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J. Biol. Chem., Vol. 276, Issue 26, 23867-23872, June 29, 2001
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From the Physiology and Biophysics, University of Texas Medical
Branch, Galveston, Texas 77550-0641
Received for publication, February 16, 2001, and in revised form, April 24, 2001
Many animal cells release ATP into the
extracellular medium, and often this release is mechanosensitive.
However, the mechanisms underlying this release are not well
understood. Using the luciferin-luciferase bioluminescent assay we
demonstrate that a Xenopus oocyte releases ATP at a basal
rate ~0.01 fmol/s, and gentle mechanical stimulation can increase
this to 50 fmol/s. Brefeldin A, nocodazole, and
progesterone-induced- maturation block basal and mechanosensitive ATP
release. These treatments share the common feature of disrupting the
Golgi complex and vesicle trafficking to the cell surface and thereby
block protein secretion and membrane protein insertion. We propose that ATP release occurs when protein transport vesicles enriched in ATP fuse
with the plasma membrane. Collagenase, integrin-binding peptides, and
cytochalasin D also block ATP release, indicating that extracellular,
membrane and cytoskeletal elements are involved in the release process.
Elevation of intracellular Ca2+ does not evoke ATP release
but potentiates mechanosensitive ATP release. Our study
indicates a novel mechanism of mechanotransduction that would allow
cells to regulate membrane trafficking and protein transport/secretion
in response to mechanical loading.
Many, if not all, animal cells release ATP (or UTP) into the
extracellular medium, and often this release is
MS1 (1-6). External ATP acts
on ATP receptors that regulate diverse functions, including pain and
touch sensation, smooth muscle contractility, synaptic transmission,
platelet aggregation, epithelial fluid secretion, and endothelial
release of vasorelaxants (6-8). Furthermore, abnormalities in ATP
release may contribute to specific human diseases, most notably cystic
fibrosis (9). Although several mechanisms have been proposed to
contribute to ATP release, including synaptic vesicular release and
various membrane ion channels (10-16), the mechanism of MS ATP release
remains unknown.
Our interest in MS ATP release was stimulated by the discovery of
Nakamura and Strittmatter (1) that mechanical stimulation of the
Xenopus oocyte evokes ATP release without causing an
increase in membrane conductance. Here, we test the hypothesis that ATP release from the oocyte is mediated by the high rate (4,000-16,000/s) of membrane fusion of vesicles involved in transporting proteins from
the Golgi complex to the cell surface (17). This idea seemed plausible
given the identification of a specific ATP transporter that
concentrates ATP in the Golgi/ER lumen 50-100-fold above that in the
cytoplasm (18, 19). To test the hypothesis we have examined the effects
of BFA and other treatments that are known to disrupt membrane
trafficking and thereby block protein secretion (20-22).
A key issue for any MS process is the pathway by which mechanical
forces are transmitted to that process. For example, specific membrane
channel proteins in bacteria and animal cells respond directly to
tension developed in the lipid bilayer (23), while other MS processes
may be activated by forces transmitted via elements of the ECM and/or
CSK (i.e. tethers), possibly interconnected by integrins
(24). In the case of the MG cation channel endogenous to the
Xenopus oocyte, a bilayer model is favored, because the channel retains its mechanosensitivity in CSK-deficient plasma membrane
vesicles (25). However, because of the oocyte's large excess membrane
area (i.e. >500%) in the form of membrane folds and
microvilli, even large oocyte deformations (e.g. caused by inflation, aspiration, or fluid jet stimulation) fail to increase membrane conductance (26). In contrast, the mildest deformation of the
oocyte will activate ATP release (1, 27). This difference may arise
because MS ATP release involves a tethered mechanism, perhaps analogous
to the integrin-dependent, stretch-sensitive transmitter
release at the frog neuromuscular junction (28). To test this
possibility, we have examined the effects of integrin-binding peptides,
as well as specific agents that disrupt ECM and CSK elements.
