1Departments of Molecular Pharmacology and Medicine, Albert Einstein College of Medicine, and 2The Albert Einstein Cancer Center, Bronx, New York
Submitted 11 May 2004 ; accepted in final form 15 October 2004
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ABSTRACT |
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caveolae; caveolin-1; caveolin-2; caveolin-3
Caveolae are a subset of specialized liquid-ordered domains, referred to as lipid rafts, which are uniquely enriched in various membrane components, such as cholesterol, sphingolipids, and glycosyl phosphatidylinositol-anchored proteins (5, 17, 22). The identification of the principal structural components of caveolae, the caveolin protein family, has provided biochemical markers for the study of these microdomains (32, 37, 50). In recent years it has become well established that caveolae are multifunctional organelles, playing important roles in a variety of cellular processes mainly through interactions with one of the caveolin proteins and a multitude of known signaling molecules, such as heterotrimeric G proteins, Src family tyrosine kinases, H-Ras, endothelial nitric oxide synthase, and the insulin receptor (23, 30, 51).
Caveolar biogenesis, or the formation of invaginations from otherwise flat plasma membrane lipid rafts, has been the subject of intense study. It is well known that this process is highly dependent on at least two molecules, cholesterol and caveolin, because treatments to eliminate either one of these players result in the loss of identifiable caveolae (42). The specific caveolin proteins involved in the formation of caveolae are known to be tissue specific, with caveolin-3 (Cav-3) being expressed in all myocytic cells, whereas Cav-1 and Cav-2 are coexpressed in most other cell types (36, 50). Expression of either Cav-1 or -3 is sufficient to drive caveolae formation, whereas the sole expression of Cav-2 is not only insufficient to support caveolar biogenesis but also insufficient to support its own stability. For example, it has been demonstrated that Cav-2 requires the expression of Cav-1; in the absence of Cav-1, Cav-2 is rapidly degraded by proteasomal mechanisms (28). Both Cav-1 and -3 form functional oligomeric complexes that insert into the plasma membrane, driving the formation of these flask-shaped organelles (29).
Interestingly, smooth muscle cells are the only adult cell type that coexpresses all three caveolin isoforms (Cav-1, -2, and -3) (47). However, only arterial smooth muscle cells coexpress all three caveolin isoforms; in contrast, venous smooth muscle cells specifically lack Cav-3 expression, but retain the coexpression of Cav-1 and -2 (43). On the basis of these findings, Sessa and colleagues (43) have postulated that Cav-3 may have a very specialized function in smooth muscle cells, possibly to maintain the contractile phenotype of arterial vascular smooth muscle cells. In accordance with this hypothesis, uterine smooth muscle cells fail to express detectable levels of Cav-3 in the nonpregnant state (13). However, bladder smooth muscle cells coexpress all three caveolins (53) and perform an essential repeated contractile function.
While the interaction between Cav-1 and -2 has been explored and verified and is now readily accepted, the ability of Cav-3 to interact with either Cav-1 or -2 has remained controversial. Several attempts have been made to analyze the potential Cav-1, -2, and-3 interaction in cell culture models, with varied results. Das and colleagues (2) demonstrated that while Cav-2 can be coimmunoprecipitated with antibodies directed against Cav-1 in Cos-7 cells, similar results could not be obtained with the immunoprecipitation (IP) of Cav-3; thus Cav-2 failed to coimmunoprecipitate with Cav-3 in this cellular context. Furthermore, it has also been shown that whereas transient transfection of Cav-2 causes an upregulation of endogenous Cav-1 in Chinese hamster ovary cells, the same is not true when Cav-3 is transfected (25). This finding supports the contention that Cav-2 interacts with and can stabilize Cav-1, whereas Cav-3 does not form a complex with Cav-1.
More recently, however, findings reported by Rybin et al. (33) challenge this notion, as these authors found Cav-2 expression in cultured cardiac myocytes. They further asserted that previous reports on the subject were premature in dismissing the presence of Cav-2 in myocyte cultures as originating from contaminating fibroblasts, and that Cav-2 was indeed expressed in their cardiac muscle cell cultures. Furthermore, these authors reported that Cav-2 coimmunoprecipitates with Cav-3 as well as cofractionates with Cav-3 by sucrose density centrifugation, thus indicating that these two caveolin proteins are capable of interacting in cultured myocytes (33). While this is the first report showing a Cav-2/-3 interaction, it is in direct contention with most other findings on the subject and thus leaves the issue unresolved.
