CD44 as a Marker for CLIC/GEEC Endocytosis

Various markers have been used to follow CIE pathways but CD44 has emerged as a highly specific cargo of the CLIC/GEEC pathway [29],[40]. We therefore first optimized and validated the conditions for quantitatively studying CIE using antibodies against CD44 as a marker. An anti-CD44 monoclonal antibody (mAb) was added together with fluorescent transferrin (Tfn-647) to cells for 2 and 10 min at 37°C. Prior to fixation, cells were placed on ice and acid stripped to remove any residual surface label, and internalized CD44 mAb was labeled with fluorescently tagged secondary antibody. For quantitative analysis of internalization, the fluorescence intensity of the internalized markers (over the entire cell) was normalized against the average fluorescence intensity of the internalized markers in control samples. By the following criteria, this procedure allowed us to use CD44 as a specific marker of the CLIC/GEEC pathway: (1) Anti-CD44 mAb internalization was distinct from uptake via clathrin coated pits, labeled with Tfn-647, and did not colocalize with caveolae, indicated by probing for endogenous caveolar proteins, CAV1 and Cavin-1 (Figure 1A). (2) CD44 mAb uptake was dynamin-independent, as determined by the use of the small molecule inhibitor dynasore. Under our experimental conditions, dynasore treatment inhibited Tfn-647 uptake but had no effect on CD44 mAb internalization (Figure 1B) [29]. (3) Uptake of CD44 mAb was completely inhibited by 7-ketocholesterol (7-KC) treatment, shown in previous studies to result in reduced membrane order [41], while Tfn-647 uptake was not significantly affected at this concentration (Figure 1B). These results confirmed that in our system anti-CD44 antibody was internalized via the cholesterol-dependent, dynamin-independent CLIC/GEEC pathway [23],[30] (Figure 1B). Internalized anti-CD44 mAb therefore represents a specific marker of the CLIC/GEEC pathway that does not require a low temperature prebinding step and does not affect the kinetics or the magnitude of the pathway being studied [29]. Specific internalization of anti-CD44 mAb was confirmed by using a control antibody (anti-GFP mAb) at the same concentration in wild type mouse embryonic fibroblast cells (WT MEFs) and with anti-CD44 mAb in COS-7 cells, which lack CD44 receptor on the cell surface [42]. Under both conditions, no internalized antibody could be detected after 2 min at 37°C whereas COS-7 cells expressing CD44-GFP showed specific uptake of the anti-CD44 mAb (Figure S1A,B). Additionally, using live cell imaging, we also observed colocalization between CD44-GFP and the fluid phase marker (Dex-647), suggesting that CD44-GFP is internalized via the CLIC pathway (Figure S2, Movie S1). These internalization conditions were subsequently used to investigate the crosstalk between the caveolar and the CLIC/GEEC endocytic systems.

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Figure 1. CD44 as a marker of CLIC/GEEC endocytosis.

(A) WT MEFs were incubated with anti-CD44 mAb and Tfn-647 for 2 min and 10 min at 37°C. Cells were placed on ice and acid washed before fixation. Internalized anti-CD44 mAb was labeled with anti-mouse Alexa Fluor-555 (AF-555) secondary antibody. For labeling endogenous CAV1 and Cavin-1, respective primary antibodies were used, followed by respective secondary AF antibodies. (B) WT MEFs were treated with 60 µM dynasore and 30 µM 7-KC respectively or left untreated (control) for 30 min at 37°C prior to uptake of endocytic markers as mentioned in (A). The bar graph represents the quantitation of internalized endocytic markers. (C) Internalization assay with anti-CD44 mAb and Tfn-647 was performed in WT, CAV1−/−, and Cavin-1−/− MEFs for 2 min at 37°C as mentioned in (A). Arrows indicate CD44-labeled puncta. (D) 40–50 cells from (C) were quantified for normalized fluorescent intensity of endocytic markers. (E) Whole cell lysates from WT, CAV1−/−, and Cavin-1−/−MEFs were immunoblotted for CD44. A representative immunoblot is shown in which actin was used as a loading control. The bar graph represents quantitation of CD44 protein levels calculated by measuring band intensities, by densitometry, in WT, CAV1−/−, and Cavin-1−/− MEFs from three independent experiments. (F) Both nonpermeabilized and permeabilized WT, CAV1−/−, and Cavin-1−/− MEFs were labeled with primary anti-CD44 mAb followed by secondary AF-488 labeling and Phalloidin-AF-596 (F-actin, not shown) as an internal control. 50–60 cells from each condition and cell type were quantified for average fluorescent intensity ratio of CD44 and F-actin (CD44:F-Actin). The graph represents ratio of CD44:F-actin values obtained in nonpermeabilized (surface) cells normalized by the corresponding value obtained in permeabilized (total) cells. (G) Internalization assay with Dex-488 was performed in WT, CAV1−/−, and Cavin-1−/− MEFs for 5 min at 37°C. 40–50 cells from each cell type were quantified for normalized fluorescent intensity of internalized Dex-488. In (F) data represent mean ± SEM of data pooled from three independent experiments, and statistical significance was calculated by one-way ANOVA analysis. In (B, D, E, G) data represent mean ± SEM of three independent experiments. *p<0.02,**p<0.01,***p<0.001,****p<0.0001 (two-tailed t-test). Scale bar: 10 µm.

https://doi.org/10.1371/journal.pbio.1001832.g001

The CLIC/GEEC Endocytic Pathway is Up-regulated in CAV1 and Cavin-1 Null Cells

Transient CAV1 over-expression, which leads to caveola formation in cells that express Cavin-1, has been shown to inhibit CIE [23],[39],[43]–[46]. To investigate whether this reflects a general role of caveolar proteins in the CLIC/GEEC pathway, we used CAV1 null (CAV1−/−) and Cavin-1 null (Cavin-1−/−) MEFs (note these cells also have lower Cavin-1 and CAV1 protein levels, respectively, and so do not allow discrimination of specific effects of loss of either protein (Figure S3A) [18]). We compared the constitutive uptake of CD44 mAb and Tfn-647 in WT, CAV1−/−, and Cavin-1−/− MEFs after 2 min at 37°C; at this early time point of internalization, peripheral labeling for CD44 mAb was observed, consistent with a previous study [29] (Figure 1C). A significant 2-fold increase in anti-CD44 mAb fluorescence, as a measure of CLIC internalization, was observed in CAV1−/− and Cavin-1−/− MEFs in comparison with WT MEFs (Figure 1D). However, there was no significant effect on Tfn-647 uptake, indicating that CME was not affected (Figure 1C,D). Additionally, CD44 mAb and Tfn-647 uptake was also performed in CAV1−/− MEFs expressing Cavin-1-specific siRNA to characterize the functional consequences, if any, of loss of Cavin-1 in CAV1−/− MEFs. No significant difference was observed in either CD44 mAb or Tfn-647 uptake between Cavin-1 siRNA (80% knock down) and control siRNA-transfected CAV1−/− cells (Figure S3B). We also examined whether the increase in anti-CD44 mAb internalization was related to altered CD44 protein expression levels in WT, CAV1−/−, and Cavin-1−/− MEFs. Western blot analysis showed no statistically significant increase in the expression level of CD44 in CAV1−/− MEFs but an increase in Cavin-1−/− MEFs when compared with WT cells (Figure 1E). However, quantitative immunofluorescence (IF) analysis showed no significant difference in the ratio of surface/total levels of anti-CD44 mAb labeling between WT, CAV1−/−, and Cavin-1−/− MEFs (Figure 1F).