The preparation of oocytes from Xenopus frogs was as
described previously (29). After surgical removal, the oocytes were incubated in Barth's solution, typically containing 1 mg/ml
collagenase Type II (Sigma) for 5 h on a rocker platform at room
temperature. In some cases, oocytes were not treated with collagenase
or were treated for longer duration (i.e. ~15 h).
Defolliculated oocytes were allowed to recover in Barth's solution for
~36 h at 17 °C. Experiments were performed on stage VI oocytes
usually 2-4 days after isolation.
ATP released from individual oocytes was monitored using the
luciferin-luciferase bioluminescence assay in a luminometer (2010 Monolight, PharMingen). The ATP monitoring concept (30) assumes that
during the measurement the luciferase activity remains constant and
degrades only a tiny fraction of the total ATP present (i.e. <1% min The measured ATP from an oocyte should reflect the balance between the
rates of ATP release and hydrolysis. However, the ecto-ATPase activity
of the oocyte is associated with the follicular cells and can be
inhibited by collagenase treatment (31). Under these conditions any
basal ATP release should result in ATP accumulation and a
time-dependent increase in bioluminescence. To measure
basal ATP release we gently aspirated an individual oocyte into the tip
(~2 mm diameter) of a Pasteur pipette and transferred it to the
luminometer cuvette. Measurements were then made every minute for at
least 15 min. To evoke MS ATP release the oocyte was stimulated in the
cuvette using a Gilson micropipettor with a P200 pipette tip to
repetitively "puff" 120 µl of the buffer at the oocyte. Special
care was taken not to directly touch or aspirate the oocyte into the
tip. For each mechanical stimulation three buffer puffs were applied
over approximately a 3-s duration. The rate of MS ATP release was
estimated by dividing the increment in ATP level (measured immediately
after stimulation) by the duration of the mechanical stimulation. To
estimate the cytoplasmic ATP concentration, the oocyte was ruptured by
repetitive aspiration into the tip of the puffer pipette.
The chemicals ATP, BAPTA.AM, A23187, BFA, cytochalasin D, progesterone,
and nocodazole were from Sigma; gadolinium (III) chloride hexahydrate
was from Aldrich; apyrase (200 units, 140 units/mg) was from Fluka; and
the peptides: GRGESP RGD (Gly-Arg-Gly-Glu-Ser-Pro, inactive), GRGDSP
RGD (Gly-Arg-Gly-Asp-Ser-Pro, anti-fibronectin-anti-vitronectin), GRGDdSP (Gly-Arg-Gly-Asp-D-Ser-Pro, selective
anti-fibronectin), and GPenGRGDSPCA RGD
(Gly-Pen-Gly-Arg-Gly-Asp-Ser-Pro-Cys-Ala(cyclical), where Pen indicates
penicillamine), selective anti-vitronectin) were from Life
Technologies, Inc. The concentration and incubation time for the
different drugs used were as follows: 5 µg/ml BFA for 2.5 h, 0.2 mM integrin-binding peptides for 1.5 h, 0.25 µM cytochalasin D for 2 h, 10 µM
BAPTA.AM or 10 µM GdCl3 for 45 min. To
facilitate the effect of nocodazole (see Ref. 32), oocytes were
previously exposed to cold treatment (i.e. 1 h in ND-96
on ice in the cold room) and then incubated in 20 µM
nocodazole for 1 h at 17 °C. Control experiments indicated that
cold treatment alone did not block either basal or MS ATP release.
A23187 was added to the cuvette to give a final concentration of 10 µM. Apyrase was also directly added to the cuvette (4 units in 2 µl, from a stock 140 units/mg). Stage VI oocytes were
matured in vitro by incubation in ND-96 plus 10 µM progesterone for ~12 h. Maturation was judged by the
appearance of a white spot on the animal pole (33). In zero-calcium
experiments, ND-96 minus CaCl2 was prepared in plastic
beakers and used in the buffer/assay mixture. Oocytes were gently
washed several times in the same medium before testing.
