Reduced Capacitative Calcium Entry Correlates with Vesicle
Accumulation and Apoptosis*
Supriya
Jayadev
,
John G.
Petranka,
Sendhil K.
Cheran,
Jennifer
A.
Biermann,
J. Carl
Barrett, and
Elizabeth
Murphy
From the Laboratory of Molecular Carcinogenesis, National Institute
of Environmental Health Sciences,
Research Triangle Park, North Carolina 27709
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ABSTRACT |
A preneoplastic variant of Syrian hamster
embryo cells, sup(+), exhibits decreased endoplasmic reticulum calcium
levels and subsequently undergoes apoptosis in low serum conditions
(Preston, G. A., Barrett, J. C., Biermann, J. A., and
Murphy, E. (1997) Cancer Res. 57, 537-542). This decrease
in endoplasmic reticulum calcium appears to be due, at least in part,
to reduced capacitative calcium entry at the plasma membrane. Thus we
investigated whether inhibition of capacitative calcium entry per
se could reduce endoplasmic reticulum calcium and induce
apoptosis of cells. We find that treatment with either SKF96365
(30-100 µM) or cell-impermeant 1,2-bis(o-amino-5-bromophenoxy)ethane-N,N,N',N'-tetraacetic
acid (5-10 mM) is able to induce apoptosis of
cells in conditions where apoptosis does not normally occur. Because
previous work has implicated vesicular trafficking as a mechanism of
regulating capacitative calcium entry, we investigated whether
disruption of vesicular trafficking could lead to decreased
capacitative calcium entry and subsequent apoptosis of cells.
Coincident with low serum-induced apoptosis, we observed an
accumulation of vesicles within the cell, suggesting deregulated
vesicle trafficking. Treatment of cells with bafilomycin (30-100
nM), an inhibitor of the endosomal proton ATPase, produced
an accumulation of vesicles, decreased capacitative entry, and induced
apoptosis. These data suggest that deregulation of vesicular transport
results in reduced capacitative calcium entry which in turn results in apoptosis.
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INTRODUCTION |
A number of recent studies have suggested that alterations in
cytoplasmic free calcium are important in apoptosis. Some groups reported that an early, sustained increase in cytoplasmic free calcium
was a necessary precedent to cell death (2-4). Others found a decrease
(5-7) or no detectable change (8-10) in cytoplasmic calcium
associated with apoptosis. In contrast to the lack of consensus on a
role for cytosolic calcium in apoptosis, a number of groups, including
our own, have recently suggested that changes in compartmentalized
calcium are important in the regulation of apoptosis (1, 6,
11-14).
Calcium within the cell exhibits a highly compartmentalized
distribution (15). Free calcium in the cytoplasm is maintained at
approximately 100 nM, whereas compartments such as the
ER1 exhibit a severalfold
higher (µM to mM) concentration of calcium (16, 17). Calcium gradients are maintained in the cell by membrane
transporters. In response to many hormones and growth factors,
transient elevations in cytoplasmic calcium occur through the opening
of channels located at the ER and plasma membranes. Cytoplasmic calcium
changes are initiated through the mobilization of signaling factors
such as inositol trisphosphate and cyclic ADP-ribose (18, 19). These
molecules act on calcium efflux channels at the ER membrane and promote
release of ER calcium stores. The subsequent depletion of ER calcium
stores is hypothesized to lead to the activation of a plasma membrane
calcium channel, the capacitative calcium entry (CCE) channel (20-22).
The ensuing influx of calcium from the extracellular milieu serves to
enhance the calcium signal and refill depleted ER stores. This coupling of ER calcium pools with capacitative entry is essential for generating transient intracellular calcium changes. The tight maintenance of
calcium gradients and the generation of transient calcium changes provides a key regulatory mechanism for cellular function (23-25).
In a previous study we investigated the role of calcium alterations
during apoptosis in two immortalized cell lines derived from
mutagenized Syrian hamster embryo cells (1). An early preneoplastic
cell line, termed sup(+), was shown to undergo a high rate of apoptosis
in low serum conditions. In contrast, a later stage preneoplastic cell
line, termed sup(
), was relatively resistant to apoptosis in low
serum. An increase in cytoplasmic free calcium was not associated with
apoptosis in this cell model; however, consistent with studies of Baffy
et al. (6) and Lam et al. (14), we found that a
decrease in ER calcium was associated with apoptosis in the sup(+)
model. The sup(
) cells, which were resistant to apoptosis in low
serum, showed no alteration in ER calcium. Pharmacological reduction of
ER calcium stores with thapsigargin resulted in apoptosis of sup(
)
cells in low serum and of sup(+) cells in 10% serum (normal growth
conditions). Conversely, maintaining ER calcium levels of sup(+) cells
in low serum prevented the DNA fragmentation characteristic of
apoptosis. Furthermore, the previous study suggested that decreased ER
calcium in apoptotic sup(+) cells was due, at least in part, to
decreased CCE leading to poor refilling of ER calcium stores (1).
In this study we sought to expand our understanding of the relationship
between mechanisms responsible for decreased calcium entry and
apoptosis. We find that preventing CCE by treating with the CCE
inhibitor SKF96365 or by chelating extracellular calcium with BAPTA
could induce apoptosis of sup(+) cells. We therefore sought to
understand the mechanism responsible for perturbation of CCE in
apoptotic sup(+) cells. Many hypotheses have been proposed for the
mechanism by which capacitative calcium entry is activated (for
reviews, see Refs. 18 and 22). Studies have suggested a role for a
diffusable second messenger (26-29), phosphorylation/dephosphorylation (27, 29-34), small G proteins (35-37), and, recently, a role for vesicular storage and regulated insertion of entry channels (38, 39).
During apoptosis we find an accumulation of vesicles within the cell,
consistent with altered vesicular trafficking. Thus we considered the
possibility that perturbation of vesicle trafficking during apoptosis
may lead to disrupted signaling to CCE. We find that perturbing vesicle
trafficking in normally growing sup(+) cells by treatment with the
macrolide antibiotic bafilomycin (Baf) results in vesicle accumulation
concomitant with decreased CCE and apoptosis. Furthermore, we find that
in the sup(
) population, which does not undergo apoptosis in reduced
serum conditions, perturbation of either CCE (with SKF96365) or vesicle
trafficking (with bafilomycin) led to apoptosis. These findings support
a role for vesicle trafficking and CCE in apoptosis.