Altogether, these results suggest that the CLIC/GEEC pathway may be upregulated in the absence of caveolar proteins. We therefore investigated the effect of loss of caveolar proteins on internalization of the fluid phase marker fluorescent dextran (Dex-488), which is partially internalized through the CLIC/GEEC pathway [23],[26]. Dex-488 was internalized at 37°C for 5 min to provide sufficient signal-to-noise ratio. Similar to CD44 mAb uptake, a significant increase in Dex-488 internalization was observed in CAV1−/− and Cavin-1−/− MEFs in comparison with WT MEFs (Figure 1G). Together, these data suggest that lack of caveolae, through two distinct genetic manipulations, causes upregulation of the CLIC/GEEC pathway. Thus, we next investigated the role of specific caveolar proteins in this regulation.

Caveolae, Caveolin-1, and Cavin-1 Inhibit Uptake via the CLIC/GEEC pathway by Independent Mechanisms

We restored the levels of CAV1 and Cavin-1 in CAV1−/− MEFs by expressing the fluorescent protein (YFP/GFP) tagged constructs, and then compared uptake of the CD44 mAb (2 min at 37°C) between transfected and untransfected cells (Figure 2A; Figure S4). We observed 30–40% transfection efficiency for almost all transient transfections and only low transgene-expressing cells were analyzed. CD44 mAb endocytosis was drastically reduced in CAV1-expressing cells (Figure 2B). More surprisingly Cavin-1 expressing CAV1−/− MEFs also showed a dramatic decrease in CD44 mAb endocytosis (Figure 2C). As CAV1−/− cells lack morphological caveolae [23] this strongly suggests that Cavin-1 inhibits CLIC endocytosis independent of CAV1 and caveolae. Additionally, exogenous expression of Cavin-1 in Cavin-1−/− MEFs was also observed to cause a significant decrease in CD44 mAb uptake (Figure 2D). Neither CAV1 nor Cavin-1 expression affected Tfn-647 uptake. Compared with untransfected cells, CAV1 over-expression resulted in 95±0.3% inhibition of CD44 mAb endocytosis, while Cavin-1 caused a 70±2.2% inhibition (Figure 2B,C). Since loss of caveolar proteins resulted in an increase in Dex-488 uptake, we also investigated the effects of reconstituted CAV1 and Cavin-1 expression in CAV1−/− MEFs on Dex-488 uptake (5 min at 37°C). Similar to CD44 mAb uptake, expression of either CAV1 or Cavin-1 resulted in a significant decrease in Dex-488 uptake (Figure S5A,B; 40±7% inhibition in CAV1-expressing cells; 44±6% inhibition in Cavin-1 expressing cells) although not as high inhibition as observed with the specific CLIC marker, CD44 mAb. We also characterized the effect of other members of the caveolin family, CAV2 and CAV3, on the CLIC/GEEC pathway. The ectopic expression of CAV3, but not CAV2, inhibited CD44 mAb uptake in CAV1−/− MEFs, while Tfn-647 uptake was not affected by expression of either of the proteins (Figure 2E,F). To test whether inhibition of the CLIC/GEEC pathway activity upon expression of caveolar proteins in our system (CAV1−/− MEFs) is due to inhibition of CLIC/GEEC carrier formation, we used electron microscopy (EM). CAV1−/− MEFs were transiently transfected with CAV1-YFP, Cavin-1-GFP, and with GFP alone respectively and then the fluid phase marker horseradish peroxidase (HRP) was added for 2 min at 37°C to label the putative CLIC/GEEC carriers [23],[29]. In comparison with cells expressing GFP alone, a significant decrease was observed in the number of CLICs in CAV1 and Cavin-1 transfected cells (GFP: 100%; CAV1: 54±4% decrease; Cavin-1: 55±8% decrease; mean ± SEM), indicating that expression of caveolar proteins can limit the formation of carriers of the CLIC/GEEC pathway (Figure 2G).

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Figure 2. Caveolar proteins specifically down-regulate CLIC endocytosis.

(A) In CAV1−/− MEFs transiently expressing CAV1-YFP and Cavin-1-GFP respectively, internalization assay was performed with anti-CD44 mAb and Tfn-647 for 2 min at 37°C. Cells were acid washed prior to fixation, and internalized anti-CD44 mAb was labeled with AF-555 secondary antibody. Arrows indicate CD44-labeled puncta. (B,C) 40–50 cells of each transfection from (A) were quantified for normalized fluorescent intensity of endocytic markers. (D) Cavin-1−/− MEFs were transiently transfected with Cavin-1-GFP, and uptake and quantification of internalized anti-CD44 mAb and Tfn-647 were performed as mentioned in (A,B). (E,F) CAV1−/− MEFs were transiently transfected with Myc-CAV2 and RFP-CAV3 respectively. Uptake and quantification of internalized anti-CD44 mAb and Tfn-647 were performed as mentioned in (B). (G) In CAV1−/− MEFs transiently expressing GFP vector, CAV1-YFP, and Cavin-1-GFP respectively, constitutive internalization of HRP (10 µg/ml) was performed for 2 min at 37°C followed by DAB reaction on live cells. Cells were fixed, embedded, and sectioned. Quantification of HRP-labeled CLICs per cell, using 16-20 cells from each transfection, is shown. For more images representing experimental variation obtained for uptake analysis, performed in untransfected and CAV1-YFP/Cavin-1-GFP-expressing CAV1−/− MEFs, see supporting information Figure S12, Figure S13, and Figure S14. In (B–G) data represent mean ± SEM of three independent experiments. *p<0.05,***p<0.0005,****p<0.0001 (two-tailed t-test). Scale bar: 10 µm.

https://doi.org/10.1371/journal.pbio.1001832.g002

These results suggested that both caveolins (CAV1 and CAV3) and Cavin-1 are capable of negatively regulating the CLIC/GEEC pathway. This raised the question of whether inhibition is caused exclusively by free caveolar proteins, independent of caveolar formation. We addressed this question by testing whether inhibition of the CLIC/GEEC pathway occurred when levels of caveolin and cavin were balanced, as indicated by their colocalization and by the immobilization of CAV1 within caveolae. Cavin-1−/− MEFs were transfected either with CAV1-YFP or with Cavin-1-mCherry alone or with both constructs simultaneously. Both CAV1 and Cavin-1 proteins were expressed in similar amounts in single and double transfections, as confirmed by quantitative Western blot analysis (Figure 3A). Moreover, confocal images showed negligible levels of cytosolic Cavin-1 present in CAV1 and Cavin-1 co-expressing cells, and quantitative colocalization analysis showed a significantly higher degree of colocalization between CAV1 and Cavin-1 at the PM in comparison with analysis of randomized pixels for the same regions (Pearson coefficient, PM: 0.80±0.02; random: −0.007±0.005; mean ± SEM, n = 40 cells, p<0.0001) (Figure 3B). In the same set of cells, anti-CD44 mAb and Tfn-647 uptake studies were performed. A significant decrease in CD44 mAb endocytosis was observed in cells co-transfected with CAV1 and Cavin-1, whereas Tfn-647 uptake was not affected (Figure 3C). To gain a quantitative estimation of free caveolar proteins present in our system upon co-expression of CAV1 and Cavin-1, we made use of fluorescence recovery after photobleaching (FRAP) to analyze the mobility and diffusion properties of CAV1. Cavin-1−/− MEFs were transiently transfected with CAV1-YFP or co-transfected with CAV1-YFP and Cavin-1-mCherry respectively. After photobleaching a defined region of interest (ROI) at the PM, we compared the rate of fluorescence recovery of CAV1 between the single and co-transfected cells. A significant decrease in the mobile fraction (CAV1: 0.82±0.03; CAV1+Cavin-1: 0.58±0.05, p<0.05) and lateral diffusion (CAV1: 0.13±0.08; CAV1+Cavin-1: 0.039±0.002; mean ± SD, n = 15 cells, p<0.001, see Text S1) of CAV1 was observed when co-expressed with Cavin-1, in comparison with expression alone (Figure 3D), suggesting that most of the CAV1 in the presence of Cavin-1 had been immobilized through the formation of caveolae. Taken together, these results suggest that the caveolar proteins, independent of caveolae, can inhibit CLIC endocytosis, but in addition, when incorporated into caveolae, there is also potent inhibition of CLIC endocytosis.