Fig. 1A shows ATP
calibration curves measured in control (ND-96), 100 µM
Gd3+, and hypotonic (ND-33) solutions. 100 µM
Gd3+ reduced assay sensitivity, whereas ND-33 increased it.
Fig. 1B illustrates an experiment simulating ATP release. In
the standard buffer/assay solution the background luminescence was
~100 relative luminescence units (i.e. same as the blank)
and equivalent to 2 fmol of added ATP. Repetitive applications of
15-fmol aliquots of ATP produced a staircase increase in luminescence,
and addition of apyrase abolished the response.
Fig. 2A indicates that the
initial transfer of the oocyte to the cuvette (i.e. at time
0) increased the base-line equivalent to adding ~20 fmol of ATP,
which was followed by a steady increase of ~10 amol/s
(8.6 ± 0.05 × 10
Brefeldin A Block of Integrin-dependent
Mechanosensitive ATP Release from Xenopus Oocytes Reveals a
Novel Mechanism of Mechanotransduction*
ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
1). Because the ATP is consumed by
the luciferase enzyme on a millisecond time scale, any change in ATP is
registered immediately. In our experiments, the luminometer cuvette
typically contained 140 µl of ND-96 buffer (in mM: 96 NaCl, 2 KCl, 1 MgCl2, 1.8 CaCl2, 5 NaHepes, pH
7.8) and 10 µl of the Sigma ATP assay reagent (1 mg/ml luciferin-luciferase mix). The assay was calibrated by plotting the log
of luminescence intensity (relative luminescence units) against the log
of ATP concentration (moles/liter) using the Sigma ATP calibration
standards. For both calibration and oocyte experiments, we measured the
ATP-induced light over a 15 s sampling periods. The background
light was measured and subtracted by running a blank. The ATP-induced
light was converted to moles of ATP according to the calibration plot
prepared each day. All treatments directly tested on ATP release were
first screened for effects on assay sensitivity.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (15K):
[in a new window]
Fig. 1.
Bioluminescence induced by applied ATP.
A, calibration plots of the bioluminescence assay in control
solution (ND-96), ND-96 plus 100 µM Gd3+, and
hypotonic solution ND-33. Whereas 100 µM Gd3+
added to the ND-96 reduced assay sensitivity, the hypotonic ND-33
increased it. Values represent the mean ± S.E. of <6 individual
experiments. B, with no oocyte present, the luminescent
response to repetitive applications of ATP (i.e. 15 fmol of
ATP in 1 µl) was measured. At the end of the experiment apyrase (4 units in 2 µl) was added. Bckg, background.
18 mol/s). In
comparison, addition of non-collagenase-treated oocytes did not
increase the luminescence above background, while addition of oocytes
treated overnight (i.e. 15 versus 5 h) with
collagenase showed a relatively larger increase in basal ATP release
(Table I). These results confirm a
previous study (31) that indicated follicle-enclosed oocytes
express significant ecto-ATPase activity, and this activity is
inhibited by collagenase. We believe the initial increase in ATP level
at zero time may be due in part to inadvertent mechanical stimulation
of the oocyte during the transfer because it is reduced along with MS
ATP release (see Fig. 3).
View larger version (20K):
[in a new window]
Fig. 2.
Basal and mechanosensitive ATP release from
Xenopus oocytes. A, the basal ATP
release from oocytes either pretreated with collagenase (1 mg/ml for
5 h) or not treated with collagenase. In each case the data
represent the mean ± S.E. (11 oocytes, 2 frogs). B,
ATP released from individual oocytes pretreated with collagenase (1 mg/ml for 5 h). The oocytes were mechanically (Mec)
stimulated by solution puffs (see "Experimental Procedures"), and
the first reading immediately after the stimulation is indicated by the
arrows. The oocyte was finally ruptured by aspiration into
the pipette tip (mean ± S.E., 8 oocytes from 2 frogs). After
oocyte rupture apyrase (4 units in 2 µl) was added.