In the current study, we employ three distinct genetic approaches to analyze the potential Cav-1/-2/-3 interaction. We first fully explored the interaction between Cav-1, -2, and -3 in Cav-1-null 3T3 mouse embryonic fibroblasts (MEFs), a fibroblast cell line deficient in the Cav-1 gene, using a variety of methods. We show that whereas stable transfection of these cells with Cav-3 does not rescue expression of Cav-2, stable transfection with Cav-1 does rescue Cav-2 levels, as predicted. Also, with the use of well-established assay systems, such as sucrose density centrifugation, coimmunoprecipitation, and solubility in cold Triton X-100, we demonstrate that Cav-3 does not interact with either Cav-1 or -2 in these fibroblastic cells.
To further dissect this issue, we next explored potential Cav-1, -2, and -3 interactions in a Cav-3-transfected myoblast cell line. Careful examination reveals that Cav-2, as well as Cav-1, coimmunoprecipitate with Cav-3 in this setting. These observations suggest that an interaction does indeed exist between Cav-1, -2, and -3 and that this interaction is muscle cell-type specific. To extend these observations to an in vivo situation, we next studied skeletal muscle samples from Cav-1 transgenic (Tg) mice, which overexpress Cav-1 in all tissues, including skeletal muscle. Our coimmunoprecipitation experiments clearly demonstrate that Cav-1 does indeed form a tight complex with Cav-3, thus confirming that a muscle-specific interaction occurs between the caveolins.
These data, indicating a cell-type-specific interaction between the caveolin protein family members, hold numerous functional implications for muscle cell development and differentiation (see DISCUSSION).
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MATERIALS AND METHODS |
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MEF culture. Primary MEFs were obtained from day 13.5 embryos and immortalized as previously described (28). Immortalized 3T3 MEFs were grown in complete medium (Dulbecco's modified Eagle's medium) supplemented with 10% fetal bovine serum, 2 mM glutamine, 100 U/ml penicillin, and 100 µg/ml streptomycin (GIBCO-BRL).
Cav-1, -2, and -3 and retroviral infection. The cDNAs encoding full-length Cav-1, -2, and -3 genes were subcloned into the pBabe-Puro retroviral expression vector (21) using a standard PCR-based strategy. It is important to note that these cDNAs were not epitope tagged. Stable infection of 3T3 MEFs and rat L6 myoblasts was conducted essentially as previously described (12, 14). Briefly, pBabe vectors were transiently transfected into the ecotrophic packaging cell line, Phoenix, using a modified calcium phosphate method (12, 14). Forty-eight hours after transfection, the viral supernatant was collected, filtered, and added to the target cells. Two infection cycles were carried out (every 12 h). After the last cycle of infection, cells were selected for 5 days in complete medium containing puromycin at a final concentration of 2.5 µg/ml. Stable expression in the target cell population was confirmed by Western blot analysis.
Transmission electron microscopy. 3T3 MEFs were fixed with 2.5% glutaraldehyde in 0.1 M cacodylate buffer, postfixed with OsO4, and stained with uranyl acetate and lead citrate. After being processed, the samples were examined under a transmission electron microscope (model 1200EX; JEOL) and photographed at a magnification of x16,000. Caveolae were identified by their characteristic flask shape, size (50100 nm), and location at or proximal to the plasma membrane (15, 19).
Immunoblot analysis. Cells were cultured in complete medium and allowed to reach 8090% confluence. Subsequently, they were washed with PBS and incubated with lysis buffer (10 mM Tris, pH 7.5; 50 mM NaCl; 1% Triton X-100; 60 mM n-octylglucoside) containing protease inhibitors (Roche Molecular Biochemicals). When necessary, protein concentrations were determined using the bicinchoninic acid reagent (Pierce). Proteins were separated by SDS-PAGE (12.5% acrylamide) and transferred to nitrocellulose. The nitrocellulose membranes were stained with Ponceau S, followed by immunoblot analysis. For all subsequent washing, the buffers contained 10 mM Tris, pH 8.0, 150 mM NaCl, 0.05% Tween 20, which was supplemented with 1% bovine serum albumin (BSA) and 4% nonfat dry milk (Carnation) for the blocking solution and 1% BSA for the antibody dilution. Horseradish peroxidase-conjugated secondary antibodies were used to visualize bound primary antibodies with the Supersignal chemiluminescence substrate (Pierce).