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Figure 3. Caveolae generation down-regulates CLIC endocytosis.

(A) Cell lysates from untransfected Cavin-1−/− MEFs or Cavin-1−/− MEFs transiently transfected with CAV1-YFP or Cavin-1-mCherry or co-transfected with CAV1-YFP and Cavin-1-mCherry were immunoblotted for CAV1 and Cavin-1. Actin was used as loading control. For quantification, the integrated intensity of protein bands was calculated. (B) Representative image of cell from (A) showing colocalization between CAV1 and Cavin-1 at different subcellular locations. Insets depict CAV1, Cavin-1 and merged labeling at the plasma membrane and cytosolic regions, marked by the square. 40 cells from (A) were subjected to Pearson correlation analysis, and a representative graph is shown. (C) Cells from (A), growing on coverslips, were subjected to internalization assay with anti-CD44 mAb and Tfn-647 for 2 min at 37°C. Cells were acid washed prior to fixation and internalized anti-CD44 mAb was labeled with AF-555 secondary antibody. 40–50 cells were quantified for normalized fluorescent intensity of endocytic markers. (D) FRAP analysis for CAV1 diffusion at the plasma membrane of Cavin-1−/− MEFs expressing CAV1-YFP alone or co-expressing CAV1-YFP and Cavin-1-mCherry. Fluorescence recovery curve and lateral diffusion analysis (inset) from 15 cells (mean ± SD) are shown. In (B) data represent mean ± SEM of data pooled from three independent experiments and in (C) data represent mean ± SEM of three independent experiments. ***p<0.0005,****p<0.0001 (two-tailed t-test). Scale bar: 10 µm.

https://doi.org/10.1371/journal.pbio.1001832.g003

Cavins 1 and 3, but Not 2 and 4, Inhibit the CLIC/GEEC Pathway

To further investigate the unexpected regulatory role of cavins in CLIC/GEEC endocytosis, we analyzed the effects of ectopic expression of each of the four members of the cavin family (Figure S6A) on CD44 mAb internalization. In CAV1−/− MEFs, moderate over-expression of Cavin-1 and Cavin-3 caused a significant decrease in CD44 mAb endocytosis, whereas in Cavin-2 and Cavin-4-expressing cells no effect on CD44 mAb endocytosis was observed (Figure 4A). Expression of both Cavin-1 and Cavin-3 resulted in significant inhibition (Cavin-1: 77.9±3.3% decrease; Cavin-3: 76.9±3.9% decrease; mean ± SEM) of CD44 mAb endocytosis when compared with control cells (100±13%; Figure 4B). These results reaffirmed the noncaveolar role of Cavin-1 as a negative regulator of CLIC/GEEC endocytosis. Furthermore, Cavin-3 was also observed to have a CAV1- and caveola-independent regulatory role in CLIC/GEEC endocytosis.

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Figure 4. Cavin-1 and Cavin-3, but not Cavin-2 and Cavin-4 negatively regulate CLIC endocytosis.

(A) CAV1−/− MEFs were transiently transfected with pIRES-GFP, pIRES-Cavin-1, pIRES-Cavin-2, pIRES-Cavin-3, and pIRES-Cavin-4 respectively and internalization assay was performed with anti-CD44 mAb and Tfn-647 for 2 min at 37°C. Cells were acid washed prior to fixation and internalized anti-CD44 mAb was labeled with AF-555 secondary antibody. (B) 40–50 cells from each transfection were quantified for normalized fluorescent intensity of endocytic markers. (C,D) 3T3-L1 cells were transiently transfected with siRNA directed specifically to Cavin-1 and Cavin-3 respectively. Uptake and quantification of internalized marker were performed as mentioned in (A,B). In (B–D) data represent mean ± SEM of three independent experiments. *p<0.05,**p<0.005,****p<0.0001 (two-tailed t-test). Scale bar: 10 µm.

https://doi.org/10.1371/journal.pbio.1001832.g004

We next assessed the effect of reducing cavin protein levels on the CLIC/GEEC pathway using a siRNA directed approach in 3T3-L1 fibroblasts, as we obtained more efficient knock down in this cell type compared with MEFs. Reduction of Cavin-1 levels (by approximately 80%) and Cavin-3 levels (by approximately 50%) resulted in an increase in CD44 mAb uptake in comparison with control siRNA-treated cells (Figure 4C,D; Figure S6B). Reduction of Cavin-3 levels also led to an increase in Tfn-647 uptake while reduction in Cavin-1 did not, suggesting Cavin-3 might influence CME.

The Scaffolding Domain of CAV1 is Essential and Sufficient for Inhibition of the CLIC/GEEC pathway

To characterize the regulatory mechanism by which CAV1 inhibits the CLIC/GEEC pathway, we investigated the role of caveolin scaffolding domain (CSD) of CAV1, as this domain has been shown to mediate several regulatory roles of CAV1 [47],[48]. We tested CAV1 and CAV3 CSD point substitution mutants (CAV1G83S; CAV3G55S) and the deletion mutant (CAV1Δ80–100) as used in previous studies [43]. Expression of the CSD mutants, unlike the WT proteins, did not inhibit CD44 mAb internalization in CAV1−/− MEFs (Figure 5A–C), suggesting that the CSD is required for inhibition of the CLIC/GEEC pathway. We next investigated whether the CSD is sufficient for inhibition by expressing the minimal CAV1 scaffolding domain as a fusion protein with GFP (CAV1-SD, amino acid 82–101) in CAV1−/− MEFs. CD44 mAb uptake was significantly decreased (58±3% decrease; mean ± SEM) in cells expressing CAV1-SD while Tfn-647 uptake was unaffected (Figure 5D). Taken together, these results suggest that inhibition of the CLIC/GEEC pathway by caveolin proteins requires an intact scaffolding domain and that this domain alone has significant inhibitory activity on CLIC endocytosis. The fact that the scaffolding domain has inhibitory activity when expressed in cells lacking endogenous CAV1 shows that the inhibitory activity of the mutant is not mediated through inhibition of interactions of proteins with endogenous CAV1 and caveolae but is an inherent property of this polypeptide. This suggests that CAV1 may affect fundamental membrane properties, as investigated in the following section.

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Figure 5. Scaffolding domain of CAV1 is crucial for inhibition of CLIC endocytosis.

