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INTRODUCTION |
Under normal conditions, healthy cell membranes resist catalysis
by secretory phospholipase A2
(sPLA2)1 (1-4).
However, they may become susceptible under circumstances that cause
alteration of membrane physical properties (1-4). Previous studies
using artificial membranes demonstrated that alterations that increase
susceptibility generally increase the anionic charge of the outer
leaflet, increase bilayer curvature, and/or decrease interactions among
neighboring phospholipids (5-9). In some cases, enhanced
susceptibility of artificial membranes depends on an increase in the
order of the phospholipids (8, 10-14). These changes increase
susceptibility by augmenting the binding of sPLA2 and/or by
improving access of membrane phospholipids to the active site of the
enzyme (5-12, 15, 16).
It is not known whether the properties that induce susceptibility to
sPLA2 in artificial membranes also contribute to the vulnerability of biological membranes to attack by the enzyme. In order
to address this issue, we manipulated various properties of erythrocyte
membranes by preparing different types of ghosts as explained in the
accompanying particle (17). We found that the factors that determined
the degree of susceptibility were increased exposure of
phosphatidylserine, an anionic phospholipid, and increased membrane
order. These interpretations agreed with those from studies of
susceptibility using artificial membranes (5-16). The next question,
then, is whether these same factors are important in the hydrolysis of
intact cells by sPLA2 under conditions at which they have
become susceptible such as in the presence of specific hormones, after
treatment with certain toxins, during apoptosis, or following cellular
trauma (2-4, 18).
One feature common among many of the conditions that render cells
susceptible to sPLA2 is the elevation of intracellular
calcium (1-4). We have used human erythrocytes as an experimental
model to determine whether phosphatidylserine exposure and/or an
increase in the order of membrane phospholipids are relevant factors in the induction of catalysis by sPLA2 when intracellular
calcium is increased. In addition, we examined other hypotheses that
have been proposed to explain the ability of certain agents to render cell membranes susceptible to sPLA2: 1) prior activation of
intracellular phospholipase(s) A2 (19, 20), 2) release of
microvesicles from the plasma membrane (1), 3) oxidation of membrane
phospholipids (21).
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EXPERIMENTAL PROCEDURES |
Materials--
Erythrocytes were obtained from healthy
individuals undergoing routine physicals at Brigham Young University
McDonald Health Center. The samples were stored overnight at 4 °C in
EDTA vacutainers from which patient identification was removed. Control
experiments comparing fresh blood with samples stored overnight
demonstrated that the storage conditions did not influence the results.
Erythrocytes were isolated by centrifugation, and resuspended to the
original hematocrit in MBSS (NaCl = 134 mM, KCl = 6.2 mM, CaCl2 = 1.6 mM, MgCl2 = 1.2 mM, Hepes = 18.0 mM, and glucose = 13.6 mM, pH 7.4, 37 °C).
Snake venom sPLA2 (monomeric aspartate 49 (AppD49) from the
venom of Agkistrodon piscivorus piscivorus) was isolated
according to published procedures and was used in all experiments
except those shown in Fig. 13 (22). Human group V and IIa
sPLA2 were provided generously by Wonhwa Cho (University of
Illinois, Chicago, IL) and Michael Gelb (University of Washington,
Seattle, WA). The final concentrations of sPLA2 used
in experiments were 1 µg/ml for AppD49 and human group V and 2 µg/ml for human group IIa.
ADIFAB, laurdan, merocyanine 540, and cis-parinaric acid were obtained
from Molecular Probes (Eugene, OR). Ionomycin and E-64d were procured
from Calbiochem (La Jolla, CA), and phenylhydrazine, diamide, and
quinine were obtained from Sigma. The scramblase inhibitor, R5421, was
a kind gift from Jeffrey T. Billheimer at Dupont Merck Pharmaceutical
Co. (Wilmington, DE). DAPA, factor Va, factor Xa, prothrombin, and
thrombin were acquired from Hematologic Technologies, Inc. (Essex
Junction, VT). Pharmacological agents were dissolved in the appropriate
solvents (Me2SO or ethanol). Control experiments
demonstrated that these solvents did not have effects on the
experimental data at the concentrations used.
Phospholipid Extraction and Thin Layer
Chromatography--
Washed erythrocytes (30 µl) were suspended in
MBSS to a final volume of 1 ml (about 1.5 × 108
cells/ml) and incubated in the presence or absence of 0.3 µM ionomycin with or without AppD49 sPLA2 for
20 min at 37 °C. Cells were then separated by centrifugation (6500 rpm for 60 s) in a microcentrifuge (about 3000 × g) and pellets were frozen in liquid nitrogen to quench the
reaction. Samples were quickly thawed, and lipids were extracted with
chloroform and methanol by the method of Bligh and Dyer (23). In brief,
100 µl of chilled MBSS was added to suspend the pellet followed by
125 µl of chloroform and 250 µl of methanol. After vortexing the
tubes for 10 s, 125 µl of water were added. The samples were
then vortexed and centrifuged for 30 s at 3000 × g. After removing half of the water layer, the protein layer
was carefully removed with a pipette tip. The remainder of the water
layer was then discarded and the residual organic layer was dried under
a nitrogen stream to ~10% of the original volume. The sample was
then spotted onto a silica gel thin-layer chromatography plate.
Phospholipids and lysophospholipids were separated by thin-layer
chromatography in 6.5:2.5:1 (v/v) chloroform:methanol:acetic acid.
