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Originally published In Press as doi:10.1074/jbc.M702117200 on July 3, 2007

J. Biol. Chem., Vol. 282, Issue 36, 26542-26551, September 7, 2007
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The Tetraspan Protein EMP2 Regulates Expression of Caveolin-1*

Ashley Forbes{ddagger}1, Madhuri Wadehra§1, Sergei Mareninov{ddagger}, Shawn Morales§, Kaori Shimazaki{ddagger}, Lynn K. Gordon||, and Jonathan Braun{ddagger}2

From the {ddagger}Molecular Biology Institute, §Department of Pathology and Laboratory Medicine, David Geffen School of Medicine, and Department of Ophthalmology, UCLA, Los Angeles, California 90095 and the ||Department of Surgery, Greater Los Angeles Veterans Affairs Healthcare System, Los Angeles, California 90099

Received for publication, March 12, 2007 , and in revised form, June 8, 2007.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Caveolin-1 is the primary component of caveolae and functions in a variety of intracellular activities, including membrane trafficking and signal transduction. EMP2 (epithelial membrane protein 2) is a tetraspan protein recently identified as a novel regulator of caveolin-1 expression. In this study, we analyzed the mechanism of EMP2-mediated caveolin-1 regulation. In NIH 3T3 cells and in the human retinal pigment epithelium cell line (ARPE-19), EMP2 regulates caveolin-1 transcription and more substantially its protein levels. EMP2-mediated down-regulation of caveolin-1 does not affect caveolin-1 translational efficiency, phosphorylation, or proteasome-mediated degradation. Analysis of caveolin-1 protein half-life indicates the EMP2-mediated loss of caveolin-1 occurs rapidly. Protease inhibition and laser confocal microscopy associates this fate with specific intracellular compartmentalization, including early lysosomal delivery. These findings elucidate a new mechanism of caveolin-1 regulation and define an additional role for EMP2 as a key regulator of cell membrane composition.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Caveolae are flask-shaped plasma membrane invaginations thought to play a role in endocytosis, cholesterol homeostasis, intracellular signaling, and cell transformation. Caveolae are found in most cell types but are particularly abundant in terminally differentiated cells such as adipocytes, endothelial cells, type 1 pneumocytes, fibroblasts, and muscle cells (1, 2). The primary structural component of caveolae is caveolin-1, a 21-24-kDa integral membrane protein believed to act as a scaffolding protein within caveolae membranes (3). Exogenous expression of caveolin-1 is sufficient to induce de novo caveolae formation (4, 5). Caveolae contribute to many cellular functions, including cholesterol binding and transport (6-8), inhibition of signaling molecules (2, 9), and suppression of oncogenic transformation (10, 11).

Transcriptional regulation of caveolin-1 expression is mediated by a variety of mechanisms (2, 8, 10-13). For example, the mitogen-activated protein kinase pathway extracellular signal-regulated kinase (ERK) has been shown to control caveolin-1 expression at the transcriptional level in NIH 3T3 cells (10), whereas activation of Src family kinases transcriptionally decreases caveolin-1 expression (2). In addition, caveolin-1 transcription is up-regulated by sterol regulatory element-binding protein-1 in response to free cholesterol, which results in caveolae formation and increased cholesterol efflux (12, 13). Conversely, high density lipoprotein exposure reduces caveolin-1 expression and prevents the uptake of oxidized cholesterol (14). Cell growth also plays a role in caveolin-1 regulation, as rapidly dividing cells display decreased caveolin-1 expression levels, whereas confluent cells have dramatically up-regulated caveolin-1 expression (11). To date, little has been reported about post-transcriptional modes of caveolin-1 protein regulation.

Caveolin-2 is another member of the caveolin gene family (3). Caveolin-2 is normally coexpressed with caveolin-1 and is thought to act as an accessory molecule, aiding in the formation of caveolae by forming hetero-oligomeric complexes with caveolin-1 (3, 15). However, caveolin-2 alone is not sufficient to facilitate caveolae formation (3). Caveolin-2 expression is linked to expression of caveolin-1. Caveolin-1 is thought to stabilize caveolin-2 protein and thereby post-transcriptionally regulate its protein levels (15).

Epithelial membrane protein 2 (EMP2) is a tetraspan protein recently identified as a novel regulator of caveolin-1 expression (16). EMP2 is a member of the GAS3/PMP22 (growth arrest-specific-3/peripheral myelin protein-22) family of proteins. Biologically, EMP2 plays an important role in blastocyst implantation with the uterine endometrium, in part through its direct association and regulation of {alpha}vbeta3 integrin (17). EMP2 controls cell membrane expression of integrins, glycosylphosphatidylinositol-anchored, and class I major histocompatibility complex proteins (16-19). EMP2 resides at the plasma membrane within lipid raft domains, and this localization is distinct from caveolin-bearing membrane structures (16). Previously, we have reported that recombinant overexpression of EMP2 in NIH 3T3 cells dramatically reduced expression of caveolin-1 and -2. Conversely, loss of EMP2 expression resulted in increased caveolin protein production (16). These findings indicate an important role for EMP2 in maintaining plasma membrane structure and function.

Here we further analyzed the mechanism of EMP2-mediated caveolin reduction. We show that EMP2 modulates caveolin-1 expression at both the transcriptional and translational levels, in both murine and human cell lines. Mechanistic analysis reveals that EMP2-mediated reduction of caveolin-1 protein involves aberrant intracellular compartmentalization and rapid degradation shortly after completion of translation. These observations thus describe a novel mechanism regulating caveolae formation.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Lines—NIH 3T3 cells were cultured in Dulbecco's modified Eagle's medium (DMEM3; Invitrogen) supplemented with 10% fetal bovine serum (Omega Scientific, Tarzana, CA), 2 mM L-glutamine, and 1 mM sodium pyruvate (Invitrogen). NIH 3T3 cells stably expressing a FLAG-tagged murine EMP2 (3T3/EMP2) or a vector control (3T3/V) were cultured as described previously (20). Cells were grown at 37 °C in a humidified 5% CO2 atmosphere.

