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Am J Physiol Cell Physiol 293: C294-C304, 2007. First published April 4, 2007; doi:10.1152/ajpcell.00413.2006
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PROTEIN AND VESICLE TRAFFICKING, CYTOSKELETON

Role of the scaffold protein RACK1 in apical expression of CFTR

Michael Auerbach and Carole M. Liedtke

Warren Alan Bernbaum, M.D., Center for Cystic Fibrosis Research, Departments of Pediatrics at Rainbow Babies & Children Hospital and of Physiology & Biophysics, Case Western Reserve University, Cleveland, Ohio

Submitted 2 August 2006 ; accepted in final form 2 April 2007


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Previous studies from this laboratory demonstrated a role for protein kinase C (PKC){varepsilon} in the regulation of cAMP-dependent cystic fibrosis transmembrane regulator (CFTR) Cl channel function via binding of PKC{varepsilon} to RACK1, a receptor for activated C kinase, and of RACK1 to human Na+/H+ exchanger regulatory factor (NHERF1). In the present study, we investigated the role of RACK1 in regulating CFTR function in a Calu-3 airway epithelial cell line. Confocal microscopy and biotinylation of apical surface proteins demonstrate apical localization of RACK1 independent of actin. Mass spectrometric analysis of NHERF1 revealed copurification of tubulin, which, in in vitro binding assays, selectively binds to NHERF1, but not RACK1, via a PDZ1 domain. In binding and pulldown assays, we show direct binding of a PDZ2 domain to NHERF1, pulldown of endogenous NHERF1 by a PDZ2 domain, and inhibition of NHERF1-tubulin binding by a PDZ1 domain. Downregulation of RACK1 using double-stranded silencing RNA reduced the amount of RACK1 by 77.5% and apical expression of biotinylated CFTR by 87.4%. Expression of CFTR, NHERF1, and actin were not altered by treatment with siRACK1 or by nontargeting control silencing RNA, which, in addition, did not affect RACK1 expression. On the basis of these results, we model a RACK1 proteome consisting of PKC{varepsilon}-RACK1-NHERF1-NHERF1-tubulin with a role in stable expression of CFTR in the apical plasma membrane of epithelial cells.

silencing RNA; downregulation; biotinylation; tubulin; NHERF1; tailless cystic fibrosis transmembrane regulator; PDZ domain

THE CYSTIC FIBROSIS TRANSMEMBRANE regulator (CFTR) is a Cl channel located in the apical membrane of epithelial cells, where it plays a key role in fluid homeostasis. Defects in CFTR channel activity or biosynthesis lead to the development of the disease cystic fibrosis, which is characterized by abnormal Cl secretion as well perturbed transport of other electrolytes. Activity of CFTR is tightly regulated by phosphorylation of its R domain by cAMP-dependent protein kinase A (PKA) and protein kinase C (PKC) (10). PKC-dependent regulation of CFTR is subtle, apparently requiring constitutive activity of a Ca2+-independent PKC isotype PKC{varepsilon} (21). We found that downregulation or inhibition of activated PKC{varepsilon} reduced or prevented cAMP-dependent activation of CFTR (17) but did not alter CFTR or Na-K-2Cl cotransporter protein and mRNA expression (18). We also reported binding of PKC{varepsilon} to the scaffold protein RACK1, a receptor for activated C kinase, and binding of RACK1 with NHERF1 at the PDZ1 domain of NHERF1 (20) and 5th WD repeat of RACK1 (22).

In the present study, we ask how a protein complex consisting of PKC{varepsilon}-RACK1-NHERF1 regulates cAMP-dependent activation of CFTR. CFTR interacts with several proteins, including ion channels, protein kinases, and phosphatases, and ATP transporters and is thought to exist in macromolecular complexes with scaffold proteins and their binding partners; many of these interactions involve PDZ-domain proteins, such as NHERF1 and beta2-adrenergic receptor (reviewed in Ref. 11). The scaffold protein, NHERF1, binds to CFTR at the COOH-terminus in a PDZ-binding domain with the amino acid sequence Asp-Thr-Arg-Leu (16). Its role in CFTR activation and trafficking is less clear (3, 29, 35). Hence, we focus in the current study on a role for RACK1 in the regulation of CFTR function.