CAV1−/− MEFs were transiently transfected with (A) CAV1G83S-mCherry, (B) RFP-CAV3 and Cav3G55S-HA (C) CAV1Δ80-100-HA and (D) CAV1-SD-GFP (GFP-tagged minimal caveolin scaffolding domain) respectively. (A–D) Internalization assay was performed with anti-CD44 mAb and Tfn-647 for 2 min at 37°C. Cells were acid washed prior to fixation and internalized anti-CD44 mAb was labeled with AF-555 secondary antibody. 40–50 cells from each transfection were quantified for normalized fluorescence intensity of endocytic markers. In (A–D) data represent mean ± SEM of three independent experiments. **p = 0.002,***p = 0.0007, ****p<0.0001 (two-tailed t-test).

https://doi.org/10.1371/journal.pbio.1001832.g005

Caveolin-1 Decreases the Mobility of Lipid Raft Markers at the Plasma Membrane

CAV1 has been shown to interfere with the mobility of membrane microdomain-associated proteins, which further blocks the integrin-mediated internalization of bacteria [49]. We investigated whether similar effects could underlie the influence of expressed CAV1 and cavins on the CLIC/GEEC pathway.

We first analyzed, using FRAP, the degree of mobility of a lipid raft-associated CLIC/GEEC cargo protein, GPI-YFP, in the presence and absence of CAV1. WT and CAV1−/− MEFs were transiently transfected with GPI-YFP or co-transfected with GPI-YFP and CAV1-mCherry. To analyze the mobility of GPI-YFP, a small ROI was bleached at the PM, half-life times of fluorescence recovery were recorded and the resulting diffusion coefficients were calculated. When compared with WT MEFs or CAV1−/− MEFs expressing CAV1, a significant increase was observed in the mobility of GPI-YFP in CAV1−/− MEFs (diffusion coefficient for WT: 0.48±0.03; CAV1−/−+CAV1: 0.50±0.02; CAV1−/− 0.67±0.03; mean ± SEM, n>20 cells, p<0.001) (Figure 6A). We also analyzed the mobility of a model GPI-anchored protein in CAV1−/− MEFs co-expressing GPI-YFP and the CSD mutant CAV1G83S. As expected, the mutant failed to decrease the mobility of GPI-YFP (diffusion coefficient 0.78±0.06) (Figure 6A), suggesting that localization of CAV1 at the PM with an intact scaffolding domain is required to restrict the mobility of the tested microdomain-associated proteins. The very short half-life times (around 8.5 seconds) observed in the above analysis make it unlikely that the changes are due to a defect in trafficking of GPI-anchored proteins, as exocytosis and endocytosis occur at longer time scales.

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Figure 6. CAV1 alters general membrane characteristics.

(A) WT and CAV1−/− MEFs were transiently transfected with GP1-YFP; CAV1−/− MEFs were also co-transfected with GPI-YFP plus CAV1-mCherry or CAV1G83S-mCherry. FRAP analysis for GPI-YFP diffusion was performed, and FRAP curves and diffusion coefficients (D) calculated from FRAP measurements in at least 30 cells per condition are shown. (B) CAV1−/− MEFs were transiently transfected with PA-CD44 and co-transfected with PA-CD44 and CAV1 respectively. Fluorescence decay curves and diffusion coefficients (D) for PA-CD44, calculated from photo-activation measurements in at least 20 cells per condition, are shown. (C) CAV1−/− MEFs were either treated with 0.05% Tween20 at 37°C prior to performing internalization assay with anti-CD44 mAb and Tfn-647 or incubated with endocytic markers at 41°C. Cells were acid washed prior to fixation and internalized anti-CD44 mAb was labeled with AF-555 secondary antibody. 40–50 cells from each treatment were quantified for normalized fluorescence intensity of endocytic markers. (D,E) CAV1−/− MEFs transiently expressing CAV1-YFP and Cavin-1-GFP respectively, were either treated with Tween20 or incubated with endocytic markers at 41°C. Uptake and quantification of internalized markers was performed as mentioned in (C). (F) In CAV1−/− MEFs transiently expressing CAV1-YFP, internalization assay were performed with Dex-488 for 5 min either at 41°C or at 37°C. 40–50 cells from each condition were quantified for fluorescence intensity of the internalized Dex-488. (G) Representative images for filipin labeling performed for CAV1-YFP-expressing treated and untreated CAV1−/− MEFs and a bar graph representing quantification for normalized fluorescence intensity of filipin are shown. In (A) and (B) data represent average mean ± SEM of pooled data from three independent experiments. Diffusion coefficients were obtained by non-linear regression of recovery and fluorescence decay curves, as described in the Materials and Methods section. In (C–F) data represent mean ± SEM of three independent experiments. *p<0.01, **p = 0.007, ***p<0.005, ****p<0.0001 (two-tailed t-test). Scale bar: 10 µm.

https://doi.org/10.1371/journal.pbio.1001832.g006

We next tested the effect of CAV1 on the mobility of the CLIC-specific cargo protein CD44. To characterize CD44 membrane diffusion ability, we made use of an expression construct generated by fusing a photo-activatable variant of green fluorescent protein (mRFP-PAGFP) to CD44 (mRFP-PAGFP-CD44, termed PA-CD44). First, we tested the internalization route of the PA-CD44 construct. PA-CD44 was photo-activated at a specific region of the PM of PA-CD44-expressing COS-7 cells. Internalized PA-CD44 showed significant colocalization with Dex-647-labeled endocytic vesicles in the photo-activated region consistent with endocytosis via the CLIC pathway (Figure S7, Movie S2). For diffusion studies, PA-CD44 was photo-activated in a small ROI at the PM and decay of fluorescence over time was measured in CAV1−/− MEFs and CAV1−/− MEFs expressing CAV1, respectively. In the presence of CAV1, the CD44 diffusion rate was significantly decreased in comparison with CAV1−/− MEFs and was on a similar time scale to that observed in the FRAP experiments (Figure 6B).

As the effects of CAV1 on CD44 and GPI-YFP may be mediated through its impact on physical properties of the PM, we then investigated whether stimulation of membrane fluidity by chemical or physical means would have any effect on internalization of cargo proteins of the CLIC/GEEC pathway using two independent treatments [49],[50]. As a chemical stimulus, we treated CAV1−/− MEFs with 0.05% Tween20 for 15 min at 37°C. After treatment, cells were incubated with anti-CD44 mAb and Tfn-647 for 2 min at 37°C. Secondly, we used high temperature, which can also alter the membrane mobility of proteins. CAV1−/− MEFs were incubated with anti-CD44 mAb and Tfn-647 for 2 min at 41°C. Interestingly, both physical and chemical stimuli caused a significant increase in CD44 mAb uptake while Tfn-647 uptake was unaffected, compared with untreated cells (Figure 6C). We further analyzed whether alterations in membrane fluidity or mobility of microdomain-associated proteins could overcome the inhibitory effect of CAV1 and Cavin-1 on their internalization by the CLIC/GEEC pathway. CAV1−/− MEFs were transiently transfected with CAV1-YFP and Cavin-1-GFP respectively. Cells were either treated with Tween20 prior to uptake or incubated with endocytic markers at 41°C, as described above. Both treatments significantly increased CD44 mAb uptake, overcoming the inhibitory effect of CAV1 (Figure 6D). In contrast the treatments did not affect endocytosis in Cavin-1 expressing cells (Figure 6E). Tfn-647 internalization was also unaffected in both treated and untreated cells (Figure 6D,E). We also tested the effect of an increase in membrane fluidity on fluid phase endocytosis. CAV1−/− MEFs were transiently transfected with CAV1-mCherry, and Dex-488 uptake (5 min) was performed at 41°C. At 41°C, Dex-488 uptake was unaffected by CAV1 expression, suggesting that higher temperature rescues the inhibition of both a specific CLIC marker and a fluid phase marker (Figure 6F). Taken together, these results suggest that CAV1 expression can modulate the mobility of membrane microdomain-associated proteins. Treatments that increase the mobility of these proteins can at least partially restore endocytosis through the CLIC pathway.

