1 Laboratoire de Neurogenèse et Morphogenèse au cours du
Développement et chez l'Adulte (NMDA), UMR 6156, Institut de Biologie
du Développement de Marseille, Faculté des Sciences de Luminy,
case 907, Université de la Méditerranée, 13288, Marseille
Cedex 09, France
2 Columbia University, New-York, NY 10032, USA
* Author for correspondence (e-mail: lebivic{at}ibdm.univ-mrs.fr)
Accepted 28 August 2002
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Summary |
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Key words: Caveolin, Epithelia, Intestine, Golgi
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Introduction |
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In this work we first showed that Caco-2 cells do not express Cav-2 at detectable levels and used them to study Cav-2 subcellular localization in the presence or the absence of Cav-1. Cav-2 was restricted at steady state to the Golgi complex of Caco-2 cells and this localization is not affected by expression of Cav-1 as opposed to what was shown in two other cells types. Furthermore, using chimeras between Cav-1 and -2, we could identify the molecular determinant responsible for this Golgi restriction as the scaffolding domain of Cav-2. This Golgi retention might be responsible for the low number of caveolae formed at the basolateral surface of Caco-2 cells expressing both caveolins. Another conclusion of this work is that expression of caveolins at the cell surface is necessary but not sufficient to promote caveolae formation and that the level of incorporation of the chimeras into lipid rafts might be regulating the building of caveolae.
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Materials and Methods |
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Antibodies
Affinity-purified anti-Cav-1 rabbit polyclonal antibody, N20, directed
against Cav-1 N-terminal residues 2-21, and the monoclonal antibody against a
Myc epitope (9 E10) were from Santa Cruz Biotechnology (Santa Cruz, CA).
Rabbit polyclonal antibody anti-Cav-2 was raised against a synthetic peptide
using the residues DFGDLEQLADSGDR of canine Cav-2
(Scheiffele et al., 1998).
Mouse monoclonal antibody anti-Cav-2 (C57820, IgG1) was purchased from
Transduction Laboratories (Lexington, KY). Mouse monoclonal antibody against
antigen 525 (Ag525 mAb), a marker of the basolateral membrane, has been
described previously (Le Bivic et al.,
1988
). Mouse mAb against sucrase-isomaltase (SI)
(Beaulieu et al., 1989
), a
marker of apical membranes, was kindly provided by A. Quaroni (Ithaca, NY).
Polyclonal anti-PLAP was from Accurate Chemical and Scientific Corp.
(Westbury, NY). Mouse anti-APN has been already described in
(Le Bivic et al., 1990
) and
mouse mAb against Giantin was a kind gift from H. P. Hauri (Basel,
Switzerland).
Western blot analysis and immunoprecipitations, flotation and
velocity gradients
Western blots were performed as already described in
(Mirre et al., 1996) while
immunoprecipitations were done as in (Le
Bivic et al., 1989
). Flotation gradients were prepared as
described in (Mirre et al.,
1996
) and velocity gradients were performed as in
(Scheiffele et al., 1998
),
except that no SDS was added to cell lysates and gradients and that Triton
X-100 final concentration was 1%. Aliquots were analyzed by SDS-PAGE and
western blotting and quantified with BioImage Quantifier software (Bio-image,
Ann Arbor, MI).
Indirect immunofluorescence and laser scanning confocal microscopy
(LSCM)
Caco-2 cells were stained for immunofluorescence as described before
(Gilbert et al., 1991) using
the following antibodies Cav-1, Cav-2, SI, Ag525 and 9E10 at 1:100 dilution,
except for Giantin used at 1:500 dilution. Secondary antibodies, i.e.,
fluorescein isothiocyanate (FITC) anti-mouse IgG or tetramethyl rhodamine
(TRITC) conjugated anti-rabbit antibodies were used at a 1:200 dilution
(Jackson). Samples were examined with a Zeiss LSM confocal system (Carl Zeiss,
Germany) and a Zeiss microscope Axiovert microscope 135M. Confocal images were
collected using argon and He-Ne lasers with attenuating filters as excitation
sources at 488 nm or 543 nm, for FITC or TRITC, respectively. For simultaneous
excitation of FITC and TRITC, a double-banded beam splitter DBSP 488/543 was
used. Excitation filters FT 510 nm or LP 560 nm, and emission filters BP
515/565 nm or LP 570 nm were used for separate acquisition of FITC and TRITC
signals.