RACK1 is a scaffold protein that stewards protein signaling requiring defined subcellular localization and substrate regulation (26). RACK1 consists of seven conserved repeating units consisting of 40-60 amino acids bounded by two internal dipeptide sequences, Gly-His (GH) and Try-Asp (WD). On the basis of the crystal structure of Gbeta{gamma}, another member of the WD40 repeat proteins, RACK1 is predicted to form a rigid circular seven-blade beta-propeller structure (33). The WD repeats are implicated in the binding of RACK1 to its protein partners (21, 25), which include PKC betaII and-{varepsilon} isotypes, phosphodiesterase PDE4D5, src tyrosine kinases, integrin beta-subunit, type 1 interferon receptor, insulin-like growth factor 1 receptor, inositol 1,4,5-trisphosphate receptors (30), PTPµ protein-tyrosine phosphatase (28), Gbeta{gamma} (7), and dopamine transporter (15). The specific interaction between RACK1 and its binding partners is complex and may involve multiple points of contact between the two proteins as shown for PDE4D4, PKC{varepsilon}, PKCbeta, beta-integrin, and interferon receptor (26). More importantly, the beta-propeller structure of RACK1 is thought important in the binding of multiple proteins, as we have found for NHERF1 and PKC{varepsilon} in airway epithelial cells. Binding of RACK1 and NHERF1 suggests a role for RACK1 in the functional expression of CFTR in the apical plasma membrane.

The goal of the current study was to determine whether this was, indeed, the functional role of the scaffold protein RACK1 in airway epithelial cells. We selected a Calu-3 cell line for study because this human serous cell line expresses CFTR and proteins involved in regulating its function, including NHERF1 (21). In addition, Calu-3 cells have been used by our laboratory (17, 18, 21) and others (13) to study protein kinase-dependent regulation of cAMP-dependent activation of CFTR. In the present study, we show that NHERF1 binds directly to tubulin and downregulation of RACK1 reduces apical expression of biotinylated CFTR. On the basis of these results, we model a "RACK1 proteome" consisting of PKC{varepsilon}-RACK1-NHERF1-NHERF1-tubulin, which localizes RACK1 to the apical plasma membrane compartment.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell Isolation and culture. Calu-3 cells were grown in a submerged cell culture on 100 mm2 tissue culture plastic, as described previously (21). Cell cultures were grown at 37°C under 5% CO2 humidified air on tissue-culture treated plastic dishes. Cells were used for experiments when confluence was reached, typically 5–6 days after subculture.

Mass spectrometry of NHERF1. To recover sufficient NHERF1 for mass spectrometry, NHERF1 was immunoprecipitated from 10 ml Calu-3 TCL using 40 µl anti-EBP50 antibody (Abcam). Immunoprecipitates were resolved on a 4–15% SDS-PAGE gradient slab gel and stained with Coomassie blue. Endogenous NHERF1 was detected as a single 50 kDa protein band. NHERF1 and proteins coimmunoprecipiating with NHERF1 were identified by sequencing the peptides formed by an in-gel tryptic digestion using liquid chromatography-tandem mass spectrometry by the Mass Spectrometry Laboratory for Protein Sequencing at the Cleveland Clinic Foundation Lerner Research Institute (Dr. M. Kinter, Director) (14). In brief, the band was cut from the gel, reduced, and alkylated with DTT and iodoacetamide. The protein was digested with trypsin. Peptides formed by the digestions were extracted from the polyacrylamide and concentrated to 30 µl. The liquid chromatography-tandem mass spectrometry instrument was Finnigan LTQ linear ion trap mass spectrometer. The HPLC column was a self-packed 8 cm x 75 µm inner diameter C18 reversed-phase capillary chromatography column. Ten microliter volumes of the extract were injected and peptides eluted from the column by an acetonitrile/0.05 M acetic acid gradient at a flow rate of 1 µl/min were introduced into the source of the mass spectrometer on-line. The microelectrospray ion source is operated at 4.0 kV. The digest was analyzed using the data dependent multitask capability of the instrument acquiring full scan mass spectra to determine peptide molecular weights and product ion spectra to determine amino sequence in successive instrument scans. This mode of analysis produces ~2,500 collisionally induced dissociation spectra of ions ranging in abundance over several orders of magnitude. The data were analyzed by using all collisionally induced dissociation (CID) spectra collected to search the NCBI nonredundant database with the search program Mascot. All matching spectra were verified by manual interpretation.