The human retinal pigmented epithelial cell line ARPE-19 (ATCC CRL-2302) was grown in-DMEM/F12 (Invitrogen) supplemented as above. To ectopically increase the expression of human EMP2, ARPE-19 (designated ARPE-19/EMP2) was stably transfected with pEGFP-hEMP2, a plasmid encoding a functional human EMP2-GFP fusion protein in the EcoRI site of the pEGFP-N3 expression vector (17). A vector control cell line was similarly prepared with pEGFP-N3, resulting in GFP expression alone (designated ARPE-19/V). ARPE-19 transfectants were produced using FuGENE 6 (Roche Applied Science), and stable clones were selected using 700 µg/ml geneticin (Invitrogen).

Northern Blot Analysis—Total RNA was isolated using RNA purification and RNA cleanup kits (Qiagen, Valencia, CA). RNA (10 µg) was subjected to agarose electrophoresis (1.6% agarose/formaldehyde), transferred to a nylon membrane (Amersham Biosciences) by capillary action, and cross-linked by UV irradiation (Stratalinker, Stratagene, San Diego). The following primers were used to generate cDNA probes: murine caveolin-1 CTACAAGCCCAACAACAAGG(C/A)GGAAGCTCTTGATGCACGGT; murine caveolin-2 ATGACGCCTACAGCCACCACA(G/G)CAAACAGGATACCCGCAATG. Probes were 32P-labeled using random primer synthesis kit (Stratagene) as described previously (20). Membranes were pre-hybridized with Rapid-Hyb buffer (Amersham Biosciences) for 1 h and then hybridized with labeled probe overnight at 65 °C. Blots were washed in low stringency buffer (65 °C, 2x sodium chloride/sodium citrate (SSC), and 0.1% SDS) and high stringency buffer (65 °C, 0.1x SSC, and 0.1% SDS), as needed, and exposed to x-ray film.

Antibodies—Anti-caveolin-1 antibodies were rabbit polyclonal anti-caveolin-1 antibody (N-20, Santa Cruz Biotechnology, Santa Cruz, CA), mouse monoclonal anti-caveolin-1-{alpha} and -beta antibody (clone 2297, BD Biosciences), and rabbit polyclonal anti-phospho-caveolin-1 (Tyr-14, Cell Signaling Technology, Beverly, MA). Mouse anti-caveolin-2 antibody (clone 65, BD Biosciences), mouse monoclonal anti-p53 antibody (2524, Cell Signaling Technology), rabbit polyclonal anti-beta-actin antibody (ab8227, Abcam Inc., Cambridge, MA), goat anti-rabbit IgG horseradish peroxidase conjugate (4050-05, Southern Biotech, Birmingham, AL), and sheep anti-mouse IgG horseradish peroxidase conjugate (NA931, Amersham Biosciences) were also used.

Additional antibodies utilized for confocal microscopy included anti-{gamma}-adaptin, anti-mouse CD107a (LAMP-1)-FITC, and anti-EEA-1 (all from BD Biosciences). Rhodamine-conjugated goat anti-rabbit IgG (H+L), FITC-conjugated anti-rabbit IgG (H+L), and Texas Red-conjugated goat anti-mouse IgG were obtained from Jackson ImmunoResearch, West Grove, PA.

Immunoblot Analysis—Cells were lysed in Laemmli buffer, separated by SDS-PAGE, and then transferred to a nitrocellulose membrane (Amersham Biosciences). For EMP2 expression, cell lysates were additionally treated with peptide:N-glycosidase (New England Biolabs) to remove N-linked glycans that interfere with the detection of EMP2 epitopes (20). Membranes were stained with Ponceau S (Sigma) to determine transfer efficiency. Membranes were blocked with 10% low fat milk or 5% bovine serum albumin in Tris-buffered saline containing 0.1% Tween 20 and probed with antibodies. Protein bands were visualized using a horseradish peroxidase-labeled secondary antibody, followed by chemiluminescence (ECL, Amersham Biosciences).

Subcellular Fractionation—3T3/V and 3T3/EMP2 cells (average cell number 3 x 106) were fractionated as described previously (21, 22). Briefly, cells were washed twice in a sucrose buffer (pH 7.4) containing 20 mM HEPES, 1 mM EDTA, 250 mM sucrose, and protease inhibitors (Complete Mini Tablets, Roche Applied Science). This sucrose buffer was used throughout all subsequent procedures that were carried out at 4 °C. Cells were resuspended in 1.0 ml of the same buffer, and homogenized three times with 10 strokes in a 2-ml Potter/Elvehjem Teflon pestle homogenizer. The original cell homogenate was transferred into a 1.5-ml tube and centrifuged at 16,000 x gmax for 15 min in an Eppendorf (model 5415R) centrifuge. The resulting pellet was washed in 0.5 ml of sucrose buffer and recentrifuged as above. The washing supernatant was combined with the supernatant of the initial pellet and was saved for preparation of the microsomal membrane fraction. The pellet was resuspended in 0.25 ml of sucrose buffer and over-layered onto a 1.12 M sucrose cushion containing 20 mM HEPES, 1 mM EDTA, and protease inhibitors and centrifuged at 101,000 x gmax for 70 min. The resulting pellet containing the mitochondria, nuclei, and cell debris was collected, washed once, and re-pelleted at 16,000 x gmax for 15 min and finally was resuspended in sucrose buffer and saved as nuclear fraction. The plasma membranes were collected at the sucrose interface, were washed once, re-centrifuged at 48,000 x gmax for 45 min, and finally resuspended in sucrose buffer and saved as plasma membrane fraction. The initial supernatant was centrifuged at 212,000 x gmax for 70 min yielding the microsomal fraction in the pellet. Equal volume equivalents of each fraction were separated by immunoblot analysis as described above.