Electron microscopy
Filter-grown confluent Caco-2 cells were processed for electron microscopy
according to a method used by (Lipardi et
al., 1998) for Fischer rat thyroid (FRT) cells. Cells were rinsed
three times with PBS and then fixed for 30-60 minutes at room temperature with
2.5% glutaraldehyde plus 0.1% tannic acid in 0.1 M sodium cacodylate buffer pH
7.3. Cells were then rinsed three times in 0.1 M cacodylate buffer and
post-fixed for 30-60 minutes with 1% osmium tetroxide in the same buffer. In
some cases, a fixative formula including potassium ferricyanide
(K3Fe (CN)6) to the osmium step was used to enhance membrane
contrast and preservation in cultured cells. Following post-fixation, filters
were abundantly rinsed with the buffer, cut from the holder, and stained en
bloc with 3% uranyl acetate in 50% ethanol (or acetone) for 20-30 minutes.
Samples were then dehydrated in a graded series of ethanols (or acetones) and
finally embedded in epon 812 (Polysciences, Warington, PA).
Immunogold electron microscopy
Filter-grown confluent Caco-2 cells were thoroughly rinsed with PBS and
fixed for 1 hour in 8% paraformaldehyde in PBS. After washing the cells were
scrapped off the dishes, collected, infiltrated with 6% gelatin in PBS at
37°C, put on ice and infiltrated with 2.3 M of sucrose in PBS. Samples
were then frozen in liquid nitrogen. Ultrathin cryosections were incubated
overnight at 4°C with anti-Cav-1 antibody (N20) diluted 1:30 or anti-Cav-2
antibody (mAb 65) diluted 1:20 in 5 to 10% goat serum in PBS. Primary
antibodies were revealed using colloidal gold 15 nm or 6 nm-conjugated goat
anti-rabbit or anti-mouse IgG respectively in the same buffer. Sections were
then fixed rapidly with 2% glutaraldehyde, rinsed in bidistilled water and
treated with 0.3% uranyl acetate and 1.8% methyl cellulose in bidistilled
water on ice.
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Results |
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Subcellular localization of Cav-1 and Cav-2
We next sought to determine the subcellular localization of Cav-1 and Cav-2
in transfected Caco-2 cells. Clones expressing either Cav-1, Cav-2 or both
were double labeled with monoclonal antibodies against endogenous markers of
Caco-2 cells and anti-Cav-1 polyclonal antibody
(Fig. 2a,c,e) or anti-Cav-2
polyclonal antibody (Fig.
2b,d,f). In agreement with a previous study
(Vogel et al., 1998),
exogenous Cav-1 was present on the basolateral membrane where it colocalized
with a basolateral antigen (Ag525) (Fig.
2c) and in intracellular compartments comprising the Golgi complex
as identified by the marker Giantin (Fig.
2e). No significant colocalization was observed with an apical
marker, SI (Fig. 2a) and only a
minor population colocalized with transferrin receptor or LAMP1 (not shown).
Cav-2 was mostly detected in the Golgi complex where it colocalized with
Giantin antibodies (Fig. 2f).
No significant overlap was observed with transferrin receptor or LAMP1 by
confocal microscopy (not shown). Strikingly, no staining of either apical or
basolateral membranes could be detected indicating that Cav-2 did not
accumulate at the plasma membrane in transfected Caco-2 cells. This was
confirmed by immunoelectron microscopy on frozen sections of Cav-1 and Cav-2
clones (Fig. 3). While gold
particles were found on the basolateral membrane with anti-Cav-1 antibodies in
Cav-1 cells (Fig. 3B), none
were detected in Cav-2 cells using anti-Cav-2 polyclonal antibodies
(Fig. 3A). In Caco-2 cells, the
Golgi complex was dispersed and very close to the basolateral membrane rather
than being concentrated close to the nucleus as in most cells. This was
confirmed by electron microscopy in which dictyosomes were seen extending
tubules and vesicles within less than 0.5 µm from the lateral membrane (not
shown).