Immunofluorescence of RACK1 and actin. Calu-3 cell monolayers were dual labeled for RACK1 and actin, as previously described (21). Briefly, cells were washed with PBS, fixed in fresh 4% paraformaldehyde for 15 min at room temperature, washed three times with PBS, and permeabilized with 0.2% Triton X-100 in 10% normal goat serum in PBS. The fixed, permeabilized cells were stained for 1 h at room temperature with a monoclonal IgM antibody directed to RACK1 (1:200 dilution) and Texas red-conjugated to phalloidin (1:80). Cell monolayers were washed three times with PBS. Secondary Oregon Green-conjugated anti-rabbit antibody for RACK1 was applied at 1:200 dilution for 1 h at room temperature. After three final washes with PBS, the polyester filters were carefully cut from their support inserts and mounted in Slow Fade mounting medium on glass microscope slides. Fluorescence was analyzed using a LSM410 confocal scanning microscope (Zeiss) equipped with an external argon-krypton laser. Optical sections of 512 x 512 pixels were digitally recorded in x16 line-averaging mode and z-sectioning was done in 1 µm increments. Images were processed for reproduction using PhotoShop software (Adobe Systems, Mountain View, CA).

Construction and expression of recombinant proteins. Recombinant RACK1 with a His6 tag, a PKA site, a thrombin cleavage site, and an HA-tag was expressed in Sf9 insect cells, as previously described (21). In brief, insect cells were maintained at 27°C in Grace's insect medium supplemented with 10% fetal bovine serum and 10 µg/ml gentamicin. Viral stocks, provided by Dr. Susan Brady-Kalnay (Case Western Reserve University), were maintained and used to transfect Sf9 cells and to express recombinant human RACK1. Three days after infection, recombinant RACK1 was isolated from cell lysates using Talon Metal Affinity Resin and evaluated by immunoblot assay.

GST tagged RACK1 was constructed using RACK1 cDNA cut from pAcHLT-c baculovirus transfer vector with full length RACK1 at Stu1 and NotI restriction sites and ligated into a pGEX-6P-1 expression vector (Amersham) at SmaI and NotI sites. pGEX-RACK1 was transformed in competent DH5{alpha} cells. GST-tagged RACK1 was expressed and verified by immunoblot analysis for RACK1 using a monoclonal antibody directed against RACK1. GST-RACK1 was detected as a 63-kDa protein. This molecular mass is consistent with the addition of the GST tag.

NHERF1 in a pGEX4T-1 vector and PDZ1 (amino acid residues 1-139) and PDZ2 (amino residues 132-299), each in a pGEX6P-1 vector, were expressed in DH5{alpha} cells, as previously described (20). Each protein was GST epitope tagged at the NH2 terminus. In brief, proteins were expressed in overnight cultures of competent DH5{alpha} cells by addition of 100 µM isopropyl-beta-D-thiogalactopyranoside and 3–6 h incubation at 37°C, cells were harvested. Recombinant protein was purified using B-PER (Pierce) extraction followed by affinity chromatography with glutathione-Sepharose B beads. The fusion protein was evaluated by immunoblot analysis for the GST tag using a polyclonal antibody to GST, or, for GST-NHERF1, a polyclonal antibody to NHERF1. Recombinant proteins displayed molecular masses of 50 kDa for GST-PDZ domain and 75 kDa for GST-NHERF1. For some experiments, the GST tag was cleaved from NHERF1 using thrombin and from PDZ domains using PreScission enzyme according to manufacturer instructions.

