimulation. A linear calibration curve was determined from the
cells incubated with varying concentrations of diBAC4(3), and this
equation was used to calculate the absolute transmembrane potential of the
cells as described by Krasznai et al.
(Krasznai et al., 1995
).
Confocal microscopy
Eosinophils were resuspended to 0.5106 cells/ml in
HybriCare media with 600 nM diBAC4(3) and loaded into
eight-chambered coverglass slides (Nalge Nunc International Corporation,
Naperville, IL) at room temperature. Cells were stimulated with 800 nM PMA and
examined using a Zeiss LSM 510 Confocal Laser Scanning Microscope (Carl Zeiss,
Oberkochen, Germany). An excitation wavelength of 488 nm was used and emission
was detected using a 505LP filter.
Superoxide assay
Superoxide assays were performed at 37°C in HBSS buffer (Hank's
balanced salt solution) supplemented with 10 mM HEPES. Cytochrome c was used
to detect the production of extracellular superoxide as previously described
(Bankers-Fulbright et al.,
1998). Inhibitors were added immediately before stimulation unless
otherwise noted. Stock solutions of diphenyleneiodonium chloride (DPI),
Gö6976, GF109203X and rottlerin (all from Calbiochem, LaJolla, CA) were
diluted in HBSS so that the final concentration of DMSO or ethanol was less
than 0.5%. The rate of superoxide production was calculated using the linear
part of the superoxide production curve (usually 20 to 50 minutes following
stimulation) and is presented as nanomoles of superoxide produced per
minute.
Intracellular pH measurements
Intracellular pH was measured as described previously
(Boyer and Hedley, 1994), with
the following changes. Purified eosinophils were resuspended to
0.5
106 per ml in physiological saline solution [140 mM NaCl, 5
mM KCl, 5 mM glucose, 1 mM CaCl2, 1 mM MgCl2, 20 mM
2-[N-morpholino]ethanesulfonic acid (MES)] supplemented with 1%
calf serum. Eosinophils were loaded with 5 µM BCECF-AM
[2',7'-bis-(2-carboxyethyl)-5-(and-6)-carboxy-fluorescein
(acetoxymethyl ester derivative)] (Molecular Probes) for 30 minutes at
37°C in the dark, washed once, resuspended in HBSS (with 25 mM PIPES, pH
7.4) and plated at 5
105 eosinophils per well in an
albumin-blocked, 96-well plate (final volume per well=200 µl). Plates were
read at 37°C on a CytoFluor Series 4000 (perSeptive Biosystems,
Framingham, MA) fluorescent plate reader with excitation at 485 nm and 450 nm
and emission at 535 nm. Calibration of the BCECF-AM dye and calculation of
intracellular pH was performed as described earlier
(Boyer and Hedley, 1994
).
Perforated whole-cell patch clamp technique
The amphotericin perforated whole-cell patch configuration was used.
Pipette electrodes were pulled to a resistance of 2-4 M from 7052 glass
(Garner Glass, Claremont, CA). The pipette tip was filled with
KMeSO4 Ringer solution (130 mM potassium MeSO3, 5 mM
NaCl, 1 mM CaCl2, 25 mM (NH4)2SO4,
pH 7.0). The pipette was then backfilled with the same solution containing 10
µM amphotericin B. High resistance seals (>5 G
) were formed
between the pipette and the cell membrane, and amphotericin was allowed to
partition into the membrane to obtain the whole-cell configuration before
currents were recorded. Typical series resistances ranged from 8 to 20
M
. Compensation for series resistance was accomplished using the series
resistance compensation circuitry available with the Axopatch 1D amplifier.
The standard recording solution for patch-clamp experiments consisted of
HybriCare media (ATCC) supplemented with 2 mM glutamine, 50 µg/ml
gentamicin sulfate, 7.5 mM (NH4)2SO4 0.3%
serum albumin and buffered with 10 mM HEPES, pH 7.4. An Axopatch 1D voltage
clamp and Digidata 1322 interface were used (both from Axon Instruments, Union
City, CA). P-CLAMP 8.0 software was used to generate the voltage-step commands
and to record the resulting currents. Analysis of the whole-cell current
traces was performed using AxoGraph 3.0.3 software from Axon Instruments.