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Caveolae formation in Cav-1 and/or Cav-2 expressing Caco-2 cells
It has been previously described that exogenous expression of canine Cav-1
in Caco-2 cells promoted the biogenesis of caveolae
(Vogel et al., 1998). Since
production of caveolae did not correlate with the amounts of exogenously
expressed Cav-1, these authors postulated that another factor necessary for
caveolae formation was missing in Caco-2 cells. Our finding that Caco-2 cells
did not express Cav-2 led us to hypothesize that it could be that factor. We
thus quantified caveolae formation in Caco-2 cells expressing either Cav-1
and/or Cav-2 (Table 1). In
Cav-1 clones, the average number of caveolae observed by mm of filter was 46,
in good agreement with what was described before
(Vogel et al., 1998
). Cav-2
clones did not show any increase in caveolae production over untransfected
cells (less than 4/mm) confirming that Cav-2 by itself was unable to promote
caveolae assembly. In cells expressing both caveolins, there was no increase
in caveolae numbers over cells expressing only Cav-1 (average 58/mm). Thus
expression of Cav-2, in Cav-1-expressing cells, was not able to stimulate
caveolae production to the levels observed in MDCK cells
(Vogel et al., 1998
;
Mora et al., 1999
). In all
clones, caveolae were restricted to the basolateral domain as in MDCK and FRT
cells (Vogel et al., 1998
;
Mora et al., 1999
). If Cav-2
acted as a co-factor in caveolae formation, its co-expression with Cav-1
should have increased the number of caveolae formed. Since it was not the
case, we looked for the localization of this protein in cells expressing also
Cav-1. Co-expression did not alter the typical staining of the Golgi complex
and in particular no staining of the plasma membrane was observed
(Fig. 4a,b). This data was
confirmed by transient expression of Cav-2-GFP fusion proteins with GFP
attached either on the N- or C- terminus of Cav-2. Both constructs gave the
same intracellular staining in Cav-1 cells with no labeling at the cell
surface (Fig. 4c,d). These
results were further confirmed by electron microscopy. No labeling of Cav-2
could be detected on the plasma membrane while Cav-1 was found at the plasma
membrane and in caveolae-like structures
(Fig. 3C,F,G). Internal
membranes and vesicles were positive for both caveolins
(Fig. 3D,E) indicating that
they were indeed in close proximity in the Golgi complex.
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Cav-1 interacts with Cav-2 without inducing a change in its
localization
The lack of Cav-2 redistribution in Cav-1 expressing cells led us to
investigate whether the two proteins could interact in Caco-2 cells when
co-expressed. In MDCK cells Cav-1 and Cav-2 form oligomers of very high
molecular weight as assessed by sucrose velocity gradients
(Scheiffele et al., 1998).
When expressed alone, Cav-1 was mainly found in the bottom part of such
gradients, indicating that it was able to form high molecular weight complexes
in Caco-2 cells (Fig. 5A) as in
other cells. Cav-2, on the other hand, migrated only in the top fractions of
the gradients, suggesting that it was not associated into a complex. When
co-expressed, however, a sizeable proportion of Cav-2 (>50%) migrated in
the bottom fractions of the gradients together with Cav-1. Thus Cav-1
expression modified the behavior of Cav-2, leading to its inclusion into high
molecular weight complexes. In Caco-2 cells expressing both Cav-1 and Cav-2,
no Cav-1 was detected in the top-half of the gradient, in contrast with cells
only expressing Cav-1, which suggests a possible regulatory effect of Cav-2 on
Cav-1 complexes (Fig. 5A). To
investigate whether Cav-1 and Cav-2 interact in the same complexes, Caco-2
cells expressing Cav-2 or both caveolins were immunoprecipitated with
antibodies against Cav-1 or Cav-2 (Fig.