Pulldown analysis and immunoprecipitation. Pulldown of endogenous tubulin from Calu-3 total cell lysates was performed using anti-tubulin antibody conjugated to agarose beads. Cells were grown to confluence, serum deprived overnight, and washed with ice-cold PBS. Cells were lysed in 1 ml lysis buffer consisting of 100 mM NaCl, 50 mM NaF, 50 mM Tris·HCl, pH 7.5, 1% Nonidet P-40, 0.25% sodium deoxycholate, 1 mM EDTA, and 1 mM EGTA, 1 mM Na vanadate, and protease inhibitor cocktail set III (Calbiochem). Lysates were clarified by pretreatment with agarose beads then incubated at 4°C for 1 h with 20-µl bead slurry. In some experiments, total cell lysates were preincubated with 50 µg GST-PDZ1 domain or 50 µg GST-PDZ2 domain at 30°C for 40 min before the addition of beads to allow binding of a PDZ domain to tubulin. Beads were recovered by centrifugation, washed extensively with PBS, and resuspended in Laemmli buffer. Samples were heated for 5 min in a boiling water bath, cooled, then subjected to 4–15% SDS-PAGE. Protein bands were transferred to polyvinylidene difluoride (PVDF) membrane paper for immunoblot analysis for NHERF1 using a polyclonal antibody to NHERF1. Protein bands immunoreactive to specific antibodies were detected using enhanced chemiluminescence. As a control for the efficiency of tubulin pulldown, immunoblots were reprobed with antibody directed against tubulin.

Proteins were immunoprecipitated from Calu-3 total cell lysate, as previously described (20, 21). Total cell lysates were clarified as described above then incubated at 4°C for 1 h with antibodies directed against the protein of interest. Immune complexes were recovered using protein A beads. Immune complexes attached to washed beads were heated to boiling for 5 min in Laemmli buffer then subjected to SDS-PAGE on 4–15% gradient slab gels for immunoblot analysis, as indicated in figure legends.

Monomeric and polymeric tubulin solutions. Monomeric tubulin (M-tubulin) was prepared by mixing 20-µl stock tubulin solution at 5 mg/ml (Cytoskeleton) with 80 µl G-PEM buffer, consisting of 80 mM 1,4 piperazinediethane sulfonic acid, pH 6.9, 1 mM GTP, 0.5 mM MgCl2, 1 mM EGTA, 0.1 µM sodium vanadate, and protease inhibitor cocktail (Calbiochem). Polymeric tubulin (P-tubulin) was prepared by preheating stock tubulin at 37°C for 20 min in G-PEM buffer supplemented with 10% glycerol to promote microtubule formation. To stabilize polymerization, 200 µl of prewarmed G-PEM/glycerol buffer was mixed with 2 µl 2 mM Taxol and incubated at room temperature for 20 min. Immediately, the Taxol solution (final concentration 20 µM) was added to the microtubules and the final mixture kept at room temp until use.

In vitro binding assays. Binding of recombinant GST-NHERF1, GST-PDZ1 domain, GST-PDZ2 domain, HA-His6-RACK1, or GST-RACK1 to tubulin was achieved by immobilizing 0.5 µg of monomeric or polymeric tubulin onto PVDF membrane paper in a slot blot apparatus then overlaying the paper with varying concentrations of a protein. Membrane papers were incubated at room temperature for 25 min, and unbound material was removed by extensive washing. Bound protein was detected by immunoblot analysis using polyclonal antibodies directed to the GST tag or a monoclonal antibody to RACK1.