Lipids were stained by iodine vapor. Spots were identified by
comparison to standards. The resulting phosphatidylcholine and
phosphatidylethanolamine spots on the silica gel were analyzed by both
phosphate assay according to the method of Bartlett (24) and by
densitometry. For densitometric measurements, samples were photographed
with a digital camera using a Coomassie Blue filter under direct light
and the digital image was quantified using standard digitizing computer software.
Fluorescence Spectroscopy--
Washed erythrocytes were
suspended in 2 ml of MBSS in a fluorometer sample cell to a final
density of about 3-4 × 106 cells/ml. Measurements
with fluorescent probes were obtained at 37 °C using a Fluoromax
(Spex Industries) photon-counting spectrofluorometer. Sample
homogeneity was maintained by continuous gentle stirring with a
magnetic stir bar. Simultaneous assessment of fluorescence intensity at
multiple excitation and emission wavelengths was obtained by rapid
sluing of monochromator mirrors using control software provided with
the instrument. Band pass was set at 4.25 nm for all experiments.
Hydrolysis by sPLA2--
Release of fatty acids from
cells was assayed with an acrylodan-labeled fatty acid-binding protein
(ADIFAB) (65 nM final, excitation = 390 nm,
emission = 432 and 505 nm; Refs. 3 and 25). The results were
quantified by calculation of the generalized polarization (GP) as
described (3, 26). The values of GP as a function of time were fit to a
double exponential equation by nonlinear regression. The amount of
hydrolysis at 100 s following sPLA2 addition was then
calculated using parameter values from the nonlinear regression results.
Prothrombinase Assay--
Exposure of phosphatidylserine in the
outer leaflet of the bilayer was detected by an increase in the
fluorescence intensity of
dansylarginine-N-(3-ethyl-1,5-pentanediyl)amide (DAPA) (3 µM final, excitation = 335 nm, emission = 545 nm; Ref. 27). To detect phospholipid translocation, DAPA, factor Va (6 nM), factor Xa (3 nM), and prothrombin (3.5-4
µM) were incubated for 300 s in MBSS in the sample
chamber of the spectrofluorometer. Cells were then added and the
mixture incubated an additional 600 s prior to addition of
ionomycin (0.3 µM) or control solvent
(Me2SO). A positive control was obtained with the addition
of thrombin (2.7 µM).
Microvesicle Release--
The release of vesicles from the
plasma membrane was monitored simultaneously with other fluorescence
observations by recording the intensity of scattered light
(excitation = 500 nm, emission = 510 nm; Ref. 4). For
simultaneous measurements with the prothrombinase assay (see above),
excitation and emission wavelengths were 600 and 610 nm, respectively.
Oxidation--
Oxidation of membrane phospholipids was monitored
by use of the fluorescent probe, cis-parinaric acid (1.12 µM final, excitation = 303 nm, emission = 416 nm; Ref. 28). Measurements of light scattering for microvesicle release
and ADIFAB fluorescence were made simultaneously. The data were
corrected for time-dependent light scattering artifacts
caused by microvesicle release.
Scanning Electron Microscopy--
Erythrocytes were prepared by
a modification of Schneider's method (29). Briefly, the preparations
were washed in 0.1 M phosphate buffer at pH 7.4. 9.5 ml of
1×108 cells/ml were incubated in a jar having a 5.5-cm
diameter, in the presence or absence of 0.3 µM ionomycin,
and allowed to settle onto cover glasses, previously coated with
poly-L-lysine, at 4 °C overnight. Samples were then
fixed in 2% glutaraldehyde for 2.5 h. Following fixation, the
cells were washed six times in sodium cacodylate buffer (pH 7.3), fixed
in 2% osmium tetroxide for 2 h at 23 °C, and washed six times
in sodium cacodylate buffer. Samples were dehydrated through a graded
series of ethanol solutions (10, 30, 50, 70, 95, and 100%) for 10 min
each then washed three times in acetone. The slides were then subjected
to critical point drying, using carbon dioxide. Finally, samples were
sputter coated with gold for 2 min. Images were obtained on a JEOL JSM
840A scanning electron microscope.
Membrane Fluidity--
Membrane order was assessed using laurdan
GP (26). Laurdan (2.5 µM final) was added to samples of
erythrocytes prepared as described above for fluorescence spectroscopy.
Fluorescence emission was then monitored as a function of time at dual
wavelengths (excitation = 350 nm, emission = 435 and 500 nm)
for at least 5 min to establish the baseline. Various agents followed
by ionomycin or control solvent were added as described in Fig. 9.
Changes in laurdan GP were then assessed by calculating the difference in the slope of laurdan GP before and after addition of ionomycin or
control solvent under each experimental condition.
Two-photon Microscopy--
The two-photon excitation images were
collected on an Axiovert 35 inverted microscope (Zeiss, Thornwood, NY),
with a Zeiss 20X LD-Achroplan (0.4 N.A., air) using a titanium-sapphire
laser excitation source (Coherent, Palo Alto, CA) tuned to 770 nm and pumped by a frequency-doubled Nd:vanadate laser (Coherent, Palo Alto,
CA) as described previously (30). The laser was guided by a
galvanometer-driven x-y scanner (Cambridge
Technology, Watertown, MA) to achieve beam scanning in both
x and y directions. A frequency synthesizer (Hewlett-Packard, Santa Clara, CA) controlled
the scanning rate of 9 s to acquire a 256 × 256-pixel frame
that covered approximately a 60 × 60-µm region. Dual images
were collected simultaneously using a beam-splitter, two emission
short-pass filters (centered at about 450 and 500 nm), and two
detectors for calculation of GP (26).