Phosphorylation Studies—To induce phosphorylation, cells were stressed with 600 mM sucrose or 5 mM H2O2 for 10 min at 37 °C (23). Cells were then washed in PBS, harvested, and assayed by immunoblot. Sucrose and H2O2 were diluted in complete growth medium.

Polysome Analysis—Cell lines were used at about 80% confluency to analyze polysome distribution. Cells were washed with ice-cold PBS containing 100 µg/ml cycloheximide, removed using a cell scraper, and lysed in polysome lysis buffer (100 mM KCl, 5 mM MgCl2, 10 mM HEPES (pH 7.4), 0.5% Nonidet P-40, 100 µg/ml cycloheximide, and 5 mM dithiothreitol). Nuclei were pelleted by centrifugation at 10,000 x g at 4 °C for 10 min. The resulting supernatant was layered onto a 15-50% (w/v) linear sucrose gradient and centrifuged at 38,000 rpm for 2 h at 4 °C in a Sorvall TH-61 rotor (Asheville, NC). Gradients were analyzed by passing the contents through a density gradient fractionator (ISCO, Los Angeles, CA) to monitor continual A260, and then 1-ml fractions were collected. Total RNA from each fraction was isolated using TRIzol LS reagent (Invitrogen). Equal volumes from each fraction were used for Northern blot analysis, as described above.

Proteasome Inhibition—Proteasome inhibition studies were also performed 3T3/V and 3T3/EMP2 cell lines at about 80% confluency. Epoxomicin (Boston Biochem, Cambridge, MA) dissolved in Me2SO was added to a final concentration of 100 nM, 1 µM, or 10 µM, or cells were given a Me2SO control or left untreated. Cells were incubated for 6 h, harvested, lysed in lysis buffer (150 mM NaCl, 10 mM Tris (pH 7.5), 1 mM EGTA, 1 mM EDTA, 1% Triton X-100, 0.5% Nonidet P-40, 1 µM phenylmethylsulfonyl fluoride, 1 µM pepstatin, 10 µM leupeptin, 2 mM sodium vanadate, and 5 µg/ml aprotinin), assayed for protein concentration, and subjected to Western blot analysis.

Metabolic Labeling and Immunoprecipitation—Metabolic labeling was performed by starvation for 20 min in DMEM without methionine and cysteine (supplemented with 5% dialyzed fetal bovine serum, 2 mML-glutamine, 1 mM sodium pyruvate, and antibiotics) followed by incubation for 1 h with 240 µCi/ml [35S]methionine and -cysteine Pro-mix (Amersham Biosciences). The media were replaced by DMEM supplemented with 2 mM methionine, and cells were analyzed at various time points ranging from 0 to 24 h. Cells were harvested in PBS, and cell pellets were frozen in liquid nitrogen and stored at -80 °C. To extract and denature proteins, cell pellets were lysed in 1% Nonidet P-40 containing 2 mM phenylmethylsulfonyl fluoride, 10 µg/ml aprotinin, 2 µg/ml, pepstatin, 10 mM iodoacetamide, 0.1 mM EDTA, 0.1 mM EGTA, 10 mM HEPES, and 10 mM KCl. Lysates were sonicated and then centrifuged to remove cell debris. Supernatants were collected, and protein concentration in the supernatant was then measured using the Bradford Assay Reagent (Bio-Rad). Samples were precleared using 8 µg of agarose-conjugated normal rabbit IgG (Santa Cruz Biotechnology) by rotating for 1 h at 4 °C. Equal amounts of protein from each sample were then incubated overnight with 30 µg of anti-caveolin-1 rabbit polyclonal antibody and protein A-agarose beads (both from Santa Cruz Biotechnology). Antibody-bound protein was collected by centrifugation, washed three times in lysis buffer, resuspended in 2x Laemmli buffer, boiled for 3 min, and equal volumes were then separated by SDS-PAGE. Resolved gels were then fixed, treated with Amplify fluorographic reagent (Amersham Biosciences), dried under vacuum, and exposed using a phosphor screen (GE Healthcare). Analysis was performed using Typhoon 9410 (GE Healthcare) and ImageQuant 5.2 software.

Lysosome Inhibition—3T3/EMP2 or 3T3/V cells were plated at 0.6 x 105/well in 6-well plates and cultured for 2 days. Cells were then placed in serum-free DMEM with lysosome inhibitors pepstatin A and leupeptin (both at 1 or 5 µM; Calbiochem) alone or in combination as described previously (24). Cells were incubated for 4 h and then harvested for Western blot analysis. Bands were quantitated using Image J 1.36b software (25).

Laser Confocal Microscopy—1 x 105 NIH 3T3 cells or 3T3/EMP2 were plated on 18-mm diameter glass coverslips (Fisher) and incubated for 48 h. Cells were fixed in methanol and blocked with 1% normal goat serum for 1 h at room temperature. Samples were incubated with the primary antibodies, as listed in the figure legends, for 1 h at room temperature and visualized using a fluorescent secondary antibody. Confocal pictures were taken using a Zeiss LSM 510 confocal microscope at x600 magnification.