Cell surface biotinylation. Polarized Calu-3 cell monolayers were grown on 24-mm diameter Transwell permeable supports. Cell-surface proteins were biotinylated using EZ-Link sulfosuccinimidyl-2-(biotinamido)ethyl-1,3-dithiopropionate (NHS-SS-biotin, 3 mg/ml in PBS, pH 8.2; Pierce) (24). Cells were rapidly cooled to 4°C, washed in PBS, pH 8.2, supplemented with 0.1 mM CaCl2 and 1 mM MgCl2, and then incubated with 3 mg sulfo-NHS-SS-biotin/ml for 30 min at 4°C. Nonreacted sulfo-NHS-SS-biotin was quenched by washing cells with PBS, pH 8.2, containing 100 mM glycine, 0.1 mM CaCl2, and 1 mM MgCl2. Cells were harvested in CFTR lysis buffer and biotinylated proteins isolated using streptavidin-agarose beads (streptavidin beads, Pierce), eluted into Laemmli SDS sample buffer supplemented with 50 mM dithiothreitol, and separated on 4–15% SDS-PAGE gradient slab gels. Biotinylated CFTR was detected by immunoblot analysis using a monoclonal antibody directed to the COOH-terminus of CFTR. In experiments in which RACK1 was downregulated (see below), supernatants from streptavidin pulldowns were subjected to gel electrophoresis and immunoblot analysis for RACK1, CFTR, NHERF1, and actin. Exposed bands were quantitated by densitometry.

Downregulation of RACK1. Endogenous RACK1 was downregulated using double stranded SMARTpool small interfering (si)RNA oligonucleotides (Dharmacon RNA Technologies). Cells were grown in culture to 60–75% confluence, collected after trypsinization, pelleted and washed twice with PBS at room temperature. Cells (6 x 106) were resuspended in 100-µl transfection solution (Amaxa Biosystems, kit "V"), mixed with 9.0 pmol siRNA oligonucleotides specific for RACK1 or, as a control, siCONTROL Non-Targeting siRNA (Dharmacon RNA Technologies). The suspension was transferred to a cuvette and electroporated using program T-024 (Amaxa Biosystems). Cells were immediately diluted with 500 ml culture medium, prewarmed to 37°C, and plated onto 24-mm diameter filter inserts at a seeding density of 2 x 106 cells/filter. Preliminary experiments were performed to determine a 48-h incubation period as optimal for maximal loss of RACK1. At this time, the expression of biotinylated CFTR was determined, as described above.

Data analysis. Immunoreactive protein bands were quantitated using a VersaDoc Imaging System (Bio-Rad). Readings from exposed bands are corrected against a blank area on the same blot of the same area. Where indicated, data were normalized by calculating a ratio of (GST signal)/(tubulin or NHERF1 signal). Ratios are reported as mean ± SE. Data are representative of at least three or more experiments, unless otherwise stated, and treatment effects were evaluated using a two-sided Student's t-test.