The physical and chemical stimuli described above alter mobility or fluidity of microdomain-associated proteins, an effect that might indicate lipid changes in the PM. As cholesterol is crucial for the function of the CLIC pathway [30], we investigated the effect of these treatments on cellular cholesterol distribution using the polyene antibiotic, filipin. CAV1−/− MEFs were transiently transfected with CAV1-YFP, treated with Tween20 or incubated with endocytic markers at 41°C, and labeled with filipin. In CAV1-expressing and treated cells, higher filipin labeling was observed (135.7±4.8% for Tween20 and 132.7±4.1% for 41°C; mean ± SEM) in comparison with treated, untransfected CAV1−/− MEFs (100±1.6%; mean ± SEM) (Figure 6G) or untreated CAV1-expressing cells. This suggests that CAV1 expression affects membrane cholesterol distribution/availability, which inhibits internalization via the CLIC/GEEC pathway. The physical and chemical stimuli restore cholesterol distribution allowing up-regulation of the CLIC/GEEC pathway. This effect is more complex than simply the availability of free cholesterol in the bulk membrane because cholesterol addition in the form of a cholesterol–cyclodextrin complex did not rescue the CAV1-mediated inhibition (unpublished data).

Cavin-1 Influences Cholesterol Distribution, Cdc42 Activity, and Endosomal Sorting

Unlike CAV1, physical and chemical stimuli could not rescue the internalization of CD44 mAb in Cavin-1-expressing cells (Figure 6E), suggesting Cavin-1 might not act by regulating the mobility of microdomain-associated surface proteins in these cells. Cavins have also been reported to respond to cholesterol levels and alter cholesterol cellular distribution [51],[52]. Thus, we next tested whether expression of Cavin-1 in cells lacking CAV1 could affect cellular cholesterol distribution by labeling Cavin-1-GFP-expressing CAV1−/− MEFs with filipin to analyze the distribution of free cholesterol. In Cavin-1-expressing cells we observed a significant increase (138±8.4%; mean ± SEM) in filipin labeling in comparison with untransfected cells (100±6.5%; mean ± SEM) (Figure 7A). These results suggest that Cavin-1 expression can alter the distribution of filipin staining, indicating a change in distribution of cholesterol in cells.

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Figure 7. Cavin-1 regulates cholesterol distribution and Cdc42 activity.

(A) Filipin labeling was performed for Cavin-1-GFP-expressing CAV1−/− MEFs, and for quantification average fluorescence intensity measured from 40–50 transfected cells was normalized to untransfected cells. (B) Whole cell lysates from CAV1−/− MEFs expressing CAV1-YFP and Cavin-1-GFP respectively were immunoblotted for Cdc42 and glyceraldehyde 3-phosphate dehydrogenase (GAPDH) was used as a loading control. The bar graph represents quantitation of protein levels, calculated by measuring band intensities by densitometry. (C) Pearson coefficient analysis determined at the plasma membrane for CAV1/Cavin-1 (positive control), Cavin-1/Cdc42, and randomized Cavin-1/Cdc42. (D) CAV1−/− MEFs were co-transfected with CRIB-YPet and Cavin-1-mCherry and quantitative colocalization analysis was performed at the PM ruffles. For 30 cells, line scans (10 pixels' length) were performed at random positions on PM ruffles and cytosol. The resultant fluorescent intensities profiles were plotted against each other and subjected to linear regression analysis. (E) CAV1−/− MEFs were transiently transfected with Cdc42-CyPet alone, with Cdc42-CyPet and CRIB-YPet simultaneously, or with Cdc42-CyPet, CRIB-YPet, and Cavin-1-Flag. Lifetime of CyPet was analyzed by FLIM-FRET and a representative graph showing lifetime of CyPet in 25 cells, mean ± SEM, is shown. (F) In Cavin-1−/− and CAV1−/− MEFs internalization assay was performed with anti-CD44 mAb and Tfn-647 for 2 and 10 min at 37°C. Endogenous CAV1 and Cavin-1 was labeled with respective primary antibodies followed by AF-488 secondary antibody labeling and for internalized anti-CD44 mAb labeling AF-555 secondary antibody was used. In (A,B,D) data represent mean ± SEM of three independent experiments. In (C) data represent average mean ± SEM of pooled data from three independent experiments; n = 40 cells. **p<0.01,***p<0.001 ****p<0.0001 (two-tailed t-test). Scale bar: 10 µm.

https://doi.org/10.1371/journal.pbio.1001832.g007

To examine the possible implications of such alteration in lipid distribution, we focused on Cdc42, whose localization and activation has been shown to depend on specific lipid organization at the PM [53]. CAV1−/− MEFs were transiently transfected with Cavin-1-GFP and endogenous Cdc42 protein levels were analyzed by Western blotting. No effect was observed on total Cdc42 protein levels in Cavin-1-expressing cells (Figure 7B, Figure S8). We next characterized the correlation between Cavin-1 and Cdc42 at the PM. CAV1−/− MEFs expressing Cavin-1-GFP were labeled for endogenous Cdc42 and subjected to Pearson coefficient analysis to calculate the correlation between the two proteins. This revealed a significant spatial correlation, compared with randomized values, between Cdc42 and Cavin-1 at the cell surface, although less than that between CAV1 and Cavin-1 (Cavin-1–Cdc42: 0.21±0.02, randomized: −0.01±0.01, CAV1–Cavin-1 PM: 0.78±0.02; mean ± SEM, n = 40, p<0.0001) (Figure 7C). This suggested that Cdc42 and Cavin-1 might colocalize within regions of the cell surface. Indeed, live-cell imaging of CAV1−/− MEFs co-transfected with Cdc42-GFP and Cavin-1-mCherry revealed that these two proteins co-accumulated in PM ruffles (Figure S9A). Further live-cell imaging demonstrated that GPI-YFP also co-accumulated with Cavin-1-mCherry in PM ruffles, suggesting that these are sites for the possible association of Cavin-1 with CLIC components (Figure S9B).

We next tested whether Cavin-1 also affected Cdc42 activity, using a fluorescently tagged CRIB domain (Cdc42/Rac-interacting binding domain from N-WASP). This construct (CRIB-YPet) binds to GTP-loaded Cdc42 and thus can act as a location biosensor for active Cdc42. CAV1−/− MEFs were either transfected with CRIB-YPet alone or co-transfected with both CRIB-Ypet and Cavin-1-mCherry. Quantitative line scan analysis of fluorescence intensity showed a high degree of colocalization between CRIB and Cavin-1 at the PM ruffles in comparison with cytosol (Pearson coefficient, PM: 0.82±0.04; cytosol: 0.35±0.01; mean ± SEM, p<0.001) (Figure 7D). This supported the notion that Cavin-1 colocalized with active Cdc42 in ruffles.