Triton X-100 insolubility. Extraction of Triton X-100 soluble proteins was performed essentially as we described previously in detail (39, 40). Briefly, cells were grown to confluence in 35-mm-diameter dishes and washed twice with ice-cold PBS. An ice-cold buffer (25 mM Mes, pH 6.5, 150 mM NaCl) containing 1% Triton X-100 plus protease inhibitors (1 ml) was gently added to the cells. After a 30-min incubation period on ice without agitation, the soluble fraction was then collected from the edge of the dish (without scraping the cells) using a 1-ml micropipette (blue tip). An equal volume of 2% SDS (1 ml) was then added to the plate to dissolve the remaining Triton X-100-insoluble material. The SDS fraction was collected using a cell scraper and briefly sonicated to decrease its viscosity. Equal volumes of the Triton X-100 soluble and insoluble fractions were then each separated by SDS-PAGE and subjected to immunoblot analysis, as described above.
Purification of caveolae-enriched membrane fractions. Caveolae enriched membrane fractions were purified essentially as previously described (35, 40). Cells grown to confluence in two 100-mm-diameter plates were washed twice in ice-cold PBS, scraped into 750 µl of Mes-buffered saline (MBS) (25 mM Mes, pH 6.5, 150 mM NaCl) containing 1% Triton X-100, passed five times through a tightly fitting Dounce homogenizer, and mixed with an equal volume of 80% sucrose prepared in MBS lacking Triton X-100. The sample was then transferred to a 4.5-ml ultracentrifuge tube and overlaid with a discontinuous sucrose gradient (1.5 ml of 30% sucrose, 1.5 ml of 5% sucrose, both prepared in MBS lacking detergent). The samples were then subjected to centrifugation at 200,000 g (44,000 rpm in a Sorval rotor TH-660) for 18 h. A light-scattering band was observed at the 5%/30% sucrose interface. Twelve 375-µl fractions were collected, and 50-µl aliquots of each fraction were subjected to SDS-PAGE and immunoblot analysis.
Velocity gradient centrifugation. Velocity gradient centrifugation was conducted as we have described (34, 37). Cells were grown to confluence in 10-cm dishes and dissociated in MBS (25 mM Mes, pH 6.5, 0.15 M NaCl) containing 60 mM n-octylglucoside. Soluble material was loaded on top of a 540% linear sucrose gradient and centrifuged at 50,000 rpm for 10 h in a SW 60 rotor (Beckman). Gradient fractions were collected from above and subjected to immunoblot analysis as described.
Immunofluorescence microscopy. Immunofluorescence was performed as described previously (11). For coimmunolocalization of Cav-2 and -3, Myc-tagged murine Cav-2 was transiently transfected into Cav-1-null cells stably expressing Cav-3 using Lipofectamine Plus (Invitrogen). Forty-eight hours after transfection, cells were processed for immunofluorescence. 3T3 MEFs were grown on glass coverslips, washed three times with PBS, and fixed for 30 min at room temperature with 2% paraformaldehyde. Fixed cells were washed with PBS and permeabilized in washing buffer (PBS containing 0.1% Triton X-100 and 0.2% BSA) for 10 min. The cells were then treated with 25 mM NH4Cl in PBS for 10 min at room temperature to quench any free aldehyde groups. The cells were rinsed with PBS and incubated with the primary antibody for 1 h at room temperature. After three washes with washing buffer (10 min each), the cells were incubated with the secondary antibody for 1 h at room temperature. Finally, the cells were washed three times with washing buffer (10 min each wash) and counterstained with Hoechst-33258 (1 µg/ml) for 15 min to visualize the nucleus. After extensive washing in PBS, slides were mounted with the Slow-Fade reagent (Molecular Probes, Eugene, OR) and observed under a confocal microscope (model MR 600; Bio-Rad). All microscopy was performed at the Analytical Imaging Facility of the Albert Einstein College of Medicine.