Samples were incubated with or without the agents indicated in Fig. 11
as described above for hydrolysis experiments. Laurdan (250 nM) was added to the samples 250 s after the addition
of the last agent in the experiment. Samples were incubated with stirring for an additional 50 s, and a 0.5-ml aliquot was then transferred to 1 ml of fresh MBSS in a heated microscopy sample dish
(36 °C). Cells were allowed an additional 5 min to settle, and
images were then obtained.
In some cases (e.g. Fig. 10), cells were incubated prior to
the onset of the experiment with 5 µM laurdan for 1 h at 36 °C, and excess laurdan was removed by centrifugation. Cells
were suspended in 2 ml of fresh MBSS and transferred to the microscopy
dish. After allowing the cells to settle, baseline images were
obtained. Ionomycin was then added directly to the sample, and
additional images were acquired. Finally, sPLA2 was added,
and the time course of changes in laurdan fluorescence was monitored by
repeated acquisition of images of the same field.
Statistical Analysis--
In all figures that contain summaries
from multiple replicates, the data are expressed as the mean ± S.E. Each replicate sample included in the data represents data from a
separate blood donor. Large comparisons of hydrolysis or light
scattering data among many groups sharing some of the same blood
samples (e.g. Figs. 2, 7, 9, and 12) were accomplished by
one-way analysis of variance followed by Dunnett's post-test for
multiple comparisons. Since the number of samples per group was
unbalanced, it was not possible to consider sample pairing in the
analyses of variance. This increased the possibility of missing real
differences that would only be identified by paired comparisons of
samples matched by blood donor. Accordingly, the various treatment
groups were also compared with the group treated with ionomycin alone
using Student's paired t test (two-tailed) for those
samples that were matched by blood donor. Although the results of this
secondary analysis agreed with those of the analysis of variance and
post-test in most instances, there was one example in which the results
were significant only when the analysis was confined to paired samples.
In this case, the level of significance was very high
(p = 0.004), and the data were therefore interpreted as
being statistically significant (see legend to Fig. 7).
When data sets were fully matched by blood sample for all treatment
groups (i.e. Figs. 3, 4, and 13), they were analyzed in two
steps. First, results within each treatment group were normalized to
the value of an appropriate internal standard matched by blood sample
(see figure legends for details). Second, the normalized values were
tested for treatment effects using Student's t test (two-tailed) with the value of 1.0 as the null hypothesis. Since multiple (two to three) treatment groups were compared with the same
standard in these cases, a correction was made to the critical value of
p accepted as indicating statistical significance
(traditionally 0.05) according to the formula p = 1-0.951/n, where n = the number of
comparisons. For n = 2, the critical value of
p = 0.025; for n = 3, the critical
value of p = 0.017.
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RESULTS |
Effect of Ionomycin--
As shown in Fig.
1, the extent of fatty acid release from
erythrocyte membranes in the presence of sPLA2 was greatly
enhanced by a 10-min prior incubation of the cells with ionomycin. The average response among multiple samples is displayed in Fig.
2. Control experiments in which
Ca2+ was replaced by EGTA in the extracellular medium
revealed that this and the other results described below for ionomycin
were due to Ca2+ entry into the cell rather than direct
effects of the ionophore. Experiments in which the time of incubation
with ionomycin was varied revealed that the effect developed after a
latency of about 100 s and reached a maximum within about 300 s (not shown). In contrast to results obtained with lymphocytes (4),
the hydrolysis was not sufficient to consume the cells, and little or
no hemolysis was observed at the end of the reaction.

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Fig. 1.
Ionomycin induces susceptibility to
sPLA2 in human erythrocytes. Erythrocytes were
incubated with 0.3 µM ionomycin (triangles) or
control diluent (Me2SO, circles) for 10 min, and
sPLA2 was then added (time 0 on graph). Hydrolysis was
monitored with ADIFAB, and data were fit by nonlinear regression as
described under "Experimental Procedures." For a statistical
analysis of the reproducibility of this result, see Fig. 2.
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Fig. 2.
Effects of various agents on the ability of
ionomycin to induce susceptibility to sPLA2. The
experiments of Fig. 1 were repeated with the following conditions.
Control: Me2SO 10 min, n = 20. Ionomycin: 0.3 µM ionomycin 10 min,
n = 23. High KCl + ionomycin: 10 min
incubation in MBSS with 89 mM KCl and 51 mM
NaCl then ionomycin added for 10 min, n = 5. Quinine + ionomycin: 10 min incubation in MBSS containing
1 mM quinine then ionomycin added for 10 min,
n = 4. E-64d + ionomycin: 36 µM E-64d 5 min then ionomycin added for 10 min,
n = 3. R5421 + ionomycin: 50 µM R5421 10 min then ionomycin added for 10 min,
n = 5. Phenylhydrazine: 0.5 mM
phenylhydrazine 10 min, n = 4. Phenylhydrazine + ionomycin: phenylhydrazine and ionomycin together 10 min,
n = 5. Diamide: 50 µM diamide
10 min, n = 4. Diamide + ionomycin: 50 µM diamide and ionomycin together 10 min,
n = 5. At the end of each of these incubations,
sPLA2 was added and the incubation continued. The
ordinate indicates the level of ADIFAB GP assessed from the
nonlinear regression of the data at 100 s following addition of
sPLA2. Asterisks represent values that differed
significantly from ionomycin alone (**: p < 0.01, by
analysis of variance as explained under "Experimental Procedures").