5B). Cav-2 was able to pull-down Cav-1 from Caco-2 cells
expressing both proteins, confirming that the two proteins indeed interact
directly or indirectly. Strikingly, Cav-2 was not detected in Cav-1
immunoprecipitates, suggesting that the stoechiometry of the complexes must be
in favor of several molecules of Cav-1 for one molecule of Cav-2 as was also
reported in MDCK cells (Scheiffele et al.,
1998
).
|
The scaffolding domain of Cav-2 is involved in its Golgi
localization
I order to understand why Cav-2 is restricted to the Golgi complex while
Cav-1 is not, in Caco-2 cells, we produced chimeras between Cav-1 and Cav-2 to
identify a region of Cav-2 that could be responsible for this localization. We
designed four chimeras (Fig. 6)
which were transfected into Caco-2 cells. Several clones were selected for
each chimera and we controlled that the anti-Cav-1 antibody did recognize the
chimera produced since it is directed against the first 20 amino acids of
Cav-1 (Fig. 7A). All chimeras
migrated at a slightly higher molecular position in the SDS-PAGE due to the
addition of the myc-epitope. Their subcellular localization was determined
using double labeling with endogenous markers and confocal microscopy.
Chimeras made of the N-terminal cytoplasmic and transmembrane domains (1 to
152) of Cav-1 (CH-I) or only the N-terminal cytoplasmic domain (1 to 119) of
Cav-1 (CH-II) were transported to the basolateral surface where they
co-localized with the Ag525 while little was found in the Golgi complex
(Fig. 7B). Strikingly, when
only the first 98 residues of the N-terminal cytoplasmic domain of Cav-1 were
grafted to the scaffolding, transmembrane and C-terminal domains of Cav-2
(CH-III), a strong co-localization with Giantin was observed
(Fig. 7B). This data indicated
that the scaffolding domain (SD) of Cav-2 was responsible for its accumulation
in the Golgi apparatus. To test this hypothesis, we designed a chimera in
which we replaced the SD of Cav-1 by the one from Cav-2
(Fig. 6). After expression in
Caco-2 cells, this chimera containing the SD of Cav-2 (CH-IV) was concentrated
in the same compartment as Giantin, demonstrating that the SD of Cav-2 was
indeed responsible for its Golgi accumulation
(Fig. 7B).
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Lipid-raft association but not oligomerization of Cav-1 correlates
with caveolae formation in Caco-2 cells
In order to understand the factors controlling caveolae formation in Caco-2
cells, we counted the number of caveolae formed in Caco-2 cells expressing the
different chimeras (Table 1).
To our surprise, only CH-I was able to promote caveolae formation at levels
similar to what was observed with Cav-1. All other chimeras showed numbers of
caveolae comparable to untransfected Caco-2 cells (less than 4/mm). Since both
CH-I and II were expressed at the cell surface, this condition was necessary
but not sufficient to form caveolae. It has been proposed that caveolae
formation might be regulated by the ability of Cav-1 to form high molecular
weight oligomers. We thus tested the state of oligomerization of the chimeras
(Fig. 8B). As opposed to Cav-1
(Fig. 8A), none of the chimeras
was able to reach a significant percentage of oligomers and indeed the
chimeras behave more like Cav-2 in this respect. Thus, oligomerization and
formation of caveolae appeared to be uncoupled events. To try to understand
why only CH-I was able to promote caveolae formation, we tested whether the
inclusion of caveolins or chimeras into Triton-resistant lipid domains could
correlate with the ability to trigger this event. For this the
Triton-resistant light membrane (or raft) fractions were obtained after
sucrose flotation gradients and analyzed by western blots with the anti-Cav-1
antibody. MDCK cells were used as a control and, in these cells, the majority
(60%) of Cav-1 was found in the raft fraction. In Caco-2 cells transfected
with Cav-1, this value was 30% and co-expression of Cav-2 raised it up to 50%
(Fig. 9A). Cav-2, when
expressed alone in Caco-2 cells, was found at low levels (10%) into rafts and
co-expression with Cav-1 increased its partition into rafts to 20%
(Fig. 9A). Surprisingly, of all
the chimeras, only CH-I was predominantly found into rafts (60%) while CH-II,
III and IV were found in this fraction at levels that were comparable to Cav-2
(Fig. 9B). These data raised
the possibility that there was a relationship between the enrichment of
caveolins into rafts and caveolae building.