We then tested whether Cdc42 could be activated by Cavin-1 expression. We used the FRET pair of Cdc42-CyPet and CRIB-YPet, whose association on GTP-loading of Cdc42 leads to a decrease in the lifetime of CyPet that can be measured by FLIM-FRET. CAV1−/− MEFs were co-transfected with Cdc42-CyPet and CRIB-YPet either in the presence or absence of Cavin-1. The lifetime of CyPet was significantly reduced, consistent with energy transfer in Cavin-1-expressing cells (1.5±0.02 nanoseconds), compared with cells lacking Cavin-1 (1.64±0.01 nanoseconds; mean ± SEM) (Figure 7E). Therefore Cavin-1 expression promotes activation of Cdc42 in CAV1-deficient cells. All together these results suggest that Cdc42 is selectively activated in PM ruffles when non-caveolar Cavin-1 is expressed in cells.

We also noted additional cellular effects upon the loss of Cavin-1. In WT MEFs, neither CAV1 nor Cavin-1 colocalized significantly with either anti-CD44 mAb or Tfn-647 after 2 or 10 min of uptake (Figure 1A). However, in Cavin-1−/− MEFs colocalization was observed between CAV1 and the internalized CD44 mAb but not with Tfn-647 after 2 min of uptake (Figure 7F). We further confirmed this association of CAV1 with CLICs by assessing its localization following inhibition of either CLICs or CME. Upon inhibition of CME by the small molecule dynamin inhibitor dyngo4a, CAV1 still colocalized with the CD44 mAb, whereas this colocalization was lost when the CLIC/GEEC pathway was inhibited by 7-KC (Figure S10). This suggests that noncaveolar caveolin is preferentially recruited into the CLIC/GEEC pathway in cells lacking Cavin-1.

Loss of Cavin-1 also affects trafficking through the CLIC/GEEC pathway. After 10 min of uptake in the Cavin-1−/− MEFs, internalized CD44 mAb and Tfn-647 were seen in the same endocytic vesicles (Figure 7F); this was not observed in WT or in CAV1−/− MEFs. These results provide evidence for a novel role for caveolar proteins in regulating not only the magnitude but also specific features of the endosomal system. In this respect we noted a significant increase in the expression levels of Tfn receptor in Cavin-1−/− MEFs (Figure S11).

Physiological Regulation of CLIC Endocytosis by Caveolins and Cavins

The inhibitory roles of caveolae, caveolins, and cavins suggest an important role for caveolae in regulating the CLIC/GEEC pathway. We therefore examined the physiological consequences of this inhibition. We first examined whether caveolae could play a role in spatial organization of the CLIC/GEEC pathway in migrating cells. A scratch-wound assay was applied to a confluent monolayer of CAV1−/− MEFs co-expressing CAV1 and Cavin-1. Fluorescently tagged CAV1 and Cavin-1 showed complete colocalization at the rear of the migrating cells (Figure 8A), consistent with previous studies showing caveolae enriched in this domain [38],[54]. CLIC endocytosis as detected by CD44 mAb uptake was dramatically reduced by the co-expression of the two caveolar proteins but the few carriers observed were invariably in areas lacking caveolae (Figure 8A). Concomitantly, in confluent monolayers of CAV1-expressing CAV1−/− MEFs, we also checked the cellular distribution of Cdc42, a pivotal regulator of polarity and the CLIC/GEEC pathway. In CAV1-YFP-expressing cells Cdc42 expression, detected by a Cdc42-specific antibody, was observed to be excluded from CAV1-positive areas. Quantification of fluorescence intensity at the cell surface, by line scan analysis, showed a significant decrease in Cdc42 expression in regions of PM expressing high levels of CAV1 compared with low- or nonCAV1-expressing regions (CAV1 high expression region: 14±0.32; low/no expression regions: 26±0.60; mean ± SEM, p<0.0001), within the same cell (Figure 8B). However, overall Cdc42 protein levels were unaltered in CAV1-YFP-expressing cells, as observed by Western blotting (Figure 7B, Figure S8).

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Figure 8. Physiological consequences of caveolin/cavin loss.

(A) Confluent monolayer of CAV1−/− MEFs co-expressing CAV1-YFP and Cavin-1-mCherry was subjected to a scratch-wound assay. Internalization assay was performed with anti-CD44 mAb for 2 min at 37°C. Cells were acid washed before fixation and internalized anti-CD44 mAb was labeled with AF-647 secondary antibody. Arrow indicates direction of wound. (B) CAV1−/− MEFs expressing CAV1-YFP were labeled for Cdc42 with specific primary antibody followed by AF-555 secondary antibody labeling. For quantitative analysis of Cdc42 expression, line scans were performed in 30–40 CAV1-YFP-expressing cells at random positions on the PM. The plot profile represents average fluorescence (± SEM) of Cdc42 labeling in mentioned areas of PM from all cells and the bar graph represents amplitude together with error obtained from nonlinear Gaussian analysis. (C) Pearson coefficient analysis of the image data set from (B). (D) CAV1−/− MEFs were co-transfected with CRIB-Ypet and CAV1-mCherry and quantitative colocalization analysis was performed at the random regions of PM. For 20–30 co-transfected cells, line scans and nonlinear Gaussian analysis were performed as mentioned in (B) at random positions on the PM. (E) WT, CAV1−/−, and Cavin-1−/− MEF monolayers were subjected to a scratch-wound assay and the extent of migration of respective cells into the wound area was measured at 0, 4, 8, and 12 h time points. Percentage of wound closure is plotted. (F) At time points 0, 4, 8, and 12 h a scratch-wound assay, followed by internalization assay, was applied to WT, CAV1−/−, and Cavin-1−/− MEFs as mentioned in (A). For labeling endogenous CAV1 and Cavin-1 respective primary antibodies followed by AF-488 secondary antibody were used. The plot profile represents average pixel intensity across the leading to trailing edge for CD44, CAV1, and Tfn-647. Arrows indicate CD44 labeling. In (C) data represents mean ± SEM of data pooled from three independent experiments; n = 40 cells. In (B, D, E) data represent mean ± SEM of three independent experiments. *p<0.05, **p<0.01, ***p = 0.001, ****p<0.0001 (two-tailed t-test). Scale bar: 10 µm.

https://doi.org/10.1371/journal.pbio.1001832.g008

To further characterize the correlation between Cdc42 and CAV1 at the cell surface, we performed the Pearson correlation coefficient analysis on the same set of images that were used for line scan analysis. This revealed significantly less correlation between CAV1 and Cdc42 at the PM in comparison with CAV1 and Cavin-1 at the PM (Pearson coefficient, CAV1–Cdc42: 0.11±0.01, randomized: −0.002±0.001, CAV1–Cavin-1 PM: 0.78±0.02; mean ± SEM, n = 40, p<0.0001) (Figure 8C). As expected from the above results, the activity of Cdc42, monitored by recruitment of the CRIB domain, was also negatively affected by CAV1 expression. Quantitative line scan analysis showed significantly fewer CRIB protein in regions of the PM expressing high levels of CAV1 in comparison with lower-expressing regions (PM region enriched in CAV1: 45±1.9; PM regions with low/no CAV1 expression: 57±2.4; mean ± SEM, p<0.001) within the same cell (Figure 8D). This suggests that CAV1 expression can result in differential distribution of CLIC components; in addition, caveolae can locally inhibit CLIC endocytosis and help polarize the pathway to the leading edge.