To measure caveolin-1 localization in different intracellular compartments, cells were processed for dual staining with antibodies for caveolin-1 and compartment markers ({gamma}-adaptin, EEA-1, or LAMP-1). To assess the lysosomal compartment, cells prior to fixation were incubated for 1 h in 0.5 µg/ml leupeptin and 0.5 µg/ml pepstatin (to reduce caveolin-1 degradation). The colocalization coefficient was quantitated by Zeiss LSM 5 PASCAL software (Carl Zeiss MicroImaging GmbH, Germany), and values represent the relative number of colocalizing pixels of caveolin compared with the total number of pixels above threshold. The mean colocalization coefficient was averaged from at least three independent images.

Statistics—Differences in caveolin levels and mean colocalization coefficients were evaluated using an unpaired Student's t test at a 95% confidence level (GraphPad Prism version 3.0; GraphPad Software, San Diego).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
EMP2 Selectively Regulates Expression of Caveolin-1 Protein—Previously, our research group identified a role for EMP2 in regulating expression of caveolin-1 and caveolin-2 in NIH 3T3 cells with either increased or decreased EMP2 protein levels (16). To further define this relationship, we performed quantitative titration analysis of caveolin-1 and caveolin-2 mRNA and protein (26). Total RNA was isolated from NIH 3T3 cells stably transfected with either EMP2 (3T3/EMP2) or a vector control (3T3/V). Samples were diluted serially in 2-fold increments from 1:1 to 1:8, then separated using electrophoresis, and transferred to a nylon membrane. Membranes were probed for expression of caveolin-1 and caveolin-2 (Fig. 1A). Strikingly, increased EMP2 levels reduced caveolin-1 steady state mRNA 3-fold and caveolin-2 mRNA levels by 2-fold. Ethidium bromide staining illustrated relative sample RNA amounts.

We next quantified steady state protein levels of caveolin-1, caveolin-2, and EMP2 in 3T3/EMP2 and 3T3/V cell lines (Fig. 1B). For analysis of caveolin-1, 3T3/V total cell lysates were diluted serially in 2-fold increments from 1:1 to 1:64, whereas 3T3/EMP2 total cell lysates were diluted serially from 1:1 to 1:4. Caveolin-1 protein was decreased by 15-fold in 3T3/EMP2 cells. To analyze caveolin-2 protein levels, 3T3/V total cell lysate was serially diluted from 1:1 to 1:8, whereas 3T3/EMP2 total cell lysate was diluted serially from 1:1 to 1:4. Caveolin-2 protein levels were decreased 2-fold in 3T3/EMP2 cells.


Figure 1
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  		FIGURE 1.
EMP2 reduces expression of caveolin-1 mRNA and protein in NIH 3T3 cells. A, total RNA was isolated from 3T3/EMP2 or vector control 3T3/V cells, diluted serially in 2-fold increments from 1:1 to 1:8, and then assayed by Northern blot for expression of caveolin-1 and caveolin-2. Ethidium bromide staining shows relative sample RNA amounts. B, total cell lysates were titrated as indicated, and samples were assayed by Western analysis for caveolin-1, caveolin-2, and EMP2 protein. C, statistical analysis of differences in mRNA and protein levels of caveolin-1 and caveolin-2. Data were analyzed using a two-tailed, unpaired Student's t test with a 95% confidence interval. Changes in expression between caveolin-1 mRNA and protein are significant, p = 0.038.

 


Figure 2
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  		FIGURE 2.
EMP2 regulates expression of caveolin-1 in human retinal pigment epithelium cells. A, Northern blot analysis of mRNA levels in ARPE-19/EMP2 or vector control ARPE-19/V cells. Ethidium bromide staining shows equal RNA loading. B, Western blot analysis of ARPE-19/EMP2 and ARPE-19/V cell lines for protein expression. Protein lysates were probed for expression of EMP2, caveolin-1, or caveolin-2. beta-Actin indicates equal protein loading. A representative experiment is shown; five independent experiments yielded similar results. C, caveolin-1 expression was quantitated and averaged from five independent experiments. Statistical differences were performed using an unpaired Student's t test. p < 0.0001.

 
Notably, changes in caveolin-1 mRNA and protein levels differed substantially, whereas changes in caveolin-2 mRNA and protein levels were comparable. To determine whether caveolin-1 mRNA and protein level differences were significantly different, a Student's t test was performed. This analysis showed that changes in expression between caveolin-1 mRNA and protein were significant (p = 0.038; Fig. 1C). Thus, EMP2 regulated expression of caveolin-1 at both the mRNA and protein levels. Conversely, regulation of caveolin-2 by EMP2 occurred only at the mRNA level.

EMP2 Regulates Expression of Caveolin-1 in Retinal Pigment Epithelium Cells—To confirm the role of EMP2 in regulating expression of caveolin-1, we examined a human retinal pigmented epithelial cell line, ARPE-19, which endogenously expresses EMP2, caveolin-1, and caveolin-2. ARPE-19 cells were stably transfected with an EMP2 overexpression construct (ARPE-19/EMP2) or a vector control plasmid (ARPE-19/V), and selected for expression using geneticin. Northern blot analysis revealed a 2-fold reduction in caveolin-1 mRNA when EMP2 is overexpressed (Fig. 2A). Conversely, caveolin-2 mRNA levels did not change. Ethidium bromide staining indicated equal RNA loading.

Western blot analysis was then used to examine changes in protein expression levels (Fig. 2B). Cell extracts were assayed for expression of EMP2, caveolin-1, caveolin-2, and beta-actin. Expression of the EMP2/GFP fusion protein is indicated by the presence of two bands, one at the expected 18-kDa size of endogenous EMP2 and another at 46 kDa, the expected size of an EMP2/GFP fusion protein. Consistent with our results in NIH 3T3 cells, caveolin-1 protein levels using an antibody specific for the full-length protein revealed a dramatic 96% reduction in caveolin-1 expression when EMP2 levels were increased by 80%. No change in caveolin-2 expression levels was observed (Fig. 2B). beta-Actin was used to show equal protein loading.