Coimmunoprecipitation studies. IP was performed as previously described (41). Briefly, 3T3 MEFs and L6 myoblasts grown to near confluence were washed twice with cold PBS and scraped into 1 ml of IP buffer [10 mM Tris (pH 8.0), 150 mM NaCl, 1% (vol/vol) Triton X-100, 60 mM n-octylglucoside], supplemented with protease inhibitors. For IP from skeletal muscle, mouse tissue was harvested, minced with scissors, homogenized in a Polytron tissue grinder for 30 s, and solubilized in IP buffer containing protease and phosphatase inhibitors. After incubation on ice for 30 min, debris was removed by centrifugation at 13,000 rpm for 10 min. Proteins (500 µg) were precleared by incubation with 30 µl of 1:1 slurry of protein A-Sepharose (Amersham Pharmacia Biotech) for 45 min at 4°C and then transferred to tubes containing fresh protein A-Sepharose and IP buffer. Anti Cav-1 and -3 MAbs were added to the mixture. For coimmunoprecipitation studies, rabbit anti-GFP IgG (FL) or mouse anti-c-Myc IgG were used as negative controls. After a 4-h or overnight incubation at 4°C, immune complexes were collected by centrifugation, washed five times in 1 ml of IP buffer, washed four times with 50 mM Tris (pH 8.0), 150 mM NaCl, 1 mM EDTA, and 1% (vol/vol) Triton X-100, and disrupted by boiling in 1% (wt/vol) SDS. Immune complexes were then resolved by SDS-PAGE and processed for immunoblotting as described above.
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RESULTS |
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However, preliminary studies (28) have indicated that these cells, like other MEF cell lines, are resistant to standard transfection protocols, thus limiting their usefulness. Therefore, we chose to overcome this issue by using a highly efficient retroviral expression vector, which allows for stable integration into the target cells. Thus the full-length untagged cDNAs encoding Cav-1, -2, or -3 were subcloned into the pBabe-puro retroviral expression vector and transfected into the ecotrophic packaging cell line, Phoenix. Virus-containing supernatants were filter-purified from infected cells and used to transduce wild-type and Cav-1-null 3T3 MEFs (12, 14). Stable cell lines were generated by selection in medium containing puromycin and analyzed for integration and expression of the desired vector by Western blot analysis (Fig. 1). Note that similar levels of expression were observed for all three proteins. In addition, stable expression of Cav-3 in wild-type 3T3 MEFs did not affect the expression of endogenous Cav-1 (data not shown).
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Thus we next examined the plasma membrane of confluent transfected 3T3 Cav-1-null MEFs for the presence of caveolae by transmission electron microscopy. As expected, Cav-1-null 3T3 MEFs do not form recognizable caveolae, whereas the introduction of either Cav-1 or Cav-3 is sufficient to restore the formation of these organelles (Fig. 2, arrows). In addition, overexpression of Cav-2 in Cav-1-null 3T3 MEFs is not sufficient to drive caveolae biogenesis (Fig. 2).
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To determine whether Cav-3 could functionally play a similar role in rescuing Cav-2 from degradation, we analyzed the Cav-2 protein content of Cav-1-null 3T3 MEFs recombinantly overexpressing Cav-3. Western blot analysis of these cells demonstrates that overexpression of Cav-3 is not sufficient to rescue Cav-2 protein levels (Fig. 3). Importantly, control experiments, in which Cav-1 was reintroduced into Cav-1-null 3T3 MEFs, demonstrate that Cav-1 overexpression does indeed result in the rescue of Cav-2 (Fig. 3). Western blot analysis for Cav-2 in Cav-2-overexpressing cells is shown for comparison. Note that in this case a near normal level of Cav-2 is achieved; however, Cav-2 is still confined to the Golgi as described below (Fig. 8B).
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To determine whether Cav-3 could functionally substitute for Cav-1 in this regard, we performed Triton X-100 solubility measurements on Cav-1-null 3T3 MEFs stably expressing Cav-3. Figure 4A shows that in these cells, Cav-2 remains in the soluble fraction, whereas in Cav-1-null 3T3 MEFs overexpressing Cav-1, as well as in wild-type MEFs, a major portion of Cav-2 localizes to the Triton X-100 insoluble fraction. As expected, Cav-1 and Cav-3 are both insoluble, thus indicating that Cav-2 follows the behavior of Cav-1, but not Cav-3 (Fig. 4B). However, Cav-3 is not quite as Triton insoluble as Cav-1; this may be due to differences in the primary sequence of Cav-3 that alters its affinity for cholesterol-rich lipid raft membrane domains. Most importantly, these data demonstrate that, in a fibroblast system, Cav-3 does not confer Triton X-100 insoluble characteristics upon Cav-2.