To test this hypothesis, we next examined the effect of loss of caveolar proteins on polarization of CLICs to the leading edge of 2D migrating fibroblasts, a key feature of the CLIC/GEEC pathway [29]. First we analyzed whether lack of caveolar proteins can alter cell migration by using a scratch-wound assay applied to confluent monolayers of WT, CAV1−/−, and Cavin-1−/− MEFs. As shown in a previous study [55], we observed a significant decrease in ability of CAV1−/− MEFs to close the wound compared with WT MEFs (after wound closure at 12 h, WT: 84±2.1%; CAV1−/−: 54±4.1%; mean ± SEM, p<0.0001) (Figure 8E). Similarly, Cavin-1−/− MEFs also showed less efficient wound closure compared with WT MEFs and showed defects in cell migration (after wound closure at 12 h, Cavin-1−/−: 60±2.5%; mean ± SEM, p<0.001) (Figure 8E). To characterize the polarization of CLICs in migrating cells, confluent monolayers of WT, CAV1−/−, and Cavin-1−/− MEFs were scratch-wounded and uptake of CD44 mAb and Tfn-647 was compared with the localization of CAV1 or Cavin-1 as cells migrated into wound. In both CAV1−/− and Cavin-1−/− MEFs, CD44 mAb internalization occurred at both the leading and trailing edge, indicating the CLIC/GEEC endocytosis was no longer polarized in these cells (Figure 8F).

Loss of caveolins and cavins has profound effects on specific tissues in vivo, including skeletal and striated muscle (reviewed by [56]). To examine whether the loss of caveolar components also affected endocytic activity in a differentiated tissue relevant to disease, we isolated mature adult muscle fibers from the flexor digitorum brevis muscle of WT and Cavin-1−/− mice. We assessed endocytic activity by adding either anti-CD44 mAb to the isolated fibers and visualizing uptake by IF, or by using HRP as a fluid phase marker for EM analysis. No significant difference was seen in CD44 surface labeling of the fibers, suggesting endogenous levels and surface accessibility of CD44 are not altered (not shown). CD44 mAb internalization was dramatically increased in fibers isolated from Cavin-1−/− mice (23.4±2.9, compared with WT: 2.2±0.3; mean ± SEM) (Figure 9A). Similarly, isolated fibers incubated with HRP as a fluid phase marker showed a highly significant 4.5-fold increase in the volume of HRP-labeled structures, as compared with WT fibers (Figure 9B–D), indicative of greatly increased fluid phase endocytosis. Taken together, these data show that the CLIC/GEEC pathway is fundamentally altered by the loss of Cavin-1 demonstrating a dramatic in vivo consequence of loss of caveolar components.

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Figure 9. Loss of Cavin-1 up-regulates endocytic activity in muscle fibers.

(A) Muscle fibers, isolated from Cavin-1−/− and WT mice, were incubated with anti-CD44 mAb for 3 min at 37°C. Fibers were acid washed on ice and internalized anti-CD44 mAb was labeled with AF-555 secondary antibody. 8–10 muscle fibers from three independent experiments were quantified for total fluorescence intensity of internalized CD44 mAb. (B,C) HRP uptake was performed in WT (B) and Cavin-1−/− (C) muscle fibers followed by DAB reaction. Arrows indicate DAB-positive intracellular structures. (D) Quantitation of DAB-labeled structures, as described in the Materials and Methods section, is shown. In (A, D) data represent mean ± SEM of three independent experiments. ***p<0.001, ****p<0.0001 (two-tailed t-test). Scale bars: (A) 10 µm, (B) 1 µm, (C) 500 nm.

https://doi.org/10.1371/journal.pbio.1001832.g009

Ethics Statement

All the animal experiments were conducted in accordance with the guidelines of the ethics committee at The University of Queensland.

Antibodies and Reagents

Mouse anti-CD44 (clone 5035-41.1D, Novus Biologicals), rabbit anti-CAV1 (BD Biosciences), rabbit anti-Caveolin2 (Sigma Aldrich), rabbit anti-Cavin-1 and Cavin-4 antibody were raised as described previously [16], rabbit anti-Cavin-3 (ProteinTech group), mouse anti-Cavin-2 (Sigma Aldrich), rabbit anti-HA (Sigma Aldrich), mouse anti-Cdc42 (Becton Dickinson), mouse anti-GFP (Roche), mouse anti-transferrin receptor antibody (Zymed), Alexa Fluor conjugated dextran (Life Technologies), anti-rabbit and anti-mouse Alexa Fluor antibodies (Invitrogen), Alexa Fluor conjugated transferrin (Invitrogen), Alexa Fluor conjugated phalloidin (Invitrogen), Filipin III (Sigma Aldrich), Dynasore (Sigma Aldrich), 7-Ketocholesterol (Sigma Aldrich), Protease Inhibitor Cocktail Set III (Merck Millipore), PhosSTOP Phosphatase Inhibitor Cocktail (Roche), Dyngo-4a (Sigma). Stealth RNAi siRNA duplex oligonucleotides targeted against mouse Cavin-1 (5′CCGCUGUCUACAAGGUGCCGCCUUU3′;5′AAAGGCGGCACCUUGUAGACAGCGG3′) and Cavin-3 (5′CCGGAGCUCUGAAGGCCCAUCAGAA3′; 5′UUCUGAUGGGCCUUCAGAGCUCCGG3′) (Invitrogen).

Cell Culture

WT, immortalized CAV1-null (CAV1−/−), Cavin-1 null (Cavin-1−/−) mouse embryonic fibroblasts (MEFs), 3T3-L1 and COS-7 cells were maintained in Dubelcco's modified Eagle's medium (DMEM; Gibco) supplemented with 10% fetal bovine serum (FBS; Cambrix) and 2 mM L-Glutamine at 37°C and 5% CO₂. For transfection, cells were either electroporated (BIORAD) or Lipofectamine 2000 (Invitrogen) was used, as per the manufacturer's instructions, and by this method 30–40% transfection efficiency was achieved. For siRNA transfection, Cavin-1 and Cavin-3-specific oligonucleotides were transfected at a final concentration of 600 pM using Lipofectamine 2000.

Internalization Assay, Cell Migration, and Immunocytochemistry

Cells grown on 12 mm coverslips were placed on 50 µl pre-warmed drops containing 10 µg/ml anti-CD44 antibody/3 mg/ml Dex-488/10 µg/ml anti-GFP antibody+10 µg/ml Tfn-647 diluted in culture media and incubated at 37°C in a 5% CO₂ incubator for time indicated. To remove any surface bound markers (CD44 mAb, GFP mAb, or Tfn-647), 2×30-second acid stripping was performed with 0.5 M glycine (pH 2.2), before fixation with 4% paraformaldehyde. The cell migration (scratch-wound) assay and immunocytochemistry were performed as previously described [29]. For cholesterol staining, cells were labeled with filipin, 50 µg/ml, for 2 h in the dark. For Cdc42 staining, cells were fixed with 10% TCA and quenched using 3×5-min washes of 30 mM glycine on ice. Permeabilized cells were blocked with 3% bovine serum albumin (BSA) and incubated in primary antibody overnight at 4°C followed by Alexa Fluor conjugated secondary antibody incubation for 40 min at room temperature.