Materials. Polyclonal anti-PKC{varepsilon}, anti-GST and anti-HA- antibodies, anti-GST and anti-HA antibodies conjugated to agarose beads, horseradish peroxidase-coupled secondary antibodies (anti-rabbit IgG, anti-goat IgG, anti-mouse IgG, and anti-mouse IgM) and protein L-agarose were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Anti-EBP50 antibody was obtained from Abcam (Cambridge, MA) and anti-mouse RACK1 monoclonal antibody from Transduction Laboratories (Lexington, KY). Paclitaxel (taxol) was purchased from Cytoskeleton (Denver, CO). A mixture of {alpha}- and beta-tubulin, purchased from Cytoskeleton, was reconstituted in DMSO at 5 mg/ml concentration. Polyclonal anti-beta-tubulin antibody was obtained from Sigma-Aldrich (St. Louis, MO). Sepharose and agarose beads and tissue culture supplies were purchased from Invitrogen-GIBCO (Gaithersburg, MD), and precast 4–15% gradient slab gels from Biorad. An enhanced chemiluminescence reagent was purchased from Amersham (Piscataway, NJ). All other chemicals were reagent grade.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Immunofluorescence of RACK1 and actin. In previous studies, we observed colocalization of RACK1 and CFTR in the apical membrane of Calu-3 cells but did not detect direct binding of RACK1 to CFTR (20). RACK1 acts as a scaffold protein with the potential to bind seven different proteins, one on each WD repeat. CFTR function and trafficking has been linked to the actin cytoskeleton (6, 31). We now ask whether RACK1 colocalizes with actin at the apical region of Calu-3 cells. Polarized Calu-3 cells were subjected to dual label immunofluorescence for actin and RACK1 and confocal microscopy. The results are shown in Fig. 1 as merged images of en face and xz-plane reconstruction images of the apical region. Actin is clearly detected in the apical region of Calu-3 cells and as a network of fluorescent stain outlining the cell periphery. RACK1, as seen previously, is localized to the apical region of airway epithelial cells. Colocalization of RACK1 and actin, indicated by an orange-red fluorescence, was not observed. Hence, we conclude that RACK1 does not directly bind to actin and that RACK1 localization to the apical membrane involves other protein(s).


Figure 1
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  		Fig. 1. Immunofluorescence of receptor-activated C kinase-1 (RACK1) and actin. Serum-deprived Calu-3 cells, grown on filter inserts, were fixed in 4% paraformaldehyde, permeabilized in 0.2% Triton X-100, and incubated with primary antibodies directed against RACK1 and Texas red-conjugated to phalloidin for 60 min before adding Oregon green-conjugated secondary antibody for RACK1 for 60 min. En face computer-generated image of an apical plane through the cell monolayer is shown. The merged image shows distinct green (RACK1) and red (actin) fluorescent patterns. RACK1 is detected only in the apical region of airway epithelial cells. Actin is detected in the apical region and as a network of fluorescent stain outlining the cell periphery. We do not detect a typical orange-red fluorescence associated with colocalization of RACK1 and actin. Cells incubated with secondary antibody alone displayed no image (data not shown).

 
Mass spectrometric analysis of endogenous NHERF1. To identify other potential binding partners, mass spectroscopic analysis was performed on immunoprecipitated NHERF1. To do this, NHERF1 was isolated from a large volume (10 ml) of Calu-3 total cell lysate and subjected to SDS-PAGE on 4–15% gradient slab gels. NHERF1 was clearly distinguishable after immunoblotting with specific antibody (Fig. 2, left panel). A duplicate lane was stained with Coomassie blue and the protein bands subjected to mass spectrometric analysis as described in MATERIALS AND METHODS (Fig. 2, right panel). The MALDI spectrum revealed seven peptides corresponding to human NHERF1 and eight peptides predicted, by sequence, for beta-tubulin, a 50 kDa monomer of microtubules.


Figure 2
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  		Fig. 2. Mass spectrometric analysis of Na+/H+ exchanger regulatory factor (NHERF1) from Calu-3 cells. Calu-3 cells were grown to confluence, serum-deprived overnight, rapidly washed in ice-cold PBS, and solubilized in lysis buffer. To purify sufficient NHERF1 by mass for analysis by mass spectrometry, NHERF1 was immunoprecipitated from 10 ml total cell lysate (TCL), subjected to SDS-PAGE on 4–15% gradient slab gels, and stained with Coomassie blue. As control, an aliquot of immunoprecipitated protein was immunoblotted for NHERF1 (left lane). NHERF1 was detected at the expected molecular mass of 50 kDa. Coomassie blue-stained gels revealed a prominent protein band at 50 kDa and minor bands at lower molecular mass (right lane). Protein bands were cut from the Coomassie-stained gel and subjected to mass spectrometric analysis, as described in MATERIALS AND METHODS.