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The results presented in Fig. 5 indicate that Cav-2 correctly targets to caveolae (fractions 5 and 6) in wild-type cells or Cav-1-null 3T3 MEFs overexpressing Cav-1, as expected (Fig. 5, A and C). Furthermore, in Cav-1-null 3T3 MEFs, Western blot analysis for Cav-2 reveals that this protein does not "float" without the presence of Cav-1, as expected (Fig. 5B). In addition, the overexpression of Cav-2 is not sufficient to localize this protein to buoyant membranes (Fig. 5D). In contrast to the results obtained with stable transfection of Cav-1, overexpression of Cav-3 in Cav-1-null 3T3 MEFs demonstrates that Cav-2 is not recruited to caveolae membrane microdomains by Cav-3 (Fig. 5E). Taken together with the results presented above, these data strongly argue that, in 3T3 MEFs, Cav-3 does not interact with Cav-2 in mouse embryonic fibroblasts.
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Oligomerization of Cav-2 is partially rescued by the introduction of Cav-3 in Cav-1-null 3T3 MEFs. Another important property of the caveolin proteins is their ability to form functional oligomeric complexes (34). After synthesis in the endoplasmic reticulum, Cav-1 can form either homo-oligomers or hetero-oligomers with Cav-2 composed of 1416 individual caveolin monomers. Cav-3 normally forms only homo-oligomeric complexes, as it is generally thought to be the sole caveolin expressed in muscle cells. In the Golgi apparatus, adjacent hetero-oligomers of Cav-1 and -2 undergo a second stage of oligomerization through contacts between the COOH- terminal domains of Cav-1 proteins (2, 36, 39, 48). This second step of oligomerization leads to the formation of a caveolin-rich scaffold. Studies (2, 36, 39, 48) employing site-directed mutagenesis have demonstrated that residues 61101 of Cav-1 are involved in the first step of oligomerization, whereas residues 168178 are involved in the second step. It remains unknown however, whether Cav-3, which shares 85% homology to Cav-1, can induce the formation of Cav-2-containing high molecular mass complexes. The oligomeric state of any caveolin protein can be assessed using a well-established velocity gradient centrifugation system (34, 37, 50).
Here, we utilize this system to determine whether overexpression of Cav-3 in Cav-1-null 3T3 MEFs can lead to the formation of oligomeric Cav-2-containing complexes. Figure 6, A and C, shows that Cav-2 is found in high molecular mass oligomers of 200400 kDa (fractions 6 and 7) in presence of Cav-1 (in wild-type 3T3 MEFs and Cav-1-overexpressing Cav-1-null 3T3 MEFs), as expected. In the absence of Cav-1, endogenous Cav-2 fails to form high molecular mass oligomers, also as expected (Fig. 6B). In addition, overexpression of Cav-2 is not sufficient to induce the formation of high molecular mass oligomers (Fig. 6D). Interestingly, we observe that recombinant expression of Cav-3 in Cav-1-null 3T3 MEFs partially restores the ability of endogenous Cav-2 to form high molecular mass oligomers (Fig. 6E).
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Neither Cav-1 nor -2 coimmunoprecipitates with Cav-3 in 3T3 MEFs. To further explore possible interactions between the three caveolin proteins, we next performed coimmunoprecipitation studies. Protein lysates were prepared from wild-type, untransfected Cav-1-null, and Cav-1-null 3T3 MEFs overexpressing each of the caveolin proteins. Lysates were immunoprecipitated with either an anti-Cav-1 or anti-Cav-3 IgG and then subjected to SDS-PAGE and subsequent immunoblot analysis with antibodies directed against Cav-2.