EMP2 Does Not Alter Caveolin-1 Translation Efficiency—Although a number of factors have been shown to control caveolin-1 transcription, little is known about post-transcriptional modes of caveolin-1 regulation (1). Given the dramatic ability of EMP2 to alter caveolin-1 protein levels, we next chose to examine how EMP2 was mediating this effect. To investigate if EMP2 was controlling the rate of caveolin-1 translation, we examined caveolin-1 polysome profiles in 3T3/EMP2 and 3T3/V cell lines. Fig. 3A shows a typical polysome distribution for sucrose gradient eluents. Transcripts that were being translated at the time of extract preparation were polysome-associated and sedimented in fractions 6-10, with the most actively translated transcripts toward the bottom of the gradient. Distribution of mRNA among the gradient fractions is shown by ethidium bromide staining (Fig. 3B). Northern blot analysis revealed that in 3T3/EMP2 cells, the majority of caveolin-1 mRNA was associated with polyribosomal fractions. In 3T3/V control cells, the majority of caveolin-1 mRNA was associated with polyribosomal fractions, whereas a small portion was associated with less dense, polysome-free fractions. These results indicate that EMP2 did not regulate caveolin-1 protein levels by decreasing translation efficiency. As a control, gradient fractions were also analyzed for beta-actin mRNA. Both cell lines showed similar high rates of translation for beta-actin.


Figure 3
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  		FIGURE 3.
EMP2 does not decrease caveolin-1 translation efficiency. Cytoplasmic extracts were fractionated through a sucrose gradient, and 10 1-ml fractions were collected while monitoring the absorbance profile. Total RNA was extracted from each fraction and then equal volumes from each sample were analyzed by Northern blot. A, UV (254 nm) chromatogram of a sucrose gradient eluent showing a typical polysome distribution. B, denaturing agarose gel, stained with ethidium bromide, showing the amount of total RNA present in each of the sucrose gradient fractions. C, Northern blot analysis of the distribution of caveolin-1 mRNA in sucrose gradient fractions reveals that EMP2 does not decrease the association of caveolin-1 mRNA with actively translation polyribosomes. beta-Actin mRNA serves as a control for active translation. Experiments were performed three times with similar results.

 


Figure 4
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  		FIGURE 4.
EMP2 alters the subcellular distribution of caveolin-1. Caveolin-1 distribution in 3T3/EMP2 and 3T3/V cells was assessed using sucrose density centrifugation. Equal volume equivalents were loaded in the cellular fractions to determine the distribution of caveolin-1 in 3T3/EMP2 (A) and 3T3/V (B) cells. These experiments were performed three times, and the caveolin-1 distribution was averaged in 3T3/EMP2 (C) and 3T3/V (D) cells. C, caveolin; PM, plasma membrane; M, microsomal; N, nuclear fractions.

 
EMP2 Alters the Subcellular Distribution of Caveolin-1—To determine whether EMP2 overexpression altered the subcellular distribution of caveolin-1, cells were separated using sucrose gradient centrifugation (Fig. 4). Similar to previously published reports (21, 22), caveolin-1 was expressed in plasma membrane, microsomal, and nuclear fractions in 3T3/V cells (Fig. 4, B and D). Similarly, caveolin-1 is expressed in plasma membrane, microsomal, and nuclear fractions in 3T3/EMP2 cells (Fig. 4, A and C). Interestingly, there is no statistical difference in the caveolin-1 plasma membrane distribution between 3T3/V and 3T3/EMP2 cells (p = 0.40). However, 3T3/EMP2 cells does exhibit a significantly reduced microsomal fraction of caveolin-1 as compared with 3T3/V (p = 0.0062).

EMP2 Does Not Change the Level of Caveolin-1 Phosphorylation—To understand the significance of EMP2 regulation of caveolin-1, phosphorylation of caveolin-1 was assessed. Caveolin-1 maintains an inactive state under normal cellular conditions but becomes phosphorylated at tyrosine 14 in response to cell signaling events (23). 3T3/EMP2 and 3T3/V cell lines were subjected to hyper-osmotic or hydrogen peroxide-induced stress, treated with 600 mM sucrose or 5 mM H2O2 for 10 min, and then subjected to immunoblot analysis (23). As a control for expression levels, we also assayed the relative amount of total caveolin-1. beta-Actin documented equal protein loading.

As shown in Fig. 5, minimal caveolin-1 phosphorylation was detected in untreated 3T3/V cells, whereas no phosphorylated caveolin-1 was detected in untreated 3T3/EMP2 cells. Treatment of cells with either H2O2 or sucrose (longer exposure) resulted in induction of caveolin-1 phosphorylation in 3T3/V cells. Phosphorylated caveolin-1 was also detectable in 3T3/EMP2 cells, although to a lesser degree. When we normalized caveolin-1 phosphorylation in 3T3/EMP2 cells to the total level of caveolin-1 expression, caveolin-1 was phosphorylated to a similar extent in these cells (data not shown). Consequently, caveolin-1 underwent normal levels of phosphorylation, and this mode of post-translational modification would not appear to contribute to reduced caveolin-1 levels in 3T3/EMP2 cells.

EMP2 Levels Alter Turnover of the Long Term Caveolin-1 Pool—Regulation of protein turnover is a key factor in controlling protein expression (27). Given the dramatically reduced protein levels of caveolin-1 in EMP2 overexpressing cells, we next chose to examine if EMP2 might be affecting caveolin-1 stability. To test this, we measured caveolin-1 turnover by pulse-chase analysis followed by immunoprecipitation of caveolin-1 protein (Fig. 6). Strikingly, EMP2 up-regulation dramatically accelerates caveolin-1 degradation. In 3T3/V cells, caveolin-1 half-life was ~5 h. However, upon overexpression of EMP2, the half-life of caveolin-1 decreased to 100 min. In addition, after 24 h no significant levels of caveolin-1 were present in 3T3/EMP2 cells. However, a sizeable fraction (33%) remained in 3T3/V cells. These findings indicate that EMP2 regulates caveolin-1 by accelerating turnover of the long term caveolin-1 compartment.