High KCl, quinine, or R5421 alone had no effect (not shown).
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Comparative Hydrolysis of Phosphatidylcholine and
Phosphatidylethanolamine--
In order to assess the relative amount
of hydrolysis of the two major glycerophospholipid species, experiments
were conducted at much higher cell densities (about 40-fold higher)
than in the experiment of Fig. 1. Under such conditions, the time
course of onset of the effect of ionomycin was much slower such that
the lag time for achieving maximum hydrolysis rates was about 20 min. The release of microvesicles in the presence of ionomycin described below was also proportionally slower. Thin-layer chromatography experiments investigating hydrolysis of erythrocytes in the presence of
sPLA2 or ionomycin alone revealed no significant hydrolysis of either phosphatidylcholine or phosphatidylethanolamine after a
20-min incubation. In contrast, both substrates were hydrolyzed significantly when erythrocytes were incubated with sPLA2
and ionomycin together (Fig. 3).

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Fig. 3.
Hydrolysis of phosphatidylethanolamine
(PE) and phosphatidylcholine (PC) in
the presence of ionomycin and/or sPLA2. Erythrocytes
were incubated with or without 0.3 µM ionomycin for 5 min; then sPLA2 was added to half of the samples and the
incubation continued for 20 min. Aliquots were removed, and
phospholipids were extracted and separated by thin-layer chromatography
as explained under "Experimental Procedures." The data were
normalized to the amount of phospholipid observed under control
conditions (no ionomycin or sPLA2). Significance was
assessed by Student's t test with adjustment for multiple
comparisons as explained under "Experimental Procedures" (*,
p = 0.013 for "PC both," and p = 0.0016 for "PE both," n = 3).
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Intracellular Phospholipase A2--
Erythrocytes were
incubated with ionomycin in the presence or absence of MAFP (19) or BEL
(31), inhibitors of intracellular phospholipases A2. As
shown in Fig. 4, the amount of hydrolysis after 100 s with sPLA2 was not statistically different
in cells treated with MAFP and ionomycin compared with cells treated
with only ionomycin. Similar results were obtained with BEL as the inhibitor (data not shown). When MAFP was incubated with erythrocytes at low cell density in the spectrofluorometer (i.e. as in
Figs. 1 and 2), it caused nonspecific perturbation of the cell membrane that directly rendered the cells susceptible to sPLA2.
Consequently, these experiments were conducted at higher cell densities
and longer incubation times similar to the experiments described above for thin-layer chromatography (i.e. Fig. 3). The remainder
of the experiments described below were completed at low cell density in the spectrofluorometer as in Figs. 1 and 2. The less specific inhibitor, AACOCF3, that had previously been shown to
inhibit an intracellular phospholipase A2 activity in
erythrocytes was also tested (32). Like MAFP and BEL, it did not reduce
the susceptibility induced by ionomycin (data not shown). Therefore,
activation of intracellular phospholipases A2 appeared
unnecessary for Ca2+-induced susceptibility to
sPLA2 in erythrocytes.

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Fig. 4.
Inhibition of intracellular phospholipase
A2 does not alter the ability of ionomycin to induce
susceptibility. Erythrocytes were incubated for 10 min in the
presence or absence of 20 µM MAFP under the conditions
described for the thin-layer chromatography experiments
(i.e. Fig. 3). Ionomycin (0.3 µM) was then
added and the incubation continued for 45 min. A 50-µl aliquot of
each sample was then transferred to a spectrofluorometer sample cell (2 ml final volume), and susceptibility was then assessed using ADIFAB as
described under "Experimental Procedures." Significance was
assessed by normalizing the data for each group to that obtained with
ionomycin alone and evaluated by Student's t test as
described under "Experimental Procedures" (***, p < 0.0001).
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Phospholipid Translocation--
Ionomycin stimulates transbilayer
migration of membrane phospholipids (33). Fig.
5 demonstrates the time course of
phosphatidylserine exposure (increase in DAPA fluorescence intensity)
upon the addition of ionomycin. Prior treatment of cells for 10 min
with R5421, an inhibitor of scramblase activity (34), caused a
substantial decrease in the exposure of phosphatidylserine stimulated
by ionomycin. Inhibition of phospholipid translocation by R5421 did not
alter the susceptibility to sPLA2 in the presence of
ionomycin (Fig. 2).

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Fig. 5.
Ability of R5421 to inhibit
phosphatidylserine translocation in the presence of ionomycin.
Cells were incubated with clotting factors Va and Xa, prothrombin, and
the fluorescent thrombin substrate DAPA for 10 min in the absence
(curve a) or presence (curve b) of 50 µM R5421 as described under "Experimental
Procedures." Ionomycin (0.3 µM) was added at the
arrow. The ordinate represents relative
fluorescence intensity of DAPA. This experiment is representative of
three independent experiments. The data have been corrected for a
linear baseline increase in fluorescence.