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EXPERIMENTAL PROCEDURES |
Materials--
Fura-2 AM, BCECF-AM, and nigericin were obtained
from Molecular Probes, proteinase K from Boehringer Mannheim, and RNase
A from 5 Prime
3 Prime. All other chemicals were acquired from Sigma/Aldrich. Institute for Biological Research medium was obtained from Life Technologies Inc. and fetal calf serum was obtained from
Summit Biotechnologies.
Cell Lines and Cell Culture--
Two Syrian hamster
embryo-derived lineages, originally immortalized via mutagenesis, were
used in these studies (40, 41). The sup(+) lineage represents an early
stage of tumorigenesis which has lost a senescence gene(s) but retains
tumor suppressor capability; whereas the sup(
) lineage represents a
later stage of tumorigenesis which has lost both senescence and tumor
suppressor genes. During normal passage, sup(+) and sup(
) cells were
maintained in Dulbecco's modified Institute for Biological Research
medium containing 10% fetal calf serum, 100 units/ml penicillin, and 100 µg/ml streptomycin. Cultures were maintained in a 37 °C
incubator with 10% CO2, 90% air. For experiments, cells
were seeded at 5 × 105 cells/100-mm culture plate or
6 × 104 cells/30-mm culture plate and grown under
normal conditions for 24 h prior to treatment. Low serum
conditions were either 0.2 or 0.8% fetal calf serum. As the serum
aged, later experiments required the higher percentage of fetal calf
serum. Four treatment groups were used in the study: 1) 10% serum
(control); 2) low serum; 3) 10% + drug (e.g. SKF96365 or
bafilomycin); and 4) low serum + drug.
Calcium Measurements--
Cells were seeded at a density of
6 × 104 cells/plate in 30-mm plates containing 22-mm
diameter round glass coverslips. Following 24 h of growth, cells
were treated for 4 h. Fura-2 AM (2 µM) was introduced to each plate 30 min prior to measurements (after ~3.5 h
of treatment). Coverslips were then washed twice with calcium- and
magnesium-free phosphate-buffered saline and placed in a custom-built holder. The entire unit was placed on the stage of a Nikon inverted epifluorescence microscope coupled to a PTI Deltascan dual wavelength excitation spectrofluorometer. Measurements were taken in a window containing 2-6 cells. Cells were excited at 340 and 380 nm and emitted
fluorescence was measured at 510 nm. Background fluorescence, measured
in non-fura-2 loaded cells, was subtracted. The ratio of fluorescence
intensity was obtained by dividing the emitted fluorescence for
excitation at 340 nm by the emitted fluorescence for excitation at 380 nm. Rmin and Rmax values
were determined as described previously (1). Intracellular calcium
concentration was calculated as described previously by Grynkiewicz
et al. (42). The cytosolic location of fura-2 was confirmed
for each treatment condition using saponin (43). Saponin was able to
release approximately 80-85% of loaded dye in all conditions.
ER and capacitative calcium entry were measured as follows: fura-2
loaded cells were treated with the ER calcium ATPase inhibitor thapsigargin (2 µM) in the absence of extracellular
calcium. Thapsigargin stimulates the release of ER calcium into the
cytoplasm thus permitting measurement of the stored ER calcium pool.
Following pool emptying, calcium is removed from the cytoplasm
presumably by extrusion via the plasma membrane transporters. Once
basal cytosolic readings were restored, extracellular calcium was
raised to 1 mM and the rapid increase in cytoplasmic
calcium (induced by ER store depletion) was measured as peak
fluorescence with fura-2.
DNA Fragmentation Analyses--
Cells were seeded at a density
of 5 × 105 cells/plate in 100-mm plates and grown for
24 h. Cells were washed with calcium- and magnesium-free
phosphate-buffered saline and 10 ml of the appropriate treatment media
was added. Following 16-24 h (see figure legends) cells were scraped
into the treatment media and pelleted. The cell pellets were washed
with calcium- and magnesium-free phosphate-buffered saline, pelleted,
and lysed in 50-100 µl of lysis buffer (10 mM EDTA, 50 mM Tris, pH 8, 0.5% sodium lauryl sarcosine, 0.5 mg/ml
proteinase K). DNA was extracted and analyzed for fragmentation as
described previously (1).
pH Measurements--
Cells were seeded at a density of 6 × 104 cells/plate in 30-mm plates containing 22-mm diameter
round glass coverslips. Following 24 h of growth, cells were
washed and treated for 4 h. BCECF-AM (4 µM) was
introduced to each plate 15 min prior to measurements. Coverslips were
then washed twice with media containing 20 mM HEPES, pH
7.4, and placed in a custom-built holder. One ml of media containing 20 mM HEPES, pH 7.4, was added to the chamber and the entire
unit was placed on the stage of a Nikon inverted epifluorescence
microscope coupled to a PTI Deltascan dual wavelength excitation
spectrofluorometer. Measurements were taken in a window containing 2-6
cells. Emission was measured at 535 nm following excitation at 439 and
505 nm. Background fluorescence, measured in an area containing no
cells, was subtracted. The ratio of fluorescence intensity was obtained
by dividing the emitted fluorescence for excitation at 505 nm by
the emitted fluorescence for excitation at 439 nm. To calibrate dye
fluorescence for each sample, the ratio of fluorescence was measured at
4-5 pH values using the high KCl-nigericin technique (44, 45).
Electron Microscopy--
Cells were seeded in 100-mm plates at a
density of 5 × 105 cells/plate and grown for 24 h. Cells were washed with calcium- and magnesium-free
phosphate-buffered saline and 5 ml of the appropriate treatment media
was added. Following 4-16 h (see figure legends), 5 ml of 2 × fixation buffer (4% glutaraldehyde, 0.2 M sodium
cacodylate, pH 7.2) was added to the plates. Cells were scraped into
the media/fix mixture and pelleted. Cell pellets were re-suspended in 1 ml of 1 × fixative and incubated at 4 °C for 2 h. Fixed
cells were washed with 0.1 M sodium cacodylate, pH 7.2, and
maintained in fresh buffer at 4 °C until further processing was
done. Samples were post-fixed in 1% osmium tetroxide, dehydrated in
graded ethanol and embedded in Poly Bed 812. The samples were cured for
3 days at 60 °C and thin sections were cut. Sections were stained
with 5% uranyl acetate followed by Reynolds lead citrate solution and then examined using a Phillips 400 transmission electron microscope.