Electron Microscopy and Quantification of CLICs Carrier

In transfected CAV1−/− MEFs, HRP uptake (10 µg/ml) was performed at 37°C for 2 min, and after brief washing with DMEM containing 1% BSA, diaminobenzidine (DAB) (10 mg/ml) reaction was performed on the live cell as previously described in [29]. Fixation, embedding, and sectioning were performed as follows. MEFs were fixed with 2.5% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.4). Cells were post-fixed with 1% osmium tetroxide for 1 h at room temperature and serially dehydrated with ethanol. Cells were embedded in increasing ratios of LX-112 resin∶ethanol to 100% resin, and polymerized overnight at 60°C. Ultrathin (60 nm) sections were cut on a Leica UC6 microtome and imaged on a JEOL1011 electron microscope at 80 kV. Quantifications were performed as follows: the perimeters of approximately 16 cells (per experimental condition) were imaged and the number of CLICs/GEEC carriers from each cell was quantified and averaged across all 16 cells. The average number of CLICs/GEEC carriers per cell was generated from two separate repeats of the same experimental conditions.

Image Acquisition (for General and Live-Cell Imaging)

Cells were grown either on 12 mm coverslips or on 35 mm glass bottom dishes (Mat-Tek Corporation) and fluorescence micrographs were captured for random fields containing transfected cells on a confocal laser-scanning microscope (Zeiss 710 META; 510 META; Carl Zeiss Inc.). Images were captured with a 63× plan Apochromat 1.4 NA Oil objective (Zeiss, Jena, Germany), using a 488 nm laser line for excitation and a 505–530 nm band pass emission filter to capture GFP and Alexa Fluor 488 fluorescence, a 561 nm laser line for excitation, and a 580–620 nm band pass emission filter to capture RFP/mCherry/Alexa Fluor 555 fluorescence, and a 633 nm laser line for excitation and a 650 long pass emission filter for emission to capture Alexa Fluor 647 fluorescence. For live cell co-imaging of Cherry and YFP, YFP fluorescence was captured using the 488 nm laser line for excitation and a 505–530 nm band pass for emission. For live cell imaging a region of interest was chosen and 2× digital zoom was applied. Optical path and emission filters were applied as appropriate for each fluorophore. Images were processed with Adobe Photoshop CS3 (Adobe, San Jose, CA) and fluorescent intensity of fluorophores on images was measured using ImageJ (National Institutes of Health, Bethesda, MD). Where indicated, correlation between fluorescence intensities between two fluorophores in different subcellular location was determined using Pearson coefficient analysis. For this, binary masks (values 0 and 1) were created to isolate pixels belonging to different regions of interest (such as the plasma membrane and/or cytosol) by multiplying (pixel by pixel) the mask image by the raw images. This method was chosen in preference to the Pearson analysis over the entire cell as it provides a better estimation of whether or not a high correlation exists between different fluorophores at a specific subcellular location, for example the plasma membrane. Then, each individual image (two channels) was used to calculate the Pearson correlation coefficient between the two fluorophores in each specific subcellular location, and the data represent an average Pearson correlation coefficient determined for at least 40 cells from at least three independent experiments.

FRAP Experiments

To assess GPI-YFP dynamics, cells were seeded on 35 mm glass bottom dishes and transfected with GPI-YFP construct [65]. Images were captured on an inverted confocal microscope using a 63× plan Apo 1.4 NA, Oil objective with 4× digital zoom, with a resolution of 0.15 µm/pixel. A circular region of interest (ROI, 4.7 µm radius) was bleached to ∼70% using the 488 nm argon laser line and 405 nm laser line at 100% transmission. Time-lapse images of the same region and a reference region of identical size were acquired before (20 frames, 5 seconds) and after (300 frames, 90 seconds) photobleaching with an interval of ∼250 milliseconds per frame using the 488 nm laser line at 2% transmission and emission detected between 500 and 530 nm. Half-life time and diffusion rate were calculated as described previously [49],[66]; also see Text S1 for details.

CAV1 dynamics were determined in Cavin-1−/− cells expressing CAV1-YFP alone or co-transfected with CAV1-YFP and Cavin-1-mCherry. Images were acquired as described above and half-times were calculated as described previously for CAV1-GFP in HeLa cells [67],[68]. The lateral diffusion nature of CAV1-YFP recovery curves at the plasma membrane was determined by performing FRAP experiments at variable ROI areas (9.6, 14.4, and 16.6 µm²) to analyze the dependency of half times with area size. Linear regression was performed and the diffusion coefficient for simple lateral diffusion was extracted from the slope as described in Text S1.

Photoactivation Experiments

To achieve spatially defined photoactivation of PAGFP, a Ti:sapphire two-photon laser (1600–1800 mW Chameleon Ultra, Coherent Scientific) tuned at 775 nm was used [69]. For lateral diffusion studies of CD44, the plasma membrane region of cells was identified with the mRFP signal of CD44-mRFP-PA-GFP. PAGFP fluorescence was activated using a constant circular region of interest (ROI, 1.98 µm radius) by a single scan with infrared laser irradiation (30% transmission), and time-lapse images of photo-activated GFP at the same region were captured every ∼250 millisecond using a 60× plan Apo 1.4 NA, Oil objective at 4× digital zoom and appropriate filter sets to capture GFP fluorescence. For each time point, the average fluorescence intensity was calculated and normalized to the value at the first frame after photo-activation. Decay curves were fitted to a double exponential decay curve and a global half-time of fluorescence decay was obtained numerically as described for FRAP experiments.

FLIM-FRET

CAV1−/− MEFs expressing FRET pair (Cdc42-CyPet and CRIB-YPet) either in presence or absence of FLAG-tagged Cavin-1 were subjected to FLIM microscopy as described previously [18].

Dynamin Inhibition and 7-Ketocholesterol Treatment

Cells were serum starved for 3 h in serum-free DMEM before incubation with 60 µM dynasore/Dyngo-4a. Internalization assay was performed in the presence of 40 µM dynasore for the desired amount of time. Cells were treated with 30 µM 7-ketocholesterol (7-KC) for 30 min at 37°C, followed by the internalization assay.

Western Blotting

Whole cell lysates were subjected to SDS-PAGE and further to Western blotting. Membranes were probed with primary antibody at the desired concentration for 1 h at room temperature or overnight at 4°C, followed by incubation with either secondary HRP-conjugated antibodies or infrared dye-labeled Odyssey secondary antibodies for 1 h at room temperature. For detection, either the Licor Odyssey infrared imaging system was used, as per the manufacturer's instructions (Licor Biotechnology), or the SuperSignal West Pico chemiluminescent substrate (Pierce) was captured on film (Kodak). Densitometric analysis of protein bands was performed either by ImageJ or by using Licor Odyssey analysis software. Also see Text S1 for details.

Muscle Fiber Isolation and Analysis of Endocytic Activity

Muscle fibers were isolated from WT and Cavin-1−/− adult mice using a method described previously [70], with modifications. For EM analysis of HRP uptake, isolated fibers cultured overnight on matrigel-coated plastic dishes were incubated in HRP (10 mg/ml) at 37°C for 5 min, washed briefly, then fixed in glutaraldehyde before DAB visualization of the HRP reaction product. Quantitation of the volume of HRP-labeled elements relative to the sampled cytoplasmic volume (volume density) was determined by point counting of peripheral areas of WT and Cavin-1−/− muscle fibers, as shown in previous studies [29]. For procedure details see Text S1.

Statistical Analyses

Statistical analyses were conducted using Microsoft Excel and Prism (GraphPad). Error bars represent either standard error of the mean (SEM) or standard deviation (SD) for at least three independent experiments, as indicated in figure legends. Statistical significance was determined either by two-tailed Student's t-test or by one-way ANOVA, as indicated in the figure legends.