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Microvesicle Release--
Upon introduction of ionomycin,
erythrocytes released microvesicles after a short lag period as
expected (1). The release of these vesicles was conveniently monitored
in real time concurrently with assessment of susceptibility by
measuring the amount of light scattered by the sample at 500 nm. As
shown in Fig. 6, the intensity of
scattered light increased about 100 s after addition of ionomycin and rose until reaching a plateau about 500 s later. We verified this interpretation by direct observation of the samples by scanning electron microscopy. As shown in Fig. 6C, treatment with ionomycin caused the erythrocytes to assume a diminished size and rounded shape
(spherocyte) and to extrude small pieces of its membrane as
microvesicles.

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Fig. 6.
Microvesicle release and shape transition
induced by ionomycin. Panel A, the intensity of light
scattering by erythrocytes was monitored as a function of time as
described under "Experimental Procedures." At the time indicated by
the arrow, 0.3 µM ionomycin (curve a) or
control diluent (curve b) was added. The curves
are offset along the ordinate for clarity. For a statistical analysis
of the reproducibility of the results, see Fig. 7. Panel B,
scanning electron micrograph of control erythrocyte. Panel
C, scanning electron micrograph of erythrocyte treated with
ionomycin. See "Experimental Procedures" for details.
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To test whether the microvesicle release was required for
susceptibility to occur, we prevented the release by adding either high
KCl (89 mM) or a Ca+2-activated K+
channel blocker (quinine) to the extracellular medium (35, 36). Cells
incubated in the high KCl buffer demonstrated a decrease, while cells
treated with quinine showed a complete inhibition, of the amount of
microvesicles present after treatment with ionomycin (Fig.
7). In the case of quinine, a decrease in
light scattering intensity was observed presumably because of the
reduction in cell size due to the shape transition (37). There was no
significant effect of either treatment on the ability of ionomycin to
induce susceptibility to sPLA2 (Fig. 2).

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Fig. 7.
Effects of various agents on the ability of
ionomycin to induce microvesicle release. The experiments of Fig.
6 were repeated in the presence or absence of the agents listed. The
ordinate indicates the change in average intensity of
scattered light from that measured immediately prior to ionomycin (or
phenylhydrazine or control diluent) addition to that observed 10 min
later. Asterisks represent values that differed
significantly from ionomycin alone (*, p < 0.05; **,
p < 0.01, by analysis of variance; ***,
p = 0.004 by paired t test; the details of
both analyses are explained under "Experimental Procedures"). High
KCl and quinine alone were indistinguishable from control. The numbers
of replicates were: Control, 37; Ionomycin, 52;
High KCl + ionomycin, 12; Quinine + ionomycin, 6;
E-64d + ionomycin, 3; Phenylhydrazine, 4;
Phenylhydrazine + ionomycin, 16. See Fig. 2 for the details
of the concentrations and incubation times.
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Microvesicle release was also inhibited by use of an erythrocyte (type
µ) calpain inhibitor, E-64d (38). One of the effects of elevated
intracellular Ca2+ in erythrocytes and platelets is
activation of calpain, an intracellular cytoskeletal protease (38). The
resulting cytoskeletal damage appears to be involved in the process of
microvesicle release (39). As expected, E-64d reduced significantly the
level of microvesicle release observed in the presence of ionomycin
(Fig. 7). However, like the other inhibitors of microvesicle release, it did not reduce ionomycin-stimulated susceptibility to
sPLA2 (Fig. 2). Higher concentrations (up to 140 µM), while able to block microvesicle release completely,
also did not lower the level of hydrolysis by sPLA2 (not
shown). Therefore, it appeared that microvesicle release was not
necessary for the cells to become susceptible to sPLA2.
Oxidation--
In contrast to the positive controls, diamide and
phenylhydrazine, ionomycin did not cause oxidation (i.e.
reduction of cis-parinaric acid intensity) of erythrocyte membranes
(Fig. 8). Erythrocytes treated with
diamide alone for 10 min did not become susceptible to
sPLA2. Likewise, diamide did not alter microvesicle release (not shown) nor the amount of hydrolysis observed when ionomycin was
present (Fig. 2). A second oxidizing agent, phenylhydrazine, also did
not cause the cells to become susceptible during a 10-min incubation
(Fig. 2). Interestingly, in contrast to diamide, phenylhydrazine significantly impaired the effect of ionomycin on susceptibility (Fig.
2). In addition, phenylhydrazine caused an increase in the intensity of
scattered light reminiscent of the effect of ionomycin to induce
microvesicle release (Fig. 7).

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Fig. 8.
Ionomycin treatment does not cause oxidation
of erythrocytes. Erythrocyte membrane oxidation was monitored with
cis-parinaric acid as described under "Experimental Procedures." At
the time indicated by the arrow, control diluent
(curve a), 0.3 µM ionomycin (curve
b), 50 µM diamide (curve c), or 0.5 mM phenylhydrazine (curve d) was added. These
experiments are representative of four independent experiments. Curves
are offset slightly along the ordinate for clarity of
presentation.
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Membrane Order--
Fluidity of the membrane was assessed by
fluorescence spectroscopy. Cells were labeled with laurdan, and
the effects of various agents on GP values were determined. In
general, an increase in the value of GP corresponds to an increase in
membrane order (26). As shown in Fig. 9,
ionomycin treatment caused a reproducible elevation of the value of GP.