Statistical Analyses--
Values are mean ± S.E. In
comparing two groups, statistical significance was determined by
Student's t test. For multiparameter comparisons,
statistical significance was determined by ANOVA, adjusting for
multiple comparisons using Fisher's test for significance. A value of
p < 0.05 was considered to be significant.
 |
RESULTS |
Capacitative Calcium Entry and Apoptosis in Sup(+)
Cells--
Sup(+) cells, an immortalized cell line derived from
mutagenesis of Syrian hamster embryo cells (40), undergo apoptosis in
response to low serum conditions (41). In a previous study, a decrease
in ER calcium was shown to precede apoptosis in sup(+) cells (1). The
decrease in ER calcium was, in turn, found to be secondary to a
decrease in the refilling pathway of CCE (1). Therefore, we were
interested in testing whether inhibition of CCE per se could
induce apoptosis of cells under conditions where apoptosis does not
normally occur (in 10% serum). We found a dose-dependent inhibition of CCE in sup(+) cells following treatment with SKF96365 (Fig. 1), an inhibitor of CCE and other
calcium channels. As little as 30 µM SKF96365 was able to
reduce CCE approximately 30%. Higher doses of SKF96365 decreased CCE
even more significantly, from the peak control calcium entry of
1.5 ± 0.9 to 0.15 ± 0.1 µM with 100 µM SKF96365 (Fig. 1). In conjunction with CCE, we also
measured ER calcium as the thapsigargin releasable calcium as described under "Experimental Procedures." Sup(+) cells treated with SKF96365 exhibited a 50% decrease in ER calcium within 4 h following
treatment. In control cells (10% serum) thapsigargin released 116 ± 15 nM ER calcium into the cytosol versus
42 ± 11 nM in SKF96365 (100 µM) treated
cells.

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Fig. 1.
SKF96365 causes a dose-dependent
inhibition of capacitative calcium entry. Sup(+) cells were
treated for 4 h with 0, 30, 50, or 100 µM SKF96365
in 10% serum media. During the final 30 min of treatment cells were
also loaded with fura-2 (2 µM). The 100 µM
SKF96365 (n = 6) treatment showed a significant
decrease (p < 0.01) in CCE compared with control,
untreated cells (n = 5). The 30 µM
(n = 2) and 50 µM (n = 3)
SKF96365 treatments also follow a trend of decreased CCE compared with
controls.
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The same doses of SKF96365 which effected CCE were also able to induce
apoptosis (Fig. 2). Within 16 h of
treatment with 30 µM SKF96365, sup(+) cells grown in 10%
serum showed the classic DNA laddering pattern indicative of apoptosis
(lane 2). Similar to the dose-dependent
inhibition of CCE, higher doses of SKF96365 were able to induce more
prominent laddering of sup(+) cells (lanes 3 and
4). Since SKF96365 is a non-selective inhibitor of CCE, we
further substantiated the relationship between CCE and apoptosis using
cell-impermeant BAPTA to chelate extracellular calcium. BAPTA chelates
calcium with one to one binding, therefore millimolar concentrations of
BAPTA are required to chelate the 1.8 mM calcium concentration present in the treatment media. Thus, lower doses of
BAPTA were unable to reduce extracellular calcium beyond the high
micromolar range (e.g. 1 mM BAPTA only decreased
the calcium concentration to ~800 µM), and
correspondingly were unable to induce apoptosis (Fig.
3, lanes 2-5). In contrast,
doses of BAPTA which significantly chelated extracellular calcium (5 mM BAPTA reduced the calcium concentration to ~390
nM and 10 mM BAPTA reduced the calcium
concentration to ~150 nM), were able to induce DNA ladders in sup(+) cells (Fig. 3, lanes 6 and 7,
for comparison, lane 8 shows low serum-induced
apoptosis).

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Fig. 2.
SKF96365 induces apoptosis of sup(+) cells in
10% serum. Sup(+) cells were treated for 16 h with 0, 30, 50, or 100 µM SKF96365 in 10% serum media. A
representative agarose gel of extracted DNA is shown. Lane
1, no addition; lane 2, 30 µM SKF96365;
lane 3, 50 µM SKF96365; lane 4, 100 µM SKF96365. All three treatments with SKF96365 produced
DNA fragmentation ladders which are indicative of apoptosis; in
contrast, control cells showed no laddering.
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Fig. 3.
BAPTA induces apoptosis of sup(+) cells in
10% serum. Sup(+) cells were treated for 16 h with 0.01-10
mM cell-impermeant BAPTA in 10% serum media. A
representative agarose gel of extracted DNA is shown. Lane
1, 10% serum control; lane 2, 0.01 mM
BAPTA; lane 3, 0.1 mM BAPTA; lane 4, 0.5 mM BAPTA; lane 5, 1 mM BAPTA;
lane 6, 5 mM BAPTA; lane 7, 10 mM BAPTA; lane 8, low serum. BAPTA
concentrations of 5 and 10 mM, which reduced free
extracellular calcium concentrations to 390 and 150 nM,
respectively, were able to induce laddering of sup(+) cells similar to
low serum treatment. Lower concentrations of BAPTA, which did not
significantly chelate extracellular calcium (e.g. 1 mM BAPTA only reduced calcium to 800 µM),
were not able to cause laddering.
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Capacitative Entry and Vesicle Accumulation in Sup(+)
Cells--
Many hypotheses have been proposed for the mechanism
responsible for CCE (for reviews, see Refs. 18, 22, 37, and 46). One
model which has been suggested is the control of calcium entry through
regulated insertion of CCE channels (38, 39). In this model, signals
from a store-depleted ER would "activate" an endomembrane compartment enabling it to fuse with the plasma membrane, inserting more entry channels into the plasma membrane. A prediction of this
model would be that inhibiting vesicle movement to and/or fusion with
the plasma membrane would inhibit CCE. Intriguingly, in many
morphological discussions of cell death, the apoptotic process is
associated with accumulation of vesicles within the cell (47-52).
Similar to many other systems that undergo apoptosis, we found that
apoptotic sup(+) cells accumulate vesicles in the cytoplasm. Sup(+)
cells maintained in low serum conditions for 16 h show a marked
increase in the number of vesicles present per cell as well as an
increase in the number of cells with vesicles (compare Fig.
4A (10%, 16 h) to Fig.
4B (low serum, 16 h)). The increase in vesiculation in
low serum-treated sup(+) cells occurs early, within 4 h an
increase in vesicles is readily apparent (Fig. 4C), preceding other known morphological markers of apoptosis such as
nuclear and cytoplasmic compaction (49, 51).

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Fig. 4.