Figure 5
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  		FIGURE 5.
EMP2 does not inhibit caveolin-1 phosphorylation. Cells were treated with 600 mM sucrose or 5 mM H2O2 for 10 min at 37 °C to induce phosphorylation, and cell lysates were prepared and subjected to immunoblot analysis. Blots were probed with a phospho-specific caveolin-1 polyclonal antibody that recognizes only the tyrosine 14-phosphorylated form of caveolin-1. Immunoblotting with mAb 2297 shows total caveolin-1 protein levels. beta-Actin indicates equal protein loading. Experiment was performed three times with similar results. Untr., untreated.

 


Figure 6
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  		FIGURE 6.
EMP2 alters the turnover of caveolin-1. 3T3/EMP2 and 3T3/V cell lines were incubated with [35S]methionine and -cysteine for 1 h, and chased in medium containing excess unlabeled methionine and cysteine for up to 24 h. A, caveolin-1 was immunoprecipitated from cell lysates, subjected to SDS-PAGE, and radioactivity detected using a PhosphorImager. B, caveolin-1 decay was quantitated with data from five independent experiments. Caveolin-1 signal strength at each time point is expressed as a percentage of signal strength at time 0. Error bars are S.D. Statistical differences between 3T3/EMP2 and 3T3/V were calculated at each time point using an unpaired Student's t test. *, p = 0.0023; **, p = 0.0220; ***, p = 0.0063.

 
EMP2 Does Not Alter Caveolin-1 Proteasome-mediated Degradation—One common mechanism for post-translational regulation involves proteolysis, and a number of proteolytic mechanisms exist in cells (27). One common system for the break down of proteins is via proteasome-mediated ubiquitin-dependent degradation. Indeed, a caveolin-1 family member, caveolin-3, undergoes ubiquitination-proteasomal degradation when it is misfolded because of sequence mutations (28). As EMP2 dramatically altered the stability of caveolin-1, we hypothesized that EMP2 might be increasing proteasome-mediated degradation of caveolin-1. To assess whether the proteasome was responsible for changes in caveolin-1 expression, we treated cells with epoxomicin, a potent and selective proteasome inhibitor, that should prevent degradation and therefore increase the levels of proteins targeted by the proteasome (29). 3T3/EMP2 and 3T3/V cells were treated for 6 h with 100 nM, 1 µM, or 10 µM epoxomicin, a Me2SO-only vehicle control, or left untreated and then assayed for protein levels of caveolin-1 and caveolin-2 by Western blot (30, 31). p53, which is rapidly degraded in the absence of epoxomicin, serves as a control (32, 33), whereas beta-actin levels indicate equal protein loading. Titration of the proteasome inhibitor revealed no significant alternations (p < 0.05) in total caveolin-1 and caveolin-2 protein levels in 3T3/EMP2 and 3T3/V cell lines (Fig. 7). Note that visualization of caveolin-1 in the 3T3/EMP2 cell line required enhanced exposure as compared with the 3T3/V cell line. As a control, p53 levels were only detectable upon proteasome inhibition. Therefore, we conclude that EMP2 is not regulating caveolin-1 by altering proteasome-mediated degradation.

Protease Inhibitors Increase Caveolin-1 Expression—Another common pathway for proteolysis utilizes lysosomes (27, 34). Lysosomes are small organelles that mediate breakdown of proteins using proteases (27), and caveolin proteins have been shown to associate with lysosomal markers (35, 36). To test the possibility that increased EMP2 may be targeting proteins to lysosomes, caveolin-1 levels were assessed in 3T3/EMP2 and 3T3/V cells treated for 4 h with leupeptin (inhibitor of serine and thiol proteases) and/or pepstatin (aspartyl protease inhibitor). In 3T3/EMP2 cells, caveolin-1 expression increased 3-fold with pepstatin but only modestly (50%) with leupeptin. A combination of the two inhibitors further increased caveolin-1 to 5- and 7-fold with 1 or 5 µM inhibitors, respectively (Fig. 8). In 3T3/V cells, each inhibitor alone affected caveolin-1 levels minimally; combination of the inhibitors increased caveolin-1 by 2-2.5-fold with 1 or 5 µM inhibitors, respectively (Fig. 8), but this did not reach statistical significance (p = 0.08). It should be noted that caveolin-1 immunoblot images show signal exposure equivalent to 3T3/EMP2 conditions. For quantitation with images in linear range, shorter times were used. These findings suggest that EMP2 promotes lysosomal degradation of caveolin-1, predominantly via aspartyl protease activity.


Figure 7
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  		FIGURE 7.
EMP2 does not alter proteasome-mediated degradation of caveolin-1. A, 3T3/EMP2 and 3T3/V cell lines were treated for 6 h with 100 nM, 1 µM, or 10 µM epoxomicin, a Me2SO only vehicle control, or were left untreated. Cells were then harvested and assayed for protein levels of caveolin-1 and caveolin-2 by immunoblot. p53 and beta-actin immunoblots serve as controls. B, caveolin-1 expression was normalized to beta-actin levels from three independent experiments in both 3T3/V and 3T3/EMP2 cells. Results are expressed as the percent increase in caveolin-1 expression over the Me2SO vehicle control. Error bars are S.E.