The results presented in Fig. 7A demonstrate that endogenous Cav-2 is coimmunoprecipitated with antibodies directed against Cav-1 in wild-type 3T3 MEFs, as well as in Cav-1-null 3T3 MEF stably overexpressing Cav-1. However, in striking contrast, Cav-2 does not coimmunoprecipitate when an antibody directed against Cav-3 is used in Cav-1-null 3T3 MEFs overexpressing Cav-3. This again indicates that Cav-2 and -3 do not interact in this fibroblast system. As important negative controls, protein A-sepharose beads alone or an irrelevant IgG antibody showed no coimmunoprecipitation of Cav-2 (Fig. 7A).
In this experiment, we also evaluated a possible interaction between Cav-1 and -3 by coimmunoprecipitation. Protein lysates from wild-type 3T3 MEFs and wild-type 3T3 MEFs recombinantly expressing Cav-3 were immunoprecipitated with monoclonal antibodies directed against either Cav-1 or -3. Western blot analysis showed that no interaction between Cav-1 and -3 is detectable (Fig. 7B). Thus, in the setting of a fibroblast cell line, Cav-1 does not interact with Cav-3.
Recombinant expression of Cav-3 in Cav-1-null 3T3 MEFs does not rescue the intracellular retention of Cav-2. Because it has been shown that the absence of Cav-1 causes intracellular retention of Cav-2 in Cav-1-null 3T3 MEFs (28), we next attempted to determine the subcellular localization of Cav-2 in wild-type 3T3 MEFs and Cav-1-null 3T3 MEFs recombinantly overexpressing Cav-1, -2, or- 3. In Fig. 8A, we show that in wild-type cells, Cav-2 and -1 follow a similar expression pattern, indicative of colocalization. Furthermore, loss of Cav-1 causes a characteristic redistribution of Cav-2 to the perinuclear region (see nuclei, labeled as N). We further demonstrate that reintroduction of Cav-1 in Cav-1-null 3T3 MEFs restores the plasma membrane localization of Cav-2 (Fig. 8B). In addition, Fig. 8B demonstrates that overexpression of Cav-2 does not compensate for the absence of Cav-1, as Cav-2 staining remains perinuclear.
We next examined the subcellular localization of Cav-2 in Cav-1-null 3T3 MEFs recombinantly overexpressing Cav-3. In these studies, Myc-tagged Cav-2 was transiently transfected into Cav-3-overexpressing Cav-1-null 3T3 MEFs. These cells were then fixed and immunostained with antibodies directed against the Myc epitope tag and Cav-3. Consistent with the biochemical data above, we find that Cav-3 is correctly targeted to the plasma membrane, whereas Cav-2 is retained in the perinuclear compartment (Fig. 8C). These data again confirm that expression of Cav-3 is not sufficient to confer membrane targeting upon Cav-2 in mouse embryonic fibroblasts. In this cell, the nucleus is labeled with Hoechst-33258 (blue). Identical experiments employing the transient expression of Cav-1 demonstrated that Myc-tagged Cav-2 was properly targeted to the plasma membrane (data not shown).
To assess the relationship between Cav-1 and -3, we next sought to determine the subcellular localization of these two proteins in wild-type 3T3 MEFs recombinantly overexpressing Cav-3. Figure 8D demonstrates that when coexpressed, Cav-1 and Cav-3 show a diffuse overlapping distribution with extensive colocalization at the level of the plasma membrane, indicating that they both colocalize to lipid raft microdomains at the cell surface.
Cav-1, -2, and -3 interact in undifferentiated L6 myoblasts recombinantly expressing Cav-3.
Because all of the findings discussed above were made in a fibroblast cell line, we next decided to verify these data in a different cellular system. For this purpose, we chose the L6 myoblast cell line because, under proliferating conditions, these cells coexpress Cav-1 and Cav-2, but not Cav-3. Induction of Cav-3 expression generally occurs when this cell line is forced to differentiate by changes in serum media concentrations, such as serum starvation. Therefore, we generated an L6 myoblast cell line recombinantly overexpressing Cav-3 (termed L6/Cav-3 cells) by using standard retroviral techniques (Fig. 9A) similar to those described above for the 3T3 MEFs. Interestingly, recombinant expression of Cav-3 in L6 cells causes a 2-fold increase in the expression levels of endogenous Cav-2, suggesting that Cav-3 expression stabilizes the Cav-2 protein product. However, no changes in Cav-1 levels were observed.