Electron micrographs of sup(+) cells
undergoing apoptosis. Sup(+) cells were treated for 4 h
(panels C, F and G) or 16 h (panels A,
B, D, and E) with: panel A, 10% serum;
panels B and C, low serum; panel D,
10% serum + 100 µM SKF96365; panels E and
F, 10% serum + 50 nM Baf; panel G,
low serum + 50 nM Baf. Control cells (A) show
"normal" morphology of elongated cells containing intact organelles
and few vesicles. Under low serum conditions (B and
C) and with SKF96365 (D) or Baf (E-G)
treatment, cells exhibit the classic morphological changes which
accompany apoptosis: nuclear condensation, rounded morphology, and
decreased cell size. Only with low serum or Baf treatment is increased
vesiculation observed (B, C, and E-G). The
increase in vesicle accumulation occurs early, preceding many of the
other markers of apoptosis (C and F). Combined
treatment with low serum and Baf appears to speed up the apoptotic
process such that by 4 h apoptotic morphology can be seen in the
cell population (G). Magnifications: A, × 6,200;
B, × 14,190; C, × 8,580; D, × 11,000; E, × 14,190; F, ×6,200; G, × 11,000.
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It is interesting to note that not all apoptotic treatments lead to
vesicle accumulation. Whereas low serum treatment can lead to vesicle
accumulation (Fig. 4, B and C) and decreased CCE (Ref. 1 and Fig. 7), SKF96365 treatment only decreases CCE (Fig. 1)
without affecting vesicle accumulation (Fig. 4D). This suggests that the sequence of events is one in which vesicle
accumulation precedes decreased CCE. Thus, pharmacological alteration
of CCE (e.g. with SKF96365) would not necessarily require
the involvement of vesicles.
Bafilomycin Increases Vesiculation and Induces Apoptosis of Sup(+)
Cells--
If vesicle accumulation is involved in the regulation of
apoptosis as opposed to a result of apoptosis, then pharmacological inhibition of vesicular trafficking should induce apoptosis.
Bafilomycin A1 (Baf), a potent selective inhibitor of
vacuolar H+-ATPase (53, 54) has previously been shown to
inhibit late Golgi and post-Golgi trafficking of vesicles to the plasma
membrane (55-57). We find that treatment of sup(+) cells in 10% serum
with 50 nM Baf for 16 h resulted in the accumulation
of large vacuoles within the cell (Fig. 4E). As with low
serum (in the absence of Baf), with Baf treatment vesiculation was
apparent early (within 4 h, Fig. 4F), preceding other
morphological markers of apoptosis such as nuclear/cytoplasmic
compaction, rounding up of cells, and decreased cell size, apparent by
16 h of treatment (Fig. 4E). Interestingly, treating
sup(+) cells with Baf in low serum served to speed up apoptosis, such
that morphological changes such as nuclear and cytoplasmic condensation
were seen as early as 4 h following treatment (Fig.
4G).
We next evaluated whether the increase in vesiculation induced by Baf
resulted in induction of apoptosis. As with SKF96365 treatment, Baf
treatment caused apoptosis in sup(+) cells under conditions where
apoptosis does not normally occur, e.g. in 10% serum. As
shown in Fig. 5, 16 h of treatment
with 50 nM Baf in 10% serum (lane 2),
conditions in which Baf induced intracellular vesicle accumulation,
resulted in apoptosis similar to that observed with low serum treatment
(lane 3).

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Fig. 5.
Bafilomycin induces apoptosis in sup(+) cells
in 10% serum. Sup(+) cells were treated for 16 h with 10%
serum (lane 1), 10% serum + 50 nM Baf
(lane 2), or low serum (lane 3). A representative
agarose gel of extracted DNA is shown. Both low serum treatment and Baf
treatment resulted in DNA fragmentation.
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Bafilomycin Does Not Affect Cytoplasmic pH of Sup(+)
Cells--
Baf acts to dissipate endosomal pH gradients and in some
cells this perturbation of vesicular pH causes a change in the
cytoplasmic pH (58). In addition, there have been reports that a
decrease in cytoplasmic pH is involved in initiating apoptosis
(59-62). To ensure that the results with Baf were due to changes in
vesiculation and not to changes in pH, we investigated whether changes
in cytoplasmic pH occur with the addition of Baf or during low
serum-induced apoptosis in sup(+) cells. By 4 h of treatment with
Baf, when vesicle accumulation was evident, there was no significant
effect on cytoplasmic pH of sup(+) cells (Fig.
6). Sup(+) cells grown in 10% serum had
a pH of 7.43 ± 0.04 and cells grown in 10% serum plus Baf had a
pH of 7.46 ± 0.02. Similar to Baf treatment, low serum did not
significantly change the cytoplasmic pH of sup(+) cells. Thus, we
conclude that Baf induces apoptosis and vesiculation without effecting
cytoplasmic pH.

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Fig. 6.
Bafilomycin does not alter cytoplasmic
pH. Sup(+) cells were treated for 4 h with 10% serum, low
serum, or 10% serum + 50 nM Baf. During the final 15 min
of treatment cells were loaded with BCECF. Values are means ± S.E. from five to nine separate experiments. Neither low serum
(7.39 ± 0.1) nor Baf treatment (7.46 ± 0.02) produced
significant changes in cytoplasmic pH compared with control levels
(7.43 ± 0.04).
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Bafilomycin Decreases Capacitative Entry--
We next examined
whether Baf treatment causes a decrease in CCE. If CCE is regulated by
insertion of calcium entry channels into the plasma membrane, we would
expect that perturbing vesicular insertion into the plasma membrane by
addition of Baf would decrease CCE. Typical traces of CCE measurements
are shown in Fig. 7A. In 10%
serum CCE is markedly higher than in low serum or Baf-treated cells.
Fig. 7B shows similar data summing up observations from 8 to
11 separate experiments. At 4 h of Baf treatment in 10% serum, CCE is reduced approximately 30% compared with control, 10% serum levels (similar to the reduction seen with low serum treatment). Interestingly, Baf treatment in low serum conditions did not produce an
additive effect on CCE at 4 h (Fig. 7B). Although
co-treatment may have sped up the apoptotic process (Fig.
4G) the non-additive effect of Baf and low serum on CCE at
4 h suggests that early events which lead to apoptosis may be the
same with both treatments.

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Fig. 7.
Bafilomycin inhibits capacitative calcium
entry. Sup(+) cells were treated as indicated for 4 h. During
the final 30 min of treatment cells were also loaded with 2 µM fura-2. Data are presented as 340/380 ratios.
A, a representative trace is shown for: 1, 10%
serum; 2, low serum; 3, 50 nM Baf + 10% serum; B, data from eight to 11 separate measurements
were averaged and averages ± S.E. are shown. All three treatment
groups (low serum, 10% + 50 nM Baf, and low serum + 50 nM Baf) showed a significant decrease (p < 0.03) in CCE compared with control, untreated cells as assessed by
analysis of variance for repeated measurements followed by the
Fisher's test.