 


Figure 8
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  		FIGURE 8.
EMP2 increases lysosomal degradation of caveolin-1. A, 3T3/EMP2 cells or 3T3/V cells were incubated with 1 or 5 µM leupeptin (Leu) or pepstatin (Pep) alone or in combination (equal µM equivalents) for 4 h. The experiment was performed three times, and a representative graph is depicted. B and C, caveolin-1 protein expression was quantitated using ImageJ 1.36b software and normalized to beta-actin expression in 3T3/EMP2 cells (B) and 3T3/V cells (C). Caveolin-1 immunoblot images show signal exposure equivalent conditions for 3T3/EMP2 and 3T3/V. For quantitation with images in linear range, shorter times were used for 3T3/V. Error bars are S.E. Statistical analysis of the data was performed using an unpaired t test. *, p = 0.01; p = 0.003 (5 µM Leu + Pep versus 0 µM inhibitor, 3T3/V).

 
Effect of EMP2 on the Intracellular Distribution of Caveolin-1—To confirm the targeting of caveolin-1 to lysosomes by EMP2, we examined the intracellular localization of caveolin-1 in 3T3/EMP2 in comparison with 3T3/V cells. As in most cell types, caveolin-1 in 3T3/V cells is localized to the plasma membrane with minimal intracellular staining (data not shown; see Ref. 18). In contrast, 3T3/EMP2 cells displayed a punctate staining of the cytoplasm with little plasma membrane staining (Fig. 9A). To identify the aberrant intracellular sites bearing caveolin-1, cells were costained for Golgi apparatus ({gamma}-adaptin), early endosomes (EEA-1), and lysosomes (LAMP-1). In 3T3/EMP2 cells, {gamma}-adaptin displayed normal peri-nuclear staining, but showed little colocalization with caveolin-1 (Fig. 9A). 3T3/V cells exhibited greater colocalization (36%) than 3T3/EMP2 (15%), indicating that more caveolin-1 was retained in the Golgi apparatus in 3T3/V cells (Fig. 9A and Table 1).


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  		TABLE 1
Colocalization of caveolin-1 with intracellular compartments

Values represent the number of pixels bearing both caveolin-1 and the compartment marker, compared with the total number of pixels bearing the compartment marker alone as calculated by Zeiss 5 PASCAL software. For LAMP-1 and caveolin-1 colocalization, samples were treated for 1 h with leupeptin and pepstatin. The mean of at least three independent images is provided.

 
However, when we assessed early endosomes (EEA-1), more than 60% of EEA-1 signal colocalized with caveolin-1 in 3T3/EMP2 (Fig. 9B and Table 1). In contrast, little caveolin-1 colocalized with EEA-1 in 3T3/V cells (Fig. 9B and Table 1). Proteins targeted to early endosomes are either recycled back to the cell surface or to late endosomes/lysosomes (34, 37). To confirm caveolin-1 in the lysosomal compartment, 3T3/EMP2 cells were treated with the protease inhibitors pepstatin and leupeptin for 1 h prior to staining to prevent caveolin-1 degradation. Dramatically following treatment, caveolin-1 was colocalized with LAMP-1 (Fig. 9C), representing 54% of the LAMP-1+ compartment (Table 1). In contrast, caveolin-1 in 3T3/V exhibited significantly less colocalization with lysosomes (23%; Fig. 9C and Table 1). The increased colocalization of caveolin-1 within lysosomes in 3T3/EMP2 cells confirm that elevated EMP2 expression leads to increased trafficking of caveolin-1 into lysosomes via early endosomes.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Caveolin-1 is an integral component of plasma membrane caveolae and plays a role in numerous cellular functions, including lipid transport, membrane trafficking of caveolin-2, and caveolae formation (4, 5). Caveolin-1 expression is transcriptionally controlled by a variety of cellular factors, but post-transcriptional modes of regulation have yet to be identified. Our present findings demonstrate a new post-transcriptional mechanism of caveolin-1 regulation. Specifically, they indicate that caveolin-1 can be redirected after initial biosynthesis to EEA-1 and LAMP-1 compartments, which are associated with rapid proteolytic degradation. Moreover, this early degradation fate is facilitated by increased levels of EMP2 expression.

Quantitative analysis of EMP2 overexpression on caveolin levels in NIH 3T3 cells indicated significant down-regulation of caveolin-1 expression mediated at both the RNA and protein levels, whereas decreases in caveolin-2 expression appear to be mediated only at the RNA level. Alternate expression profiles of caveolin-1 and caveolin-2 are not surprising, as caveolin-1 and caveolin-2 are differentially expressed in different cell types (3). Furthermore, Scherer et al. (3) have reported that NIH 3T3 cells transformed with v-Abl display dramatically decreased caveolin-1 levels, whereas caveolin-2 levels are unaffected. Given the vital role of caveolin-1 in the formation of caveolae and the accessory role of caveolin-2, it is logical that expression of these proteins would be differentially regulated.


Figure 9
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  		FIGURE 9.
EMP2 alters the intracellular distribution of caveolin-1. Confocal microscopy was used to examine the distribution of caveolin-1 in 3T3/V and 3T3/EMP2 cells. Cells were assayed for colocalization as follows. A, caveolin-1 (red) with {gamma}-adaptin (FITC); B, caveolin-1 (red) and EEA-1 (FITC); or C, caveolin-1 (red) and LAMP-1 (FITC). To determine LAMP-1 and caveolin-1 colocalization, samples were treated for 1 h with the lysosomal inhibitors leupeptin and pepstatin. Pictures were taken at a magnification of x600. The boxed area in 3T3/EMP2 cells is enlarged x3 to show the colocalization of caveolin-1 with intracellular markers (inset). Scale bar, 10 µm.