The effects of various conditions on basal and MS ATP release from
Xenopus oocytes (see "Experimental Procedures" for details)
18 mol/s and as percentage of
control. Number of oocytes tested for each condition and the
reversibility of the block are also shown.
View larger version (18K):
[in a new window]
Fig. 3.
The effect of various agents on ATP-induced
bioluminescence before and after mechanical (Mec)
stimulation. A, in control oocytes and after
preincubation in B, brefeldin A (5 µg/ml/2.5 h);
C, progesterone (10 µM/~12 h); D,
anti-fibronectin peptide (GRGDdSP) (0.2 mM/1.5 h);
E, nocodazole (20 µM/1 h, see "Experimental
Procedures"); and F, cytochalasin D (0.25 µM/2 h). Values represent the mean ± S.E. of 12 (A), 25 (B), 12 (C), 8 (D),
19 (E), and 18 (F) individual experiments.
Fig. 2B shows MS ATP release from individual oocytes pretreated with collagenase. The first mechanical stimulation increased the ATP level from 25.14 ± 3.1 to 250 ± 30 fmol of ATP. Because the stimulus was applied over approximately a 3-s duration (see "Experimental Procedures"), the rate of MS ATP release was estimated to be ~70 fmol/s. Subsequent stimuli produced smaller increments in ATP release (i.e. ~50 fmol), possibly reflecting a reduction in the MS releasable pool and/or assay sensitivity (c.f. Fig. 1B). Note this apparent saturating response to gentle mechanical stimulation was ~4 orders of magnitude less than the ATP released by mechanical damage of the oocyte (see below). Mechanical rupture of the oocyte increased the luminescence equivalent to 300,000 fmol of ATP, which decayed rapidly in the first minute, presumably due to ATP hydrolysis by released/activated ATPase. Addition of apyrase further reduced the ATP-generated luminescence.
Several mechanisms may contribute to ATP release, and different
mechanisms may underlie basal and MS release. For example, the oocyte
expresses hemi-gap and Gd3+-sensitive MG channels, both of
which have been implicated in ATP release from epithelial cells (12,
15). However, mechanical stimuli much stronger than that required to
cause ATP release, fails to activate a membrane conductance increase
(1, 26, 27). Furthermore, 10 µM Gd3+, which
completely blocks hemi-gap and MG channel activity (33, 34), does not
affect ATP release (Table I). 100 µM Gd3+
caused an apparent, partial inhibition of ATP release (data not presented), but this may be due to the higher Gd3+ reducing
assay sensitivity (see Fig. 1A). It has recently been proposed that strong hyperpolarization of the oocyte (i.e.
to 
200 mV) may activate an ATP-selective conductance (16). However, our studies indicate similar hyperpolarizations activate a nonselective conductance that fails to saturate with prolonged hyperpolarization and
recovers only slowly (i.e. >10 min) following
repolarization (34). These properties are consistent with reversible
membrane dielectric breakdown. Whatever the underlying mechanism, this voltage-activated conductance does not mediate the non-electrogenic MS
ATP release measured at resting potentials (1).
Non-electrogenic ATP release could be mediated by exocytosis, a
specific ATP transporter or both. To test the hypothesis that ATP
release is associated with fusion of protein carrier vesicles with the
plasma membrane, we examined the effects of different agents that
disrupt the Golgi complex and thereby block membrane/protein trafficking. BFA reversibly blocks anterograde vesicular transport through a redistribution of the Golgi complex into the ER (22) and
thereby inhibits protein secretion from animal cells, including the
Xenopus oocyte (35, 36). We found that BFA treatment also blocks basal and MS release (Table I, Fig. 3B), and the
block is reversed 3 h after removal of BFA (data not presented).