This effect was blocked by EGTA demonstrating that it required
Ca2+ and was not simply a direct effect of intercalation of
ionophore into the membrane. Incubation of the cells in high KCl,
E-64d, or R5421 had no significant effect on the response to ionomycin. Since phenylhydrazine treatment inhibited the ability of ionomycin to
induce susceptibility, we also considered its effect on membrane order.
In contrast to the other agents tested, phenylhydrazine did cause a
significant decrease in the GP value.

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Fig. 9.
Effects of various agents on laurdan GP
assessed by steady state fluorescence. The ordinate
indicates the change in average GP slope before and after the addition
of ionomycin. Asterisks represent values that differed
significantly from ionomycin alone (*, p < 0.05; **,
p < 0.01, by analysis of variance as explained under
"Experimental Procedures"). The numbers of replicates
were: Control, 13; Ionomycin, 20; EGTA + ionomycin, 4; High KCl + ionomycin, 3; E-64d + ionomycin, 6; R5421 + ionomycin, 3;
Phenylhydrazine + ionomycin, 4. Concentrations and
incubation times were the same as described in the legend to Fig. 2
with the exception that cells were incubated 5 min with phenylhydrazine
prior to ionomycin addition. In the EGTA experiment, 1 mM
EGTA replaced the 1.6 mM Ca2+ in the
MBSS.
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Control observations revealed that the effect of phenylhydrazine on
laurdan GP was caused, at least in part, by time-dependent changes in the optical density of phenylhydrazine. We therefore repeated some of the experiments of Fig. 9 using two-photon microscopy to detect laurdan GP under conditions at which laurdan fluorescence arising directly from the membrane could be distinguished from indirect
optical effects of the experimental agents. As shown in Fig.
10A, untreated cells
displayed a non-uniform distribution of laurdan GP values. Higher
values were concentrated along the rims of the diskocytes.
The addition of ionomycin increased the GP value of these peripheral
regions and expanded their size (Figs. 10B and
11A). In agreement with the measurements displayed in Fig. 9, phenylhydrazine blocked the response to ionomycin (Fig.
11B).

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Fig. 10.
Change in laurdan GP distribution during
hydrolysis by sPLA2 after addition of ionomycin assessed by
two-photon microscopy. The colors represent the relative
fluidity of each area of the membrane. Blue proceeding
through red indicates an increase in membrane order.
Panel A, two-photon micrographs of erythrocytes prior to
treatment (GP = 0.019 ± 0.18, mean ± S.D. from fits
to the Gaussian distribution for the image); panel B, 10 min
after ionomycin addition (GP = 0.12 ± 0.18); panel
C, 1 min after sPLA2 addition (GP = 0.16 ± 0.16); panel D, 4 min after sPLA2 addition
(GP = 0.24 ± 0.15).
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Fig. 11.
Effects of various agents on laurdan GP
assessed by two-photon microscopy. Panel A,
distribution of GP values for erythrocytes in the absence (closed
squares) or presence (open circles) of ionomycin
obtained from images such as those shown in Fig. 10. The
curves represent nonlinear regression of the data using the
Gaussian distribution. Panel B, the average value of GP was
calculated from fits to the Gaussian distribution for images such as
those illustrated in Fig. 11A under the indicated conditions
and subtracted from the average control value. Data are expressed as
the mean ± S.E. for 5-7 images per condition. See Fig. 2 for the
details of the concentrations and incubation times.
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Two-photon images of erythrocyte ghosts also revealed a non-uniform
distribution of laurdan GP (17). Hydrolysis by sPLA2 appeared related to the presence of regions of high GP since these regions expanded and became more ordered following sPLA2
addition (17). As shown in Fig. 10, the same phenomenon was observed
with intact erythrocytes treated with ionomycin. After addition of sPLA2, regions of low fluidity (yellow to
red color) expanded systematically and became more ordered
(Fig. 10, C and D).
The possibility that the susceptibility of erythrocytes to
sPLA2 was dependent on membrane order was further
investigated using merocyanine 540. Merocyanine 540 binds to the outer
leaflet of erythrocyte membranes and induces the shape transition from diskocytes to spherocytes without release of microvesicles or flip-flop
of membrane lipids (40, 41). Addition of merocyanine 540 to
erythrocytes also caused a significant increase in laurdan GP
(0.17 ± 0.009 GP units, mean ± S.E., n = 3, p < 0.003 by Student's one-sample t test)
comparable to that produced by ionomycin (e.g. Fig.
11B). Likewise, the agent rendered the membranes susceptible to sPLA2 (Fig. 12, control
and ionomycin data from Fig. 2 included for comparison).

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Fig. 12.
Effect of merocyanine 540 on susceptibility
to sPLA2. Hydrolysis of erythrocyte phospholipids by
sPLA2 was assessed in the presence of 10 µM
merocyanine 540 (MC540) (10 min incubation prior to sPLA2
addition) as described in Fig. 2. The "control" and "ionomycin"
data of Fig. 2 are included for comparison. The amount of hydrolysis
observed with merocyanine 540 was significantly different from the
control (no merocyanine) by analysis of variance (*, p < 0.05, n = 4).
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Human sPLA2--
Fig.
13 displays repetition of key
experiments using human group V sPLA2 instead of the snake
venom enzyme. The extent of hydrolysis was about half of that observed
with the AppD49 enzyme as reported previously (4). Nevertheless, the
fundamental trends observed with ionomycin and phenylhydrazine were
similar for the human sPLA2 compared with the venom enzyme
(compare Figs. 2 and 13). Experiments were also repeated with human
group IIa sPLA2. In this case, however, the activity was
very low, and quantitative interpretation of the data was not
feasible.