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Discussion |
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Interactions between Cav-1 and Cav-2
When expressed in Caco-2 cells, Cav-1 and Cav-2 showed different
subcellular localization. Cav-1 was found in the Golgi complex, in
intracellular vesicles and in the basolateral membrane confirming and
extending a previous study performed with an antibody recognizing only plasma
membrane Cav-1 (Vogel et al.,
1998). In addition we showed that Cav-1 is below detection levels
at the apical membrane as opposed to what was observed in MDCK cells.
Exogenous Cav-2 was accumulated in the Golgi complex of Caco-2 cells as
identified by colocalization with Giantin. No Cav-2 could be detected at the
cell surface under our culture conditions suggesting that it might never reach
the plasma membrane. Accumulation of Cav-2 in the Golgi complex of FRT and
K562 cells has been reported (Mora et al.,
1999
; Parolini et al.,
1999
). In both cases, this localization was observed in the
absence of Cav-1 expression and Cav-1 transfection induced a partial
redistribution of Cav-2 to the cell surface demonstrating that Golgi retention
was in part a consequence of a lack of Cav-1. In Caco-2 cells, however, Cav-1
expression did not modify Cav-2 localization suggesting that another limiting
factor was missing to export Cav-2 to the plasma membrane. While Cav-1 could
not re-localize Cav-2, both proteins could associate with each other since
antibodies against Cav-2 immunoprecipitated Cav-1 in co-expressing cells. This
association correlated with the ability of Cav-1 to recruit Cav-2 into high
molecular weight complexes as it was shown in MDCK, FRT and K562 cells
(Scheiffele et al., 1998
;
Mora et al., 1999
;
Parolini et al., 1999
). On the
other hand, Cav-2 seemed to stabilize Cav-1 present in these larger complexes
since in its absence Cav-1 was also detected in complexes of lower molecular
weight. Formation of Cav-1/Cav-2 complexes might start early when both
caveolins exit the endoplasmic reticulum. The localization of Cav-2 in the
Golgi complex at steady state implies that Cav-1 might still interacts with
Cav-2 in this compartment. This interaction appeared to be rather stable since
complexes of Cav-1 and Cav-2 could be co-immunoprecipitated with Cav-2
antibodies even 15 hours after a metabolic pulse (not shown). Whether this
interaction needs to be broken to allow Cav-1 exit from the Golgi complex
remains to be investigated. Alternatively, a pool of Cav-1 might never
associate with Cav-2 and reach the cell surface as in Cav-1 expressing cells.
It is worth to note that a similar situation was recently found in mouse
macrophages. In these cells Cav-2 is present primarily in the Golgi complex
while Cav-1 is accumulated at the cell surface
(Gargalovic and Dory, 2001
)
suggesting that the two proteins can be uncoupled in cells and that the role
of Cav-2 might be more than a simple accessory protein for Cav-1.
Role of the scaffolding domain in the accumulation of Cav-2 in the
Golgi complex
To identify the determinants responsible for the accumulation of Cav-2 in
the Golgi complex of Caco-2 cells we have chosen to use chimeras between Cav-1
and 2, a strategy respecting the normal conformation of caveolins, which are
suspected to have a hairpin structure. Using this approach we demonstrated
that the switch of the SD of Cav-1 by the same region from Cav-2 was enough to
ensure the accumulation of Cav-1 in the Golgi complex. So far, several studies
have been performed to dissect the molecular requirements regulating the
subcellular localization of Cav-1 but these studies used a totally different
strategy. Instead of chimeras, truncated proteins were produced and
transfected to follow their intracellular behavior
(Schlegel and Lisanti, 2000;
Machleidt et al., 2000
;
Luetterforst et al., 1999
).