Progesterone maturation of the oocyte is associated with the
disappearance of the Golgi apparatus and block of protein secretion
(37). We found that basal and MS ATP release is also absent in matured oocytes (Fig. 3C, Table I). The microtubule network is
critical in maintaining the integrity and function of the Golgi complex (38), and its depolymerization by nocodazole disrupts the Golgi complex
and vesicle trafficking (39). We found that nocodazole also blocks
basal and MS ATP release (Table I, Fig. 3E), and this block
was reversed within 3 h after nocodazole exposure (data not
presented). The above treatments did not alter the normal oocyte
resting membrane potential (~30 mV) nor the amount of ATP released
upon oocyte rupture (i.e. ~3 nM) (data not presented).
To determine the pathway by which mechanical forces are transmitted to the ATP release process, we tested specific agents that interfere with ECM, membrane, and CSK elements. Evidence for ECM involvement was indicated by prolonged exposure of oocytes to collagenase (i.e. 15 versus 5 h), which blocked MS ATP release by 99% (Table I). In contrast, basal ATP release appeared enhanced (i.e. 129%, see Table I), probably because prolonged collagenase exposure produces a more complete inhibition of ecto-ATPase activity. To determine whether ECM-integrin interactions are important in ATP release, we tested specific RGD-containing peptides that disrupt integrin binding to ECM proteins (24, 28). We found that the peptide (GRGDdSP), which inhibits integrin binding to fibronectin, but not to vitronectin, significantly reduced basal and MS ATP release (Fig. 3D, Table I). The nonspecific anti-fibronectin-anti-vitronectin peptide (GRGDSP) produced a similar effect (data not presented). However, neither the specific anti-vitronectin peptide (GPenGRGDSPCA) (Table I) nor an inactive control peptide (GRGESP) inhibited ATP release (data not presented). Because integrins mechanically link the ECM to the actin CSK (24), we tested the effect of disruption of actin microfilaments. We found that treatment of oocytes with cytochalasin D blocked both basal and MS release (Fig. 3F, Table I). The selective block of MS ATP release by collagenase indicates that basal ATP release can proceed even when ECM elements critical for mechanosensitivity are disrupted. However, intact integrin and microfilament interactions are apparently required for basal ATP release. These interactions may be involved in transmitting background mechanical forces (i.e. arising from gravity and/or CSK tension) that modulate the release mechanism.
We found that external Ca2+ was not required for basal or
MS ATP release (Figs. 4, A and
B). Furthermore, 10 µM A23187 did not
stimulate ATP release. However, A23187 treatment consistently potentiated (~10-fold) MS ATP release (Fig. 4C, Table I).
The A23187 potentiation was blocked when measured in
zero-Ca2+ external solution (data not shown) and by
preincubation of the oocyte in 10 µM BAPTA-AM (Fig.
4D). We also tested A23187 on progesterone matured oocytes,
which causes massive cortical granule-membrane fusion, in addition to
activating the Ca2+-sensitive Cl conductance
(e.g. see Fig. 8 in Ref. 26). However, as with immature
oocytes A23187 did not increase ATP release (data not presented),
indicating that cortical granules do not contain ATP and
Ca2+-activated Cl channels do not mediate ATP
efflux (see also Fig. 4C).
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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The Xenopus oocyte belongs to a growing list of animal
cells that exhibit MS ATP (or UTP) release (1-6). Here, we provide new
details on a novel mechanism that may explain why basal and MS ATP
release is a general feature of animal cells. In summary, MS ATP
release can occur without membrane damage based on its reversible block
by specific agents (e.g. BFA, nocodazole, and integrin-binding peptides) and irreversible block by
progesterone-induced maturation. It is also not mediated by the
endogenous MG, hemi-gap, or Ca2+-activated Cl
channels. Instead, we propose that (i) ATP release is dependent upon
BFA-sensitive trafficking of ATP-enriched vesicles that transport protein from the Golgi to the cell surface, (ii) the vesicle
trafficking and/or membrane fusion is facilitated by mechanical forces
transmitted from the ECM to the CSK possibly via membrane integrins,
(iii) increase in [Ca2+]i acts cooperatively with
mechanical forces to promote ATP release, and (iv) progesterone
maturation of the oocyte blocks ATP release, implying that ATP release
may play some role in the normal maintenance of the follicle and/or
immature oocyte.