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To examine whether the introduction of recombinant Cav-3 could affect the caveolar targeting of the endogenous Cav-1 and Cav-2, we next performed sucrose density centrifugation. For this purpose, we compared the behavior of L6 cells, with and without Cav-3 expression (L6 vs. L6/Cav-3). The results presented in Fig. 9C clearly demonstrate that recombinant expression of Cav-3 in L6 myoblasts increases the caveolar localization (see especially fraction 5) of endogenous Cav-1 and Cav-2.
Next, using velocity gradient centrifugation, we analyzed the oligomeric state of endogenous Cav-1 and Cav-2 in the presence of recombinant Cav-3. Again, we directly compared the behavior of L6 cells, with and without Cav-3 expression (L6 vs. L6/Cav-3). Figure 9D shows that the presence of Cav-3 favors the oligomerization of endogenous Cav-1 and Cav-2. Note the disappearance of the oligomers from fractions 5 and 6 in L6/Cav-3 cells. These results provide independent support for our coimmunoprecipitation studies, showing that all three caveolins (Cav-1, -2, and -3) form a stable complex in L6 cells.
Cav-1 and Cav-3 colocalize in proliferating L6/Cav-3 cells. To further examine the interaction between the caveolins, we next performed immunofluorescence microscopy on L6/Cav-3 cells to determine the subcellular localization of these proteins. Cells were fixed and dually immunostained with antibodies against Cav-1 and Cav-3. Microscopic analysis of these cells demonstrates the well-defined colocalization of Cav-1 and -3, at the level of the plasma membrane (Fig. 10). Therefore, in undifferentiated myoblasts, like in the mouse embryonic fibroblasts above, Cav-1 and Cav-3 are directed to the plasma membrane. More important, however, are the findings that all three caveolin proteins interact and are part of the same complex, and that this interaction occurs only in muscle cells.
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Thus we next focused our attention on this Tg mouse model to extend our findings with L6 myoblasts, i.e., that Cav-1 and -3 interact in a muscle-specific manner. Skeletal muscle extracts from wild-type and Cav-1 Tg mice were prepared and subjected to IP with an anti-Cav-1 antibody or an anti-Cav-3 antibody. Immunoblot analysis of the resultant protein complexes reveals that Cav-1 and Cav-3 coimmunoprecipitate with antibodies directed against either Cav-1 or Cav-3 (Fig. 11). Thus these in vivo data verify our results obtained in L6/Cav-3 myoblasts and further indicate that Cav-1 and Cav-3 are capable of interacting, but that this interaction is dependent on the cell type in which the two proteins are coexpressed. One limitation of this system, however, is that skeletal muscle does not normally express Cav-2, and thus we were unable to ascertain whether this protein also interacts with Cav-3 in this setting. Thus complex formation between Cav-1 and Cav-3 does not require coexpression with Cav-2.
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DISCUSSION |
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We next attempted to extend these results in a different cellular system using L6 myoblasts recombinantly overexpressing Cav-3 that also express endogenous Cav-1 and -2. In these cells, coimmunoprecipitation experiments demonstrate that, contrary to our results in fibroblasts, Cav-1, -2, and -3 all coimmunoprecipitate. These data indicate that a muscle cell-type-specific interaction occurs between the caveolin protein family members. To confirm these findings in vivo, we next performed similar experiments on Cav-1 Tg mouse skeletal muscle samples. In these experiments, we demonstrate that Tg Cav-1 and endogenous Cav-3 do indeed coimmunoprecipitate. Therefore, this study shows for the first time that a muscle-specific interaction occurs between Cav-1, -2, and -3.
The tissue distribution pattern of the Cav-1, -2 and -3 has long been the subject of intense study. While no comprehensive report has ever been published, a careful review of the literature provides a consensus indicating that Cav-1 and -2 are found in most cell types with the exception of myocytes, whereas Cav-3 is confined to myocytic cells (36, 50). It is well known that both Cav-1 and -3 are capable of driving caveolae formation, whereas Cav-2 does not possess this property. Cav-2 is also known to require Cav-1 as a chaperone protein to localize it to the plasma membrane, because ablation of Cav-1 results in Golgi retention and the rapid proteasomal degradation of Cav-2 (28). With the recent publication of findings indicating that Cav-2 and -3 are coexpressed in ventricular myocardium (33), the question of whether Cav-3 can also act as a molecular chaperone for Cav-2 has taken on new importance. Sequence analysis of Cav-1 and Cav-3 reveals that these two proteins share 85% homology and 65% identity as well as an identical COOH-terminal palmitoylation pattern (3). However, it has previously been shown in Chinese hamster ovary and Cos-7 cells that Cav-2 and Cav-3 do not interact as determined by coimmunoprecipitation experiments (2, 25). To further address this issue, we undertook the present study using molecular genetic approaches in both myocytes and fibroblasts. Thus we are able to clearly demonstrate that the interaction between the caveolin proteins is cell-type specific.