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Results |
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|
One consequence of proton channel activation in eosinophils should be
attenuation of NADPH oxidase-dependent depolarization. If the proton channel
is activated, then blocking proton efflux should increase the magnitude of
depolarization. To test this, we stimulated eosinophils with PMA in the
presence of 250 µM ZnCl2; this concentration of ZnCl2
fully blocks PMA-stimulated eosinophil proton currents but does not fully
inhibit PMA-stimulated NADPH oxidase activity
(Bankers-Fulbright, 2001). As
predicted, treatment with ZnCl2 enhanced the magnitude of
eosinophil depolarization (+43.0±4.2 mV; P=0.0006), in
contrast to stimulation with PMA alone (+16.2±1.3 mV)
(Fig. 2).
|
To confirm that the outcome observed with ZnCl2 was due to
effects on proton transport, we altered the proton channel activation
threshold by changing the extracellular pH (pHe). If proton efflux
plays a role in regulating eosinophil depolarization, increasing the
activation threshold (lowering pHe) should increase the magnitude
of depolarization and decreasing the threshold (increasing pHe)
should attenuate depolarization (Cherny et
al., 1995; Åhlin et al.,
1995
; Henderson et al.,
1987
). Notably, the voltage threshold for activation of the proton
conductance depends primarily on the difference between, not the absolute
values of, intracellular and extracellular pH
(Cherny et al., 1995
). Thus, we
first measured eosinophil intracellular pH (pHi) under different
pHe conditions to document this difference
(Fig. 3A). Eosinophils in
standard buffer (pH 7.4) have a resting pHi of 7.12±0.30.
Decreasing pHe to 7.0, 6.5 and 6.0 induces corresponding
significant decreases in pHi (7.00±0.05, 6.71±0.04
and 6.53±0.04, respectively; all P<0.00001 compared with
pHe 7.4). Eosinophils stimulated with PMA for up to 2 hours
consistently maintain their resting pHi at all pHe (data
not shown). Thus, at pHe greater than 7.0, the ratio of
intracelluar to extracellular proton concentration favors efflux; at
pHe less than 7.0, proton efflux is diminished. PMA-stimulated
NADPH oxidase activity is also affected by changing pHe
(Fig. 3B). Interestingly, the
rate of superoxide production is not significantly affected by decreasing
pHe to 7.0 (P=0.48) or 6.5 (P=0.065). However,
the PMA-stimulated NADPH oxidase rate is inhibited by approximately 50% at
pHe 6.0 (P=0.005).
|
To document the effect of pHe on eosinophil depolarization, we analyzed diBAC4(3)-loaded eosinophils at pHe between 6.0 and 8.0. Membrane potential measurements of eosinophils maintained at pHe 6.0 and pHe 8.0 showed some variability, but no significant differences in resting membrane potential (19.8 vs 33.2, respectively; P=0.06) (Fig. 3C). As predicted, eosinophils at pHe 8.0 showed a decreased magnitude of PMA-stimulated depolarization (+8.9±1.4 mV) compared with cells stimulated at pHe 7.4 (+19.2±2.2 mV; P=0.002). Similarly, decreasing pHe to 7.0, 6.5 and 6.0 (reducing proton efflux) increased the magnitude of depolarization to +33.9±2.6 mV (P=0.0001), +44.4±1.4 mV (P=0.004) and +42.2±4.6 mV (P=0.010), respectively. There was no significant difference between the rate (P=0.082) or magnitude (P=0.092) of PMA-stimulated eosinophil depolarization at pHe 6.0 vs 6.5. In addition, these results were not significantly different from the values obtained following zinc inhibition of the proton channel. Thus, eosinophil depolarization is predictably affected by changing proton efflux through modifications of the proton gradient, as well as through blocking the proton channel with ZnCl2.
The NADPH oxidase and its associated proton channel are often activated by
the same stimuli (DeCoursey et al.,
2000; DeCoursey et al.,
2001b
; Cherny et al.,
2001
). We have previously shown that superoxide production by
PMA-stimulated eosinophils is inhibited by the selective PKC
blocker,
rottlerin (Bankers-Fulbright,
2001
). However, proton channel activation in these cells is not
blocked by rottlerin alone, although the pan-specific PKC blocker, GF109203X,
does block proton channel activity. To determine which PKC isoforms were
modulating eosinophil depolarization, we stimulated cells with PMA in the
presence of different PKC inhibitors (Fig.
4). None of the PKC inhibitors significantly affected eosinophil
resting membrane potential. The pan-specific PKC blocker, GF109203X,
completely inhibited PMA-induced depolarization (P=0.00006).