This approach led to the identification of several regions of Cav-1 that
control its subcellular localization or membrane association. In particular,
it was found that the C-terminal end of Cav-1
(Schlegel and Lisanti, 2000
)
or Cav-3 (Luetterforst et al.,
1999
) was enough to ensure Golgi association. Thus one possibility
is that by introducing the SD of Cav-2 into Cav-1, a plasma membrane targeting
signal was removed from Cav-1 leading to the preponderance of the Golgi
C-terminus signal. It was also demonstrated that part of the SD of Cav-1
appears to control its exit from the Golgi
(Machleidt et al., 2000
)
confirming this hypothesis. Thus, the SD of Cav-2 allows this protein to have
a different subcellular localization from Cav-1, strongly suggesting that the
two proteins have different fates and probably different partners and
functions. This is also supported by the fact that the SD of Cav-1 binds to a
consensus sequence in its partners while Cav-2 failed to interact with these
proteins under the same conditions (Couet
et al., 1997
).
Role of Cav-1 and Cav-2 in caveolae formation
Previously, we had shown that Caco-2 cells did not express Cav-1 and did
not show any morphologically recognizable caveolae
(Mirre et al., 1996). This was
confirmed later and in addition it was shown that the expression of canine
Cav-1 in these cells led to caveolae formation
(Vogel et al., 1998
).
Surprisingly, not only the number of caveolae produced was 10 times lower than
in MDCK cells but, it was also independent of the levels of Cav-1 expressed
whereas, in lymphocytes, a correlation between Cav-1 expression levels and the
number of newly generated caveolae was observed
(Fra et al., 1995
). We
hypothesized that the lack of Cav-2 expression could have been the reason why
caveolae formation was not optimal in Caco-2 cells. However, our data
demonstrated that Cav-2 was not able to trigger or stimulate caveolae
formation. Cav-2 expression did not increase significantly the number of
caveolae in Caco-2 cells that expressed levels of Cav-1 equivalent to MDCK
cells. Intriguingly, these results are different from those described in FRT
cells (Mora et al., 1999
)
which only express Cav-2. Expression of Cav-1 in these cells promoted caveolae
formation (Lipardi et al.,
1998
) to numbers roughly similar to those found for MDCK cells
(Mora et al., 1999
). A likely
explanation is that in FRT cells, the expression of Cav-1 induced a partial
relocation of Cav-2 to the cell surface that we did not observe in Caco-2
cells. As a consequence, in Caco-2 cells, Cav-2 cannot play its potential
regulatory role on the number of caveolae produced at the cell surface. This
hypothesis is at odds with the fact that in Cav-2 deficient mouse there is an
intact caveolar membrane system, suggesting that Cav-2 has no regulatory role
on the number of caveolae (Razani et al.,
2002
).
The formation of caveolae is correlated to concentration of caveolin
into rafts
The mechanisms regulating the formation of caveolae are still poorly
understood but a major factor could be the concentration of caveolin 1
molecules in some regions of the plasma membrane to trigger the building of
the caveolae scaffold. How this is achieved is not clear but here, we show
that when the concentration of Cav-1 or chimeras reaches a given level (more
than 30%) in the floating fraction of sucrose gradients it is correlated to a
visible effect on caveolae formation. Thus, it is possible that the enrichment
of caveolin molecules into lipid rafts creates a microenvironment that favors
the association of these proteins into a coat and thus into the formation of
invaginated caveolae. It has been shown that Cav-1 binds directly to
cholesterol (Murata et al.,
1995) and this could explain the concentration of this protein in
caveolae that are rich in cholesterol. There is however no data on the
affinity of Cav-2 for cholesterol so far but the fact that it is not highly
enriched in rafts in Caco-2 cells seems to indicate that its binding to
cholesterol is different from the one of Cav-1. Furthermore a recent study has
shown that Cav-2 can be found in lipid droplets while Cav-1 was not in the
same conditions, indicating that the two proteins might have different lipid
affinity (Fujimoto et al.,
2001
). Do caveolae invaginate from lipid rafts? There is no answer
yet to that question. One hypothesis however, is that caveolin molecules when
they reach a local concentration in a given lipid environment can bend the
plane of the membrane to give rise to the pear shape that is characteristic of
these structures. This hypothesis will be tested in vitro using purified Cav-1
with adequate mixes of lipids.
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Acknowledgments |
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