Cav-2 has long been considered an accessory protein for Cav-1, with little ascribed functional role. In fact, the recent generation of Cav-1- and Cav-2-null mice has shown that all but one of the phenotypes recognized in the Cav-1-null mouse are due to a loss of Cav-1, not the subsequent Golgi retention and degradation of Cav-2 (27, 28, 31). However, the role of Cav-2 in the myocardium has not been addressed, as it was thought that this protein was not expressed in this tissue. The presence of Cav-2 in isolated cardiac myocytes has now been confirmed (33). Yet, as Cav-2 remains trapped in the Golgi complex without a molecular chaperone and it has been reproducibly shown that Cav-2 and -3 do not interact, it does not follow that these two proteins should be coexpressed in myocytes. Our new data resolve this conundrum, providing evidence that Cav-3 can interact with Cav-2, but only when these two proteins are coexpressed in myocytes.
What is the nature of the muscle-specific interaction of all three caveolin isoforms? One possibility is that a muscle-specific accessory protein acts as a bridge to allow the interaction of Cav-3 oligomers with Cav-1/2 hetero-oligomers. Alternatively, a muscle-specific chaperone protein may allow the hetero-oligomerization of all three caveolins (Cav-1, -2, and -3) at the level of the endoplasmic reticulum/Golgi. In this case, we would envision a transient interaction and the chaperone would not be part of the final caveolin hetero-oligomeric complex. Finally, in either case, such a factor is probably expressed in myoblasts before terminal muscle differentiation/fusion, as recombinant expression of Cav-3 in undifferentiated L6 myoblasts allows the formation of mixed caveolin oligomers (containing Cav-1, -2, and -3). However, this factor is apparently not expressed in fibroblastic cells. Thus our current studies provide a systematic basis for identifying such a muscle-specific accessory protein(s) or chaperone(s) that mediate caveolin hetero-oligomerization.
These data, indicating a cell-type-specific interaction between the caveolin protein family members, hold numerous functional implications for muscle cell development and differentiation. For instance, it is well known that caveolin protein family expression is limited to Cav-1 and -2 in cultured skeletal myoblasts during the proliferative portion of the life cycle (26). Conversely, in terminally differentiated myocytes, Cav-1 and -2 protein expression is nearly undetectable (26), whereas Cav-3 is highly expressed (47). However, during a significant portion of the differentiation process, from myoblast to myocyte, all three caveolin proteins are detectable in the same cell (7, 26). Perhaps, rather than just coincidental coexpression, the concomitant expression of all three caveolin protein family members may play a necessary role in muscle cell development and the terminal differentiation process.
In further support of the findings outlined here, it has recently been shown that in smooth muscle cells of the murine bladder, Cav-1, -2, and -3 are all coexpressed and can be coimmunoprecipitated (53). While it has long been thought that all three caveolin proteins were expressed in this cell type, the functional significance of this phenomenon remained elusive. However, in this report, it was also shown that a selective loss of Cav-3 (as in Cav-3-null mice) leads to a 2-fold increase in morphologically identifiable plasmalemmal caveolae in bladder smooth muscle cells (53). This finding strongly indicates that Cav-3 coexpression may normally act to suppress Cav-1-driven caveolar invagination in smooth muscle cells. Conversely, a selective loss of Cav-1 (as in Cav-1-null mice) leads to a dramatic reduction (
6.5-fold) of plasmalemmal caveolae in bladder smooth muscle cells (53). In contrast, ablation of Cav-2 expression (as in Cav-2-null mice) had little or no effect on caveolae formation, assessed morphologically. Finally, dual ablation of Cav-1 and Cav-3 expression (as in Cav-1/-3 double-knockout mice) was required for the complete ablation of caveolae in these smooth muscle cells.
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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.
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