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Fig. 13.
Ionomycin induces susceptibility to
catalysis by human Group V sPLA2. The experiments
shown in Fig. 1 were repeated with or without phenylhydrazine (0.5 mM). For statistical analysis, the results were normalized
to the amount of hydrolysis observed in the "Ionomycin" treatment
group. Significance was assessed by Student's t test as
explained under "Experimental Procedures" (control,
p = 0.015; Phenylhydrazine + ionomycin,
p = 0.002, n = 4).
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DISCUSSION |
The ability of intracellular Ca2+ to govern the
susceptibility of cell membranes to sPLA2 has been observed
in a number of cell types. For example, S49 lymphoma cells normally
resist the action of sPLA2 until treated with agents that
elevate intracellular Ca2+ levels such as ionophore,
lysolecithin, or the plant toxin thionin (3, 4). A similar phenomenon
has been observed in other cells such as HL-60, MOLT-4, Raji,
erythrocytes, and platelets (1, 4, 42).
While it is clear that this phenomenon is general, at least among blood
cells, the mechanisms involved are much less established. Nevertheless,
a few hypotheses have been proposed based on a variety of observations:
1) prior activation of intracellular phospholipase(s) A2
(19, 20, 43, 44); 2) release of microvesicles from the plasma membrane
(1); 3) oxidation of membrane phospholipids (21); 4) transbilayer
migration of phosphatidylserine and phosphatidylethanolamine (2, 9, 16,
45); and 5) changes to other microscopic physical properties of the
membrane (17). Based on the data shown in Figs. 2 and 6-8, the first
three hypotheses were excluded as explanations for the susceptibility
to sPLA2 observed in the presence of ionomycin. As
discussed below, the results of this study combined with those of the
accompanying article (17) contend that alterations to specific physical
properties related to membrane fluidity are responsible for
susceptibility to the enzyme. Importantly, these results validate the
assumption that information obtained from studies of artificial
bilayers relates to biological membranes.
The results reported with erythrocyte ghosts in the accompanying paper
(17) suggest that exposure of phosphatidylserine can promote
susceptibility, although multiple regression analysis revealed that it
was a less important contributor than membrane properties assessed by
laurdan. What the experiments with ghosts were unable to determine was
whether phosphatidylserine exposure was required or instead simply
ancillary or even redundant for making the membrane susceptible to the
enzyme. The logic of the two studies was to identify first in the
ghosts possible candidates for the relevant membrane changes and then
ask whether those changes applied to Ca2+ ionophore
treatment of intact erythrocytes. In the case of phosphatidylserine exposure, the appropriate conclusion is that such may promote susceptibility, but it is not an absolute requirement during ionomycin treatment of erythrocytes. This assertion is based on two results. First, R5421 treatment inhibited the exposure of phosphatidylserine substantially (Fig. 5) but did not affect the amount of hydrolysis catalyzed by sPLA2 in the presence of ionomycin (Fig. 2).
Second, merocyanine 540, which does not induce translocation of
phosphatidylserine (41), was able to cause susceptibility. These
observations corroborate results obtained with S49 cells in which it
was shown that susceptibility to sPLA2 during apoptosis
precedes significant exposure of phosphatidylserine (46).
Comparison of the laurdan results shown in Figs. 9-11 with the
susceptibility data (Fig. 2) suggests that changes in membrane order
could be largely responsible for the induction of susceptibility by
ionomycin. The agreement with the results from erythrocyte ghosts
described in the accompanying article (17) is strong. First, membrane
order was found to be the major predictor of susceptibility in ghosts
when the various factors were considered together in multiple
regression analysis. Second, comparison of the levels of susceptibility
and change in GP induced by ionomycin (Figs. 2 and 11) with those
reported for the ghosts demonstrates that the similarity is
quantitative as well as qualitative. In addition, the data obtained
with merocyanine 540 suggest that the relationship between membrane
order and susceptibility is a general phenomenon rather than depending
on influx of calcium (Fig. 12). Importantly, these results support the
concept that principles learned from biophysical studies with
artificial membranes apply to biological systems (5-16). Attempts have
been made previously with cultured cells to determine whether changes
in membrane order detectable with fluorescent probes might explain the
action of Ca2+ to render them vulnerable to
sPLA2 (3, 4). The results from those studies were
inconclusive probably because of the diversity of membranes accessible
to the probes. These studies with erythrocytes have the experimental
and interpretive advantage of avoiding complications due to
intracellular membranes.
It is likely that regional increases in the order of membrane lipids
increases susceptibility both by enhancing the binding of the enzyme as
well as creating membrane defects that facilitate migration of
phospholipids into the active site of sPLA2 as discussed (17). As with the ghosts, the microscopy images supported the idea that
hydrolysis was focused at such regions of reduced fluidity (Fig. 10).
How an elevation in intracellular Ca2+ concentration would
cause this increase in membrane order is not clear. One likely
possibility is that Ca2+ ions entering the cell bind to
phospholipids, especially phosphatidylserine and phosphatidylinositols,
on the inner leaflet of the membrane. This binding would cause the
lipids to become more ordered on the inner leaflet. Increased order on
the inner face would then be likely to enhance the ordering of lipids
on the outer face leaflet since the physical properties of
phospholipids across membranes are coupled. Typical biochemical effects
of Ca2+ such as involvement of calmodulin and kinases
appear not to be involved based on results with S49 lymphoma cells (4).