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Since Baf is known to inhibit the vacuolar proton ATPase within
minutes, we also investigated the effect of Baf on CCE at early time
points. Acute treatment with Baf had no effect on CCE and even
following 30 min of treatment, Baf had no significant effect on CCE
(remaining at 82% ± 36 of control (n = 6)). It was only after prolonged treatment with Baf that reduced CCE could be
observed, with 2 h of treatment reducing CCE by 45%
(n = 4) and 4 h treatment reducing CCE by 32%.
Vesiculation, CCE, and Apoptosis in Sup(
) Cells--
Finally, if
vesicle trafficking and CCE are involved in the apoptotic process in
general, then the same relationships should hold true in other cell
systems. Thus, to further substantiate our findings we expanded our
study to sup(
) cells. Sup(
) cells do not exhibit decreased ER
calcium or undergo apoptosis under low serum conditions; however, our
previous study established that pharmacological reduction of ER calcium
during low serum treatment of sup(
) cells resulted in apoptosis (1).
Thus, we investigated whether decreasing CCE or altering vesicle
trafficking could induce apoptosis in sup(
) cells. We found that
sup(
) cells treated with SKF96365 showed decreased CCE and underwent
apoptosis (Fig. 8). Unlike sup(+) cells,
sup(
) cells did not show decreased CCE with low serum treatment alone
(Fig. 8A). Consistently, sup(
) cells treated with low
serum did not exhibit the classic laddering pattern indicative of
apoptosis but instead showed a smeared DNA pattern, indicative of
necrotic death (Fig. 8B, lane 2). In contrast, sup(
) cells
treated with SKF96365 in conjunction with low serum showed both
decreased CCE (Fig. 8A) and apoptosis (Fig. 8B, lanes 3-6). As with sup(+) cells, reduction in CCE temporally preceded apoptosis, with over an 80% decrease in CCE apparent within 4 h
and DNA ladders apparent only after 16 h.

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Fig. 8.
SKF96365 inhibits capacitative calcium entry
and induces apoptosis of sup( ) cells. Panel A,
sup( ) cells were treated for 4 h with 10% serum
(n = 7), low serum (n = 6), or low
serum + 50 µM SKF96365 (n = 3). During
the final 30 min of treatment cells were also loaded with 2 µM fura 2-AM. Data are presented as 340/380 ratios ± S.E. Unlike the sup(+) cell line, sup( ) cells do not exhibit a
significant decrease in CCE with low serum treatment. However, combined
treatment with SKF96365 and low serum results in a significant
(p < 0.01 versus low serum or 10%
treatment) inhibition of CCE. Panel B, sup( ) cells were
treated for 16 h with: lane 1, 10% serum; lane
2, low serum; lane 3, low serum + 1 µM
SKF96365; lane 4, low serum + 5 µM SKF96365;
lane 5, low serum + 10 µM SKF96365; and
lane 6, low serum + 50 µM SKF96365. Low serum
shows a smeared pattern of DNA indicative of necrotic death of the
sup( ) population. In contrast, SKF96365 treatment shows laddering,
indicative of apoptotic death, higher doses show more prominent ladders
with 50 µM exhibiting the most prominent ladders.
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Since we hypothesize that vesicle trafficking affects CCE and in turn,
apoptosis, we next determined the effect of perturbing vesicle
trafficking in sup(
) cells. As with the sup(+) population, sup(
)
cells maintained in 10% serum conditions show a minimal number of
vesicles present within the cell (Fig.
9a). Sup(
) cells grown in
low serum did not exhibit an accumulation of vesicles (Fig.
9b), consistent with our suggestion that aberrant vesicle trafficking is part of (and is upstream of) the signal for decreased CCE and apoptosis. Since sup(
) cells do not exhibit reduced CCE or
undergo apoptosis in low serum, they should not show vesicle accumulation in this condition. Conversely, we would hypothesize that
altering vesicle trafficking with an agent such as Baf should result in
the same effects as was observed in the sup(+) population, decreased
CCE and apoptosis. Indeed we find that Baf treatment of sup(
) cells
in low serum results in a marked accumulation of vesicles (Fig.
9c) and approximately 60% reduction in CCE within 4 h
(Fig. 10A). Furthermore, Baf
treatment of sup(
) cells was able to change necrotic death (evidenced
in low serum as smeared DNA, Fig. 10B, lane 1) to apoptotic
death (evidenced by the classic DNA ladders, Fig. 10B, lane
2).

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|
Fig. 9.
Vesicle accumulation in sup( ) cells.
Sup( ) cells were treated for 4 h with: a, 10% serum;
b, low serum; c, low serum + 50 nM
Baf. Control cells (a) show "normal" morphology of
elongated cells containing few, if any, vesicles. In contrast to sup(+)
cells, low serum-treated sup( ) cells (b) show little
difference from the control population, cells remain elongated and
contain few vesicles. Combined treatment with Baf and low serum,
however, results in a significant accumulation of vesicles.
|
|

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[in a new window]
|
Fig. 10.
Bafilomycin inhibits capacitative calcium
entry and induces apoptosis of sup( ) cells. Panel A,
sup( ) cells were treated for 4 h with low serum
(n = 6) or low serum + 50 nM Baf
(n = 5). During the final 30 min of treatment cells
were also loaded with 2 µM fura 2-AM. Data are presented
as 340/380 ratios ± S.E. Baf treatment significantly
(p < 0.03) decreased CCE. Panel B, sup( )
cells were treated for 16 h with: lane 1, low serum; or
lane 2, low serum + 50 nM Baf. Low serum shows
the smeared pattern of DNA indicative of necrotic death of the sup( )
population whereas Baf treatment shows the classic laddering indicative
of apoptotic death.
|
|
 |
DISCUSSION |
Calcium regulation of apoptosis has been studied in a myriad of
systems (63-65). As more studies have focused on calcium and apoptosis, there has been increasing controversy as to the exact role
that calcium plays. Our work and that of others, have suggested that
decreased ER calcium stores are important in activation of apoptosis
(1, 6, 12, 13). We have shown previously that decreased ER calcium
correlated with decreased CCE (1). However, it was not clear from the
previous study whether the decrease in CCE was a cause or an effect of
apoptosis. To address this question, we inhibited CCE by addition of
SKF96365, a nonspecific inhibitor of the CCE channel (66). We find in
the sup(+) and sup(
) cell lines that SKF96365 inhibits CCE within
4 h of treatment. We therefore examined whether SKF96365
inhibition of CCE per se could induce apoptosis. As shown in
Figs. 2 and 8, at doses which inhibit CCE, SKF96365 also induces
apoptosis. Since SKF96365 is reported to inhibit other calcium entry
pathways (66), we confirmed the interconnection between CCE and
apoptosis using an extracellular calcium chelator-BAPTA. Similar to low
serum or SKF96365 treatment, chelating extracellular calcium with
cell-impermeant BAPTA resulted in DNA fragmentation. Thus, inhibiting
the ER refilling pathway seems sufficient to signal apoptosis of sup(+)
and sup(
) cells.