 
Association of caveolin-1 mRNA with polyribosomal subunits indicated that EMP2 did not alter caveolin-1 translation efficiency. Similarly, proteasome inhibitor assays excluded EMP2-dependent, proteasome-mediated degradation of caveolin-1 protein levels. Strikingly, when we analyzed caveolin-1 expression after 1 h of biosynthetic labeling, its turnover rate was dramatically affected by EMP2 expression levels. 3T3/EMP2 cells (t1/2 = 1.5 h) exhibited 3.3 times faster caveolin-1 degradation compared with 3T3/V cells (t1/2 = 5 h). When we next assessed lysosomal degradation of caveolin-1, normal levels of protein were restored following 4 h of treatment with typical proteolytic inhibitors. Finally, confocal analysis revealed that in cells with elevated EMP2, caveolin-1 predominantly localized with EEA-1 and LAMP-1 compartments.

The process by which EMP2 promotes targeting of caveolin-1 to this degradation pathway is unknown. Lysosomal inhibitors modestly increased caveolin-1 levels at high concentrations, suggesting that the endogenous protein traffics to the lysosome. Moreover, pepstatin treatment alone increased caveolin-1 in both 3T3/EMP2 and 3T3/V, suggesting that caveolin-1 degradation is primarily via aspartyl proteases. Previous studies have found that caveolin-1 can colocalize with lysosomes (35, 36). We observed that a small percentage of caveolin-1 was delivered to lysosomes in 3T3/V cells, suggesting that degradation of caveolin-1 via lysosomes is not an artifact of EMP2 overexpression. It is also interesting to note that confocal analysis and subcellular fractionation of 3T3/EMP2 cells reveal a significant fraction of caveolin-1 on the plasma membrane. This suggests that caveolin-1 traffics to the membrane in these cells, where it is then consequently transported to lysosomal compartments.

Biologically, the reciprocal regulation of EMP2 and caveolin-1 levels has important implications for the surface receptor repertoire. For example, EMP2 strongly associates with P2X7 (38). P2X7 is a ligand-gated channel implicated in inflammasomes, a structure mediating cytoplasmic protein translocation to proteolytic endosomes (39, 40). One speculation is that EMP2 might play a role in structures that translocate proteins to an endosomal compartment and proteolytic fate.

It is known that caveolin-1 and EMP2 strongly modulate the surface availability and lipid raft association of large but distinct groups of membrane proteins. Le and co-workers (41, 42) reported that caveolin-1 acts as a negative regulator by preventing caveolae from internalizing. Upon removal of caveolin-1, caveolae internalize more rapidly (41). Consequently, by decreasing caveolin-1 protein levels, EMP2 may be facilitating increased internalization of caveolae-associated plasma membrane proteins. Conversely, increased EMP2 levels promote the surface expression of several integrin isoforms, GPI-linked proteins, and major histocompatibility complex class 1 proteins (16-19). The regulation by EMP2 of caveolin-1 expression thus would appear to profoundly reshape the cellular surface receptor repertoire and its attendant responsiveness to the local environment.

Regulation of caveolin-1 expression by EMP2 may also play a role in cellular transformation. Reduced expression of caveolin-1 has been linked to cancer in a number of cell types (43). Caveolin-1 was first discovered as a primary substrate for tyrosine phosphorylation in Rous sarcoma virus-transformed chicken embryonic fibroblasts, suggesting a role for caveolin-1 as a tumor suppressor protein (44). Transformation of NIH 3T3 cells with various activated oncogenes (v-Abl, Bcr-Abl, and H-Ras (G12V)) down-regulates caveolin-1 (10, 26), and caveolin-1 is often reduced in breast and prostate cancer cell lines (45-48), sometimes in association with hypermethylation (45, 49). Mechanistically, the tumor suppressor role of caveolin-1 may reflect its inhibition of signaling activity of proto-oncogenes, presumably through its roles in surface receptor expression and signaling (43). Indeed, specific knockdown of caveolin-1 is sufficient to confer a tumorigenic phenotype in NIH 3T3 cells (11).

We recently reported that elevated expression of EMP2 is a central molecular feature of endometrial carcinoma with unfavorable outcome (50). Closely related proteins in the PMP22/GAS3 family are also linked to malignant phenotype as follows: PMP22 in pancreatic cancer (51), osteosarcoma (52-54), mammary carcinoma (55), schwannoma (56); EMP1 in brain tumors (57); and EMP3 in prostate cancer (58), neuroblastoma, and glioma (59). The basis of this association with aggressive malignant phenotype is largely unknown. However, we speculate that reciprocal down-regulation of caveolin-1 may define one of the pathways linking overexpression of these tetraspan proteins to aggressive malignancy.


   FOOTNOTES
 
* This work was supported by National Institutes of Health Grants GM7185 (to A. F.), AI52031 (to S. M.), HD48540 (to J. B.), and CA9120 (to M. W.), the Lalor Foundation (to M. W.), the Giannini Family Foundation (to M. W.), and the Ruzic Foundation (to J. B.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

1 Both authors contributed equally to this work. Back

2 To whom correspondence should be addressed: Pathology and Laboratory Medicine, 13-222 Center for Health Sciences, Geffen School of Medicine, UCLA, Los Angeles, CA 90095. Tel.: 310-794-7953; Fax: 310-825-5674; E-mail: jbraun{at}mednet.ucla.edu.

3 The abbreviations used are: DMEM, Dulbecco's modified Eagle's medium; PBS, phosphate-buffered saline; GFP, green fluorescent protein; FITC, fluorescein isothiocyanate. Back


   ACKNOWLEDGMENTS
 
We thank Agnes Nagy for help with the subcellular fractionation experiments. The UCLA Jonsson Comprehensive Cancer Center flow cytometry core was recipient of National Institutes of Health Grant CA16042.



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HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
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Copyright © 2007 by the American Society for Biochemistry and Molecular Biology.
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