Additionally, the PKC
-selective blocker, rottlerin, significantly
inhibited PMA-stimulated depolarization (P=0.02), and a blocker of
Ca2+ dependent PKC isoforms, Gö6976, had a minimal but
statistically significant (P=0.03) effect.
|
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Discussion |
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Although both neutrophils and eosinophils are granulocytes and use the same
NADPH oxidase to generate superoxide, the amount and location of the oxidase
is different in these cell types. Thus, extrapolating membrane potential
results from recent neutrophil data is not necessarily predictive of the
depolarization magnitude or time course in eosinophils. Eosinophils express
more NADPH oxidase than neutrophils in both humans and guinea pigs
(Yagisawa et al., 1996;
Someya et al., 1997
) and the
majority of the oxidase is located on the plasma membrane
(Lacy et al., 2003
). By
contrast, the vast majority of neutrophil oxidase is located intracellularly.
This difference in location is consistent with the distinct functional roles
of neutrophils and eosinophils. Neutrophils kill primarily by phagocytosing
their targets, whereas eosinophils kill non-phagocytosable targets by
secreting toxic agents.
In these experiments, membrane potential measurements were made using the
cell-permeable fluorescent dye, diBAC4(3). A primary concern when
working with membrane potential-sensitive dyes is the possibility of reporting
intracellular compartment membrane potentials instead of the plasma membrane
potential and the dye's possible sensitivity to the oxygen radicals. Our data
probably reflect the plasma membrane potential for several reasons. First,
although the sulfur-containing, cationic carbocyanine dyes have been reported
to be quenched during superoxide production, this is unlikely to occur with
diBAC4(3), an anionic, non-sulfur-containing oxonol dye
(Whitin et al., 1981). Second,
eosinophils, in contrast to neutrophils or monocytes, are not primarily
phagocytic and thus are less likely to accumulate extracellular dye in
phagosomes. Third, confocal microscopy documents that the dye is localized to
the cytosol of the eosinophil and not sequestered in the secondary granules or
other intracellular organelles. Fourth, altering the external conditions of
the cell by adding ZnCl2 or changing the pHe affects
diBAC4(3) fluorescence as would be predicted if the dye was
measuring the plasma membrane potential. Additionally, the effects of
ZnCl2 and pHe changes on eosinophil membrane potential
are consistent with previous reports
(Henderson et al., 1987
;
Susztak et al., 1997
;
Banfi et al., 1999
).
Proton efflux through the NADPH oxidase-associated proton channel appears
to be required for continued superoxide production by neutrophils and
eosinophils. Previous studies, using ZnCl2 to block proton efflux,
document inhibited superoxide production (Henderson et al., 1988a;
Bankers-Fulbright et al.,
2001). This is probably due to the lack of electrogenic
compensation for electron transfer through the membrane by NADPH oxidase,
allowing unattenuated depolarization and proton accumulation at the inner
membrane surface (Henderson et al., 1988a). Although originally thought to be
part of the NADPH oxidase complex, the proton channel now appears to be a
distinct protein that is, nevertheless, concomitantly regulated by PKC
(DeCoursey et al., 2001a
).
Stimuli such as arachidonic acid and PMA, which activate the proton channels
in neutrophils and eosinophils, also stimulate electron transport through the
oxidase (DeCoursey et al.,
2000
; DeCoursey et al.,
2001b
; Cherny et al.,
2001
). Although PMA-stimulated NADPH oxidase activity can be
blocked with the PKC
selective inhibitor, rottlerin, the proton
conductance is insensitive to PKC
inhibition
(Bankers-Fulbright, 2001
).
Consistent with our previous data, we report here that rottlerin also inhibits
PMA-stimulated eosinophil depolarization
(Fig. 4). Thus, the proton
channel and electron transport appear to be regulated independently by
different PKC isoforms.
Pharmacological specificity is always a concern when using inhibitors. At
the concentrations used in this paper, rottlerin is selective for PKC
and does not inhibit the activity of any other known PKC isoforms. However,
rottlerin can block calmodulin-dependent (CaM) kinase III (Geschwendt et al.,
1994). CaM kinase III is unlikely to be the target of rottlerin in
PMA-stimulated eosinophils because stimulation with PMA does not increase
intracellular calcium concentrations (data not shown). Additionally, we have
previously shown that PKC
is indeed activated in PMA-stimulated
eosinophils, and this activation is inhibited by rottlerin
(Bankers-Fulbright, 2001
).
In summary, we have shown that PMA-stimulated eosinophils depolarize to
voltages sufficient to activate the proton conductance, and proton efflux can
regulate the extent of depolarization. PKC appears to be necessary for
PMA-induced depolarization and superoxide production, but not for proton
channel activation in human eosinophils.
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Acknowledgments |
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