The signal connecting a decrease in ER calcium to CCE has been studied
by a number of investigators and continues to be a very active area of
investigation (22, 46, 67, 68). At this point the precise mechanism of
CCE regulation remains largely unknown, despite intense investigation.
It has been reported that a soluble messenger is responsible for
stimulating CCE (26-29), although others have questioned this
mechanism (37). Alterations in phosphorylation have also been proposed
to be important in CCE (27, 29-34), although again there are data to
the contrary (22). One consistent finding across laboratories is that
addition of GTP
S or GDP
S can inhibit CCE (35, 36, 69, 70). The heterotrimeric G proteins and most small G proteins are activated by
GTP
S and inhibited by GDP
S (71). Inhibition by both GTP
S and
GDP
S is consistent with the involvement of the Rab proteins (71-73), which are small G proteins important in vesicle trafficking (72, 73). Further support suggesting that vesicle transport is involved
in the regulation of CCE has come from: 1) studies showing that CCE
exhibits cold sensitivity (39), and, 2) studies showing that CCE is
inhibited by the lysosomotropic inhibitor primaquine (38). It is
intriguing to speculate that perhaps CCE is regulated in a manner
similar to GLUT4 regulation. In the GLUT4 model, insulin stimulation of
cells leads to the fusion of vesicles containing the GLUT4 transporter
with the plasma membrane (74-76). Thus, increased glucose uptake
results from increased transporters at the plasma membrane. Whether
regulated insertion of CCE channels represents an important and
universal mechanism for modulating calcium entry following store
depletion remains to be determined. Nonetheless, studies from a number
of groups as well as the current study suggest a role for vesicle
trafficking and insertion of CCE channels as a means of modulating
CCE.
In light of a possible role for vesicle trafficking in CCE, we were
struck by the accumulation of vesicles that occurs in sup(+) and other
cells undergoing apoptosis. Indeed, in many morphological discussions
of apoptosis, accumulation of vesicles has been defined as one of the
hallmarks of apoptosis (47-52). The accumulation of vesicles in early
apoptosis is consistent with altered vesicle trafficking and might be
related to inhibition of CCE. To test further this hypothesis, we
inhibited vesicular transport by addition of bafilomycin. Bafilomycin,
a known specific inhibitor of the vacuolar H+-ATPase (53,
54), has been previously shown to inhibit late Golgi and post-Golgi
trafficking of vesicles to the plasma membrane (55-57). As expected
from such an effect, Baf was able to induce the accumulation of
vesicles within the cytoplasm of sup(+) and sup(
) cells. More
importantly, we found that Baf treatment of sup(+) cells in 10% serum
and sup(
) cells in low serum, significantly decreased CCE and induced
apoptosis of cells. The current study not only establishes a connection
between vesicle trafficking and CCE but also between these two events
and apoptosis. Although a number of studies have suggested a connection
between CCE and vesicle trafficking, this is the first study to address
the interconnection of these events with apoptosis.
Mechanistically, it appears that the means by which signaling to CCE
becomes disrupted during apoptosis involves vesicular trafficking. One
of the classical morphological characteristics which defines apoptosis
is vesicle accumulation near the plasma membrane (47-52). In the
sup(+) population undergoing apoptosis we also observe vesicle
accumulation. Furthermore, in cells exhibiting vesicle accumulation
associated with low serum or Baf, we observe decreased capacitative
entry and apoptosis. The converse does not appear to be true, however,
cells treated with SKF96365 have decreased CCE but they do not show
accumulation of vesicles (Fig. 4D). These findings serve to
order events into a scheme whereby vesicle accumulation precedes
aberrant CCE and decreased CCE precedes apoptosis. Thus, as shown with
SKF96365, inhibition of CCE entry without affecting vesicular
trafficking appears to be sufficient to induce apoptosis of cells. The
universality of this mechanism of regulating CCE and apoptosis remains
to be determined.
 |
ACKNOWLEDGEMENTS |
We thank John L. Horton for assistance with
EM work and Dr. Francis M. Hughes Jr. and Dr. Gary St. John Bird for
careful review of the manuscript.
 |
FOOTNOTES |
*
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement" in
accordance with 18 U.S.C. Section
1734 solely to indicate this fact.
To whom correspondence and reprints should be addressed: NIEHS,
P.O. Box 12233, MD D2-03, Research Triangle Park, NC 27709. Tel.:
919-541-4976; Fax: 919-541-3385; E-mail:
jayadev{at}niehs.nih.gov.
 |
ABBREVIATIONS |
The abbreviations used are:
ER, endoplasmic
reticulum;
CCE, capacitative calcium entry;
Baf, bafilomycin;
GLUT4, glucose transporter type 4;
BCECF-AM, 2',7'-bis-(carboxyethyl)-5(6')-carboxyfluorescein acetoxymethyl ester;
GTP
S, guanosine 5'-O-(3-thiotriphosphate);
GDP
S, guanyl-5'-yl thiophosphate;
BAPTA, 1,2-bis(o-amino-5-bromophenoxy)ethane-N,N,N',N'-tetraacetic
acid.
 |
REFERENCES |
-
Preston, G. A.,
Barrett, J. C.,
Biermann, J. A.,
and Murphy, E.
(1997)
Cancer Res.
57,
537-542[Abstract]
-
McConkey, D. J.,
Nicotera, P.,
Hartzell, P.,
Bellomo, G.,
Wyllie, A. H.,
and Orrenius, S.
(1989)
Arch. Biochem. Biophys.
269,
365-370[Medline]
[Order article via Infotrieve]
-
McConkey, D. J.,
Aguilar-Santelises, M.,
Hartzell, P.,
Eriksson, I.,
Mellstedt, H.,
Orrenius, S.,
and Jondal, M.
(1991)
J. Immunol.
146,
1072-1076[Abstract/Free Full Text]
-
Bellomo, G.,
Perotti, M.,
Taddei, F.,
Mirabelli, F.,
Finardi, G.,
Nicotera, P.,
and Orrenius, S.
(1992)
Cancer Res.
52,
1342-1346[Abstract]
-
Magnelli, L.,
Cinelli, M.,
Turchetti, A.,
and Chiarugi, V. P.