The ability of phenylhydrazine to impede the effects of ionomycin on
susceptibility and membrane order was unexpected. The mechanism of this
inhibition is not yet clear. Phenylhydrazine has been reported to cause
a variety of effects on erythrocytes such as proteolysis, hemolysis,
formation of Heinz bodies, and alterations to phospholipid distribution
and dynamics (47-53). However, these effects of phenylhydrazine are
unlikely to be relevant to ionomycin-induced susceptibility and
membrane order shown in Figs. 2, 9, and 11 since they were observed
only after prolonged incubation with the agent for a period exceeding
1 h. In contrast, our results occurred immediately. We considered
possible direct effects of phenylhydrazine on sPLA2 by
monitoring the consequence of phenylhydrazine incubation on the ability
of the enzyme to hydrolyze artificial liposomes. In this case, no
inhibition by phenylhydrazine was observed. It is also unlikely that
these results reflected a direct effect of phenylhydrazine on
ionomycin. Repetition of the experiments in Fig. 5 in the presence of
phenylhydrazine demonstrated that the agent did not alter the ability
of ionomycin to induce translocation of phosphatidylserine to the
membrane exterior (not shown). Also, a different oxidizing agent,
diamide, did not interfere with the responses to ionomycin (Fig.
2).
The data in this study also support the possibility that the shape
transition from diskocytes to spherocytes induced by ionomycin (Fig. 6)
is related to susceptibility. The agents in Fig. 2 that did not block
hydrolysis by sPLA2 after addition of ionomycin also did
not alter the shape transition (based on visual inspection of the
images used to generate Fig. 11). In contrast, phenylhydrazine inhibited both. Likewise, merocyanine 540 caused both the shape transition and increased susceptibility. Nevertheless, it is doubtful that the important factor is the actual shape of the erythrocyte per se since it was shown in the accompanying article (17)
that the overall morphology of erythrocyte ghosts was unrelated to hydrolysis by sPLA2. Likewise, it is unlikely that the
decreased cell volume resulting from K+ efflux during
Ca2+ uptake (35) could be the basis of enhanced
susceptibility to sPLA2. This assertion is based on the
observation that high KCl medium, sufficient to block the reduction in
cell volume (36), failed to inhibit the vulnerability of the cells to
attack by sPLA2. It is more likely that the molecular
processes leading to the shape transition during ionomycin treatment
also promote the alterations in membrane microscopic properties that
result in enhanced hydrolysis by sPLA2.
Erythrocytes are a common model system used for studying plasma
membrane structure and its relationship with membrane proteins, cytoskeleton, and a variety of pathologies. Changes that occur upon
elevation of intracellular Ca2+ are thought to be
representative of similar changes that occur in other cells during
processes such as platelet activation and apoptosis (36, 54).
Nevertheless, the intracellular Ca2+ concentration required
for these phenomena in erythrocytes is much higher than that achieved
in other cells (1, 34-36, 54, 55). Intracellular Ca2+ in
erythrocytes treated with ionophore equilibrates rapidly and completely
with extracellular Ca2+ which is high micromolar to
millimolar in most investigations (1, 34-36, 54, 55). This raises
possible concerns regarding the relevance of observations in
erythrocytes to physiological or pathological states (36). Such
concerns are challenges one commonly faces when using a simplified
system as a model. However, the benefit of obtaining information
leading to testable hypotheses that can then be applied to more complex
systems often exceeds the disadvantages.
The results from this study offer this benefit and also potentially
relate to pathological conditions at which intracellular Ca2+ levels are very high. First, as stated above, it has
been difficult to identify in nucleated cells what changes in the
plasma membrane were involved in the induction of susceptibility due to
the diversity of membranes present in the cells. By using erythrocytes,
we have identified a testable mechanism that can now be investigated in cultured cells using imaging technology such as the two-photon method.
Second, many studies have suggested that membrane changes identified in
erythrocytes during Ca2+ loading such as PS exposure,
microvesicle release, and diminished membrane fluidity may relate to
alterations present in several erythrocyte pathologies: cell aging,
secondary effects of hypertension, spherocytosis, thalassemias, and
sickle cell disease (e.g. Refs. 36 and 56-61). For example,
sickle cells contain high levels of intracellular Ca2+ (62,
63) that may explain alterations to their membranes such as vesicle
release (60), phosphatidylserine exposure (61), and possible enhanced
susceptibility to sPLA2 in the absence of ionophore (64).
Furthermore, Ca2+-induced membrane changes in erythrocytes
are analogous to events occurring in lymphocytes during apoptosis (54).
Recent data has revealed that cells undergoing apoptosis also become
vulnerable to sPLA2 early in the apoptotic process (46).
These observations suggest that one physiological role of
sPLA2 is to help clear aging, damaged, or dying cells in
which intracellular Ca2+ levels may become very high. Such
could help explain participation of the enzyme in pathological
conditions involving damaged cells such as ischemia, sepsis, and
inflammatory disease. Based on the data of Fig. 13, it is more likely
that the group V enzyme would be involved in these processes
physiologically than the group IIa. Possible relevance of these results
to the more recently discovered group X enzyme has not yet been investigated.