The Xenopus oocyte releases ATP at a basal rate of around 0.01 fmol/s, and gentle mechanical stimuli can transiently increase this 5,000-fold to 50 fmol/s. In comparison, mechanical rupture of the oocyte results in the release of 300,000 fmol of ATP. This indicates a cytoplasmic ATP concentration of around 1 mM (i.e. for an oocyte volume ~0.5 µl). However, this is likely an underestimate of the actual concentration given the rapid degradation of released ATP by activated ATPase activity (Fig. 2B). Furthermore, this estimate does not include ATP compartmentalized within mitochondria and the ER/Golgi cisternae. Clearly, the basal and MS-released ATP represents a tiny fraction of the total oocyte ATP.
According to our primary hypothesis, both basal and MS ATP
release arise from fusion/exocytosis of vesicles involved in
transporting proteins from the Golgi complex to the cell surface. This
hypothesis is supported by the block of ATP release by BFA, nocodazole,
and progesterone-induced maturation of the oocyte. All three treatments cause disappearance of the Golgi complex and thereby block protein transport/secretion from the Xenopus oocyte (20-22, 36,
38). According to Zampighi et al. (17), the vesicles that
transport proteins to the cell surface fuse with the cell membrane at a rate of at least 4,000 s1. To account for
basal ATP release, each vesicle would need to release 2.5 × 1021 mol of ATP (i.e. 1,500 ATP
molecules), consistent with a vesicular ATP concentration of ~5
mM at the time of exocytosis (i.e. for a vesicle
diameter/volume of 0.1 µm/5.2 × 1019
liters, see Ref. 17). This concentration is only ~5 times
higher than the estimated cytoplasmic concentration and is reasonable given the identification of a specific ATP transporter that can concentrate ATP in the Golgi/ER lumen 50-100 times above that in
external medium (18, 19). Presumably, the ATP released reflects the
residual ATP left in the vesicle after ATP-dependent processing of cargo proteins is complete (18). Although we currently favor ATP release by exocytosis, our luminescence measurements cannot
exclude the possibility that block of membrane trafficking does not
also inhibit membrane insertion of electroneutral ATP transporters.
However, even if some resident Golgi ATP transporters reach the cell
membrane (i.e. despite Golgi retention mechanisms, see Ref.
40), in order for them to mediate ATP efflux they would require
obligatory ADP/AMP influx (18, 41). In this case, conditions that
reduced local external ADP/AMP (e.g. block of ecto-ATPase
activity or buffer puffs directed at the surface) would be expected to
reduce rather than increase ATP efflux (c.f. Table I).
Furthermore, because BFA block of ATP release is >95% complete within
2.5 h, it would imply the hypothesized membrane-inserted ATP
transporters have a membrane half-life of less than 1 h. In comparison, the measured half-life of membrane proteins expressed in
oocytes ranges between 4 and 36 h (17, 42, 43).