(1993)
Biochem. Biophys. Res. Commun.
194,
1394-1397[CrossRef][Medline]
[Order article via Infotrieve]
-
Baffy, G.,
Miyashita, T.,
Williamson, J. R.,
and Reed, J. C.
(1993)
J. Biol. Chem.
268,
6511-6519[Abstract/Free Full Text]
-
Magnelli, L.,
Cinelli, M.,
Turchetti, A.,
and Chiarugi, V. P.
(1994)
Biochem. Biophys. Res. Commun.
204,
84-90[CrossRef][Medline]
[Order article via Infotrieve]
-
Iseki, R.,
Kudo, Y.,
and Iwata, M.
(1993)
J. Immunol.
151,
5198-5207[Abstract/Free Full Text]
-
Zhu, W.-H.,
and Loh, T.-T.
(1995)
Life Sci.
57,
2091-2099[CrossRef][Medline]
[Order article via Infotrieve]
-
Beaver, J. P.,
and Waring, P.
(1994)
Immunol. Cell Biol.
72,
489-499[Medline]
[Order article via Infotrieve]
-
Lam, M.,
Dubyak, G.,
Chen, L.,
Nunez, G.,
Miesfeld, R. L.,
and Distelhorst, C. W.
(1994)
Proc. Natl. Acad. Sci. U. S. A.
91,
6569-6573[Abstract]
-
Bian, X.,
Hughes, F. M., Jr.,
Huang, Y.,
Cidlowski, J. A.,
and Putney, J. W., Jr.
(1997)
Am. J. Physiol.
272,
C1241-C1249[Abstract/Free Full Text]
-
Richter, C.
(1993)
FEBS Lett.
325,
104-107[CrossRef][Medline]
[Order article via Infotrieve]
-
Lam, M.,
Dubyak, G.,
and Distelhorst, C. W.
(1993)
Mol. Endocrinol.
7,
686-693[Abstract]
-
Pozzan, T.,
Rizzuto, R.,
Volpe, P.,
and Meldolesi, J.
(1994)
Physiol. Rev.
74,
595-636[Free Full Text]
-
Meldolesi, J.,
and Pozzan, T.
(1998)
Trends Biochem. Sci.
23,
10-14[CrossRef][Medline]
[Order article via Infotrieve]
-
Chen, W.,
Steenbergen, C.,
Levy, L. A.,
Vance, J.,
London, R. E.,
and Murphy, E.
(1996)
J. Biol. Chem.
271,
7398-7403[Abstract/Free Full Text]
-
Murphy, C. T.,
Poll, C. T.,
and Westwick, J.
(1995)
Cell Calcium
18,
245-251[Medline]
[Order article via Infotrieve]
-
Marks, A. R.
(1997)
Am. J. Physiol.
272,
H597-H605[Abstract/Free Full Text]
-
Putney, J. W., Jr.
(1986)
Cell Calcium
7,
1-12[Medline]
[Order article via Infotrieve]
-
Fasolato, C.,
Innocenti, B.,
and Pozzan, T.
(1994)
Trends Biochem. Sci.
15,
77-83
-
Berridge, M. J.
(1995)
Biochem. J.
312,
1-11[Medline]
[Order article via Infotrieve]
-
Berridge, M. J.
(1995)
BioEssays
17,
491-500[Medline]
[Order article via Infotrieve]
-
Sargeant, P.,
and Sage, S. O.
(1994)
Pharmacol. & Ther.
64,
395-443[CrossRef][Medline]
[Order article via Infotrieve]
-
Whitfield, J. F.,
Bird, R. P.,
Chakravarthy, B. R.,
Isaacs, R. J.,
and Morley, P.
(1995)
J. Cell. Biochem.
22,
74-91
-
Randriamampita, C.,
and Tsien, R. Y.
(1993)
Nature
364,
809-814[CrossRef][Medline]
[Order article via Infotrieve]
-
Parekh, A. B.,
Terlau, H.,
and Stuhmer, W.
(1993)
Nature
364,
814-818[CrossRef][Medline]
[Order article via Infotrieve]
-
Davies, E. V.,
and Hallett, M. B.
(1995)
Biochem. Biophys. Res. Commun.
206,
348-354[CrossRef][Medline]
[Order article via Infotrieve]
-
Thomas, D.,
and Hanley, M. R.
(1995)
J. Biol. Chem.
270,
6429-6432[Abstract/Free Full Text]
-
Randriamampita, C.,
and Tsien, R. Y.
(1995)
J. Biol. Chem.
270,
29-32[Abstract/Free Full Text]
-
Montero, M.,
Garcia-Sancho, J.,
and Alvarez, J.
(1993)
J. Biol. Chem.
268,
13055-13061[Abstract/Free Full Text]
-
Hardie, R. C.,
Peretz, A.,
Suss-Toby, E.,
Rom-Glas, A.,
Bishop, S. A.,
Selinger, Z.,
and Minke, B.
(1993)
Nature
363,
634-637[Medline]
[Order article via Infotrieve]
-
Sargeant, P.,
Farndale, R. W.,
and Sage, S. O.
(1994)
Exp. Physiol.
79,
269-272[Abstract]
-
Parekh, A. B.,
and Penner, R.
(1995)
Proc. Natl. Acad. Sci. U. S. A.
92,
7907-7911[Abstract]
-
Bird, G. S.,
and Putney, J. W., Jr.
(1993)
J. Biol. Chem.
268,
21486-21488[Abstract/Free Full Text]
-
Fasolato, C.,
Hoth, M.,
and Penner, R.
(1993)
J. Biol. Chem.
268,
20737-20740[Abstract/Free Full Text]
-
Putney, J. W., Jr.,
and Bird, G. S.
(1993)
Cell
75,
199-201[Medline]
[Order article via Infotrieve]
-
Somasundaram, B.,
Norman, J. C.,
and Mahaut-Smith, M. P.
(1995)
Biochem. J.
309,
725-729[Medline]
[Order article via Infotrieve]
-
Somasundaram, B.,
Mahaut-Smith, M. P.,
and Floto, R. A.
(1996)
J. Biol. Chem.
271,
26096-26104[Abstract/Free Full Text]
-
Koi, M.,
and Barrett, J. C.
(1986)
Proc. Natl. Acad. Sci. U. S. A.
83,
5992-5996[Abstract]
-
Preston, G. A.,
Lang, J. E.,
Maronpot, R. R.,
and Barrett, J. C.
(1994)
Cancer Res.
54,
4214-4223[Abstract]
-
Grynkiewicz, G.,
Poenie, M.,
and Tsien, R. Y.
(1985)
J. Biol. Chem.
260,
3440-3450[Abstract]
-
Murphy, E.,
Freudenrich, C. C.,
Levy, L. A.,
London, R. E.,
and Lieberman, M.
(1989)
Proc. Natl. Acad. Sci. U. S. A.
86,
2981-2984[Abstract]
-
Shrode, L. D.,
Klein, J. D.,
Douglas, P. B.,
O'Neill, W. C.,
and Putnam, R. W.
(1997)
Am. J. Physiol.
272,
C1968-C1979[Abstract/Free Full Text]
-
Thomas, J. A.,
Buchsbaum, R. N.,
Zimniak, A.,
and Racker, E.
(1979)
Biochemistry
18,
2210-2218[Medline]
[Order article via Infotrieve]
-
Parekh, A. B.,
and Penner, R.
(1997)
Physiol. Rev.
77,
901-930[Abstract/Free Full Text]
-
Wyllie, A. H.,
Kerr, J. F. R.,
and Currie, A. R.
(1980)
Int. Rev. Cytol.
68,
251-306[Medline]
[Order article via Infotrieve]
-
Wyllie, A. H.,
Morris, R. G.,
Smith, A. L.,
and Dunlop, D.
(1984)
J. Pathol.
142,
67-77[Medline]
[Order article via Infotrieve]
-
Wyllie, A. H.
(1997)
Int. Rev. Cytol.
1987,
755-785
-
Munker, R.,
Greither, L.,
Darsow, M.,
Pander, S.,
and Wilmanns, W.
(1992)
Ann. Hematol.
65,
50-52[Medline]
[Order article via Infotrieve]
-
Vollenweider, I.,
and Groscurth, P.
(1991)
Electron Microsc. Rev.
4,
249-267[Medline]
[Order article via Infotrieve]
-
Payne, C. M.,
Bernstein, C.,
and Bernstein, H.
(1995)
Leuk. & Lymphoma
19,
43-93[Medline]
[Order article via Infotrieve]
-
Bowman, E. J.,
Siebers, A.,
and Altendorf, K.
(1988)
Proc. Natl. Acad. Sci. U. S. A.
85,
7972-7976[Abstract]
-
Hanada, H.,
Moriyama, Y.,
Maeda, M.,
and Futai, M.
(1990)
Biochem. Biophys. Res. Commun.
170,
873-878[Medline]
[Order article via Infotrieve]
-
Zeuzem, S.,
Feick, P.,
Zimmermann, P.,
Haase, W.,
and Kahn, R. A.
(1992)
Proc. Natl. Acad. Sci. U. S. A.
89,
6619-6623[Abstract]
-
Henomatsu, N.,
Yoshimori, T.,
Yamammoto, A.,
Moriyama, Y.,
and Tashiro, Y.
(1993)
Eur. J. Cell Biol.
62,
127-139[Medline]
[Order article via Infotrieve]
-
Palokangas, H.,
Metsikko, K.,
and Vaananen, K.
(1994)
J. Biol. Chem.
269,
17577-17585[Abstract/Free Full Text]
-
Heming, T. A.,
Bidani, A.,
Hinder, F.,
and Traber, D. L.
(1995)
J. Exp. Biol.
198,
1711-1715[Abstract/Free Full Text]
-
Li, J.,
and Eastman, A.
(1995)
J. Biol. Chem.
270,
3203-3211[Abstract/Free Full Text]
-
Perez-Sala, D.,
Collado-Escobar, D.,
and Mollinedo, F.
(1995)
J. Biol. Chem.
270,
6235-6242[Abstract/Free Full Text]
-
Gottlieb, R. A.,
Nordberg, J.,
Skowronski, E.,
and Babior, B. M.
(1996)
Proc. Natl. Acad. Sci. U. S. A.
93,
654-658[Abstract/Free Full Text]
-
Morana, S. J.,
Wolf, C. M.,
Li, J.,
Reynolds, J. E.,
Brown, M. K.,
and Eastman, A.
(1996)
J. Biol. Chem.
271,
18263-18271[Abstract/Free Full Text]
-
McConkey, D. J.,
and Orrenius, S.
(1996)
J. Leukocyte Biol.
59,
775-783[Abstract]
-
Trump, B. F.,
and Berezesky, I. K.
(1995)
Growth Factors and Tumor Promotion: Implications for Risk Assessment, pp. 121-131, Wiley-Liss, Inc., New York
-
McConkey, D. J.,
and Orrenius, S.
(1996)
Stem Cells
14,
619-631[Abstract]
-
Merritt, J. E.,
Armstrong, W. P.,
Benham, C. D.,
Hallam, T. J.,
Jacob, R.,
Jaxa-Chamiec, A.,
Leigh, B. K.,
McCarthy, S. A.,
Moores, K. E.,
and Rink, T. J.
(1990)
Biochem. J.
271,
515-522[Medline]
[Order article via Infotrieve]
-
Putney, J. W., Jr.
(1997)
Cell Calcium
21,
257-261[Medline]
[Order article via Infotrieve]
-
Hofer, A. M.,
Fasolato, C.,
and Pozzan, T.
(1998)
J. Cell Bio.
140,
325-334[Abstract/Free Full Text]
-
Jaconi, M. E. E.,
Lew, D. P.,
Monod, A.,
and Krause, K.-H.
(1993)
J. Biol. Chem.
268,
26075-26078[Abstract/Free Full Text]
-
Petersen, C. C. H.,
and Berridge, M. J.
(1995)
Biochem. J.
307,
663-668[Medline]
[Order article via Infotrieve]
-
Hall, A.
(1990)
Science
249,
635-640[Medline]
[Order article via Infotrieve]
-
Novick, P.,
and Zerial, M.
(1997)
Curr. Opin. Cell Bio.
9,
496-504[CrossRef][Medline]
[Order article via Infotrieve]
-
Olkkonen, V. M.,
and Stenmark, H.
(1997)
Int. Rev. Cytol.
176,
1-85[Medline]
[Order article via Infotrieve]
-
Bradbury, N. A.,
and Bridges, R. J.
(1994)
Am. J. Physiol.
267,
C1-C24[Abstract/Free Full Text]
-
Holman, G. D.,
Leggio, L. L.,
and Cushman, S. W.
(1994)
J. Biol. Chem.
269,
17516-17524[Abstract/Free Full Text]
-
Baldwin, S. A.,
Barros, L. F.,
and Griffiths, M.
(1995)
Biosci. Rep.
15,
419-426[Medline]
[Order article via Infotrieve]
Copyright © 1999 by The American Society for Biochemistry and Molecular Biology, Inc.