It is possible that the ATP released from the oocyte plays no additional functional role but merely represents the unavoidable consequence/cost of requiring ATP in the Golgi/ER lumen as a substrate and energy source for processing secretory and membrane-bound proteins (18, 44, 45). The absence of endogenous ATP receptors on the oocyte (e.g. see Ref. 1) would seem to rule out any autocrine signaling role. However, it is possible that released ATP activates endogenous ATP receptors on follicular cells (e.g. see Ref. 46) and thereby participates in a regulatory cell-cell loop involving the electrically coupled oocyte and follicular cells (47). In this case, changes in external "mechanical tone" exerted on the oocyte by the surrounding follicular cell layer may regulate MS ATP release. It has been shown that direct oocyte inflation or oocyte aspiration can induce ATP release as monitored by purinergic receptors heterologously expressed in the oocyte (1). However, our preliminary results indicate that osmotic swelling of the oocyte (i.e. in ND-33) does not significantly increase ATP release.2
The demonstration that the BFA-sensitive process that mediates constitutive protein secretion and membrane protein insertion is mechanosensitive has implications beyond the oocyte. For example, this process may serve as a general intrinsic mechanism for regulating animal cell growth and proliferation in response to mechanical loading. At least consistent with this idea is the demonstration that stretch-induced hypertrophy of isolated cardiac myocytes is mediated by increased release of angiotensin II (48) and that mechanical strain-induced proliferation of fetal lung cells is associated with increased constitutive release of glycosaminoglycans and proteoglycans (49). It is also interesting that MS ATP release from the oocyte shares some similarities with stretch-induced enhancement of transmitter release at the frog NMJ (28). For example, both MS processes are blocked by integrin-binding peptides, and both can occur when elevation of [Ca2+]i is prevented (Ref. 28 and Fig. 4D). However, whereas elevation of [Ca2+]i alone can evoke synaptic transmitter release, it only serves to facilitate MS ATP release. This facilitation may be related to the demonstrated Ca2+ dependence of vesicle fusion during intra-Golgi transport (50). Finally, our results provide possible clues to the mechanism underlying CFTR-dependent ATP release from specific cell types (see Ref. 51). For example, cAMP activation of CFTR has been shown to be associated with a sustained increase in ATP release (33 fmol/s) from CFTR-transfected oocytes (13) but not from CFTR-expressing Calu-3 cells (4). Interestingly, in CFTR-transfected Xenopus oocytes (as well as in human epithelial cells) it has been demonstrated that cAMP activation of CFTR occurs mainly by increased membrane insertion of new CFTR channels (52-54), whereas in other cell types, including Calu-3 cells, cAMP directly activates already inserted CFTR channels (55, 56). Furthermore, BFA and nocodazole block CFTR activation in the former cell type (53) but not the latter (56). A plausible explanation for these cell type-specific effects is that CFTR confers cAMP sensitivity on membrane trafficking and thereby ATP release.
In conclusion, our study reveals a novel mechanism of
mechanotransduction that involves ATP release via BFA-sensitive vesicle trafficking between the Golgi complex and the cell surface. Since this
mechanism is common to most animal cells, it would explain why
constitutive ATP release is widespread in animal cells (57). The idea
that the process that determines constitutive protein secretion and
membrane protein insertion is MS has implications for MS regulation of
cell growth and development. This process may also account for why
overexpression or activation of specific membrane proteins
(e.g. CFTR) in some cells has been associated with increased
ATP release.
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ACKNOWLEDGEMENT |
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We thank Dr. Javier Navarro for the use of his luminometer and Jay Steer for computer assistance.
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FOOTNOTES |
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* This work was supported by the National Institutes of Health and by Muscular Dystrophy Association.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
To whom correspondence should be addressed: Physiology and
Biophysics, University of Texas Medical Branch, 301 University Blvd.,
Galveston, TX 77550-0641. Tel.: 409-772-5464; Fax: 409-772-3381; E-mail: ohamill@utmb.edu.
Published, JBC Papers in Press, April 24, 2001, DOI 10.1074/jbc.M101500200
2 R. Maroto and O. P. Hamill, unpublished observations.
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ABBREVIATIONS |
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The abbreviations used are: MS, mechanosensitive; BFA, brefeldin A; MG, mechanically gated; ECM, extracellular matrix; ER, endoplasmic reticulum; CSK, cytoskeleton; BAPTA, 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid; AM, acetoxymethyl ester.
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REFERENCES |
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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