 
Direct binding of NHERF1 with tubulin. To determine whether NHERF1 directly binds to tubulin, we performed solid phase slot blot assays using commercially available tubulin and recombinant GST-tagged NHERF1, and, as a control, RACK1. Monomeric and polymeric tubulin were prepared immediately before use and vacuumed onto PVDF paper then overlaid with solutions containing GST-NHERF1 or GST-RACK1. NHERF1, but not RACK1, was observed to bind to monomeric and polymeric tubulin (Fig. 3A). Immunoblots were reprobed for tubulin and quantitated to allow normalization of the GST signal to the tubulin signal. NHERF1 binding to polymeric tubulin is 1.78-fold higher than binding to monomeric tubulin (P < 0.001). NHERF1 binds to tubulin in a dose-dependent manner with an EC50 value of 2.8 µg, or a nominal 0.03 nmol (Fig. 3B). We detected very little binding of RACK1 to monomeric or polymeric tubulin; ratios obtained in Fig. 3B were not significantly different than zero. Because of the preferential binding of NHERF1 to tubulin, we selected NHERF1 for additional studies as described below.


Figure 3
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  		Fig. 3. Direct binding of NHERF1 to tubulin. A: direct binding of NHERF1 and/or RACK1 to tubulin was tested in a solid phase slot blot binding assay. Monomeric (M-tubulin) and polymeric (P-tubulin) tubulin were prepared just before use and 0.5 µg vacuumed onto PVDF paper. Tubulin was overlaid with 2.0 µg glutathione S-transferase (GST)-NHERF1 or GST-RACK1 and incubated at RT for 25 min. Unbound material was removed by washing and bound protein detected by immunoblot analysis for the GST-tag. Membrane paper was reprobed for tubulin. Exposed bands were quantitated by densitometry and data normalized as the ratio of (GST signal)/(tubulin signal) for each slot. Results are shown for a typical solid phase slot blot binding assay in which experimental conditions are replicated in quadruplicate. Data are reported as means ± SE for the number of replicates. Quantitation by densitometry indicates that NHERF1 binding to P-tubulin is 1.78-fold higher than binding to M-tubulin. *P < 0.001 compared with the ratio for M-tubulin. We detected very little binding of RACK1 to M- or P-tubulin. B: dose-dependent binding of NHERF1 to tubulin. P-tubulin (0.5 µg) was immobilized on PVDF paper then overlaid with the indicated amount of recombinant NHERF1 in 50 µl total volume. Bound protein was detected by immunoblot analysis for NHERF1 and exposed bands quantitated by optical density (OD) scan using a VersaDoc Imaging System. Immunoblots were reprobed for tubulin and quantitated by densitometry. The NHERF1 densitometric value was normalized to the tubulin signal and the normalized ratio used to determine binding constants. Binding constants, EC50, were calculated from %maximal binding as 2.8 µg, or a nominal 0.03 nmol. Results are representative of three independent experiments, each performed in triplicate or quadruplicate.

 
Binding of PDZ domains of NHERF1 to tubulin. NHERF1 consists of two PDZ domains which have been shown to mediate binding to other cellular proteins (11, 12). To determine whether a PDZ domain binds directly to tubulin, we performed solid phase binding assays using recombinant GST-tagged PDZ1 or PDZ2 domains of human NHERF1. The PDZ1, but not PDZ2, domain (Fig. 4A) binds to tubulin in a dose-dependent manner (Fig. 4B). The EC50, or effective concentration at which 50% binding is observed, was calculated as 1.5 µg, or a nominal 0.03 nmol.


Figure 4
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  		Fig. 4. Direct binding of PDZ domains to tubulin. A: binding of GST-tagged PDZ1 and GST-tagged PDZ2 domains was determined using a solid phase slot blot assay. M-tubulin (0.5 µg) and P-tubulin were immobilized on PVDF paper, overlaid with solutions containing 2 µg PDZ1 or PDZ2 domain, and incubated at room temperature for 25 min. Unbound material was removed by washing and bound protein detected by immunoblot analysis for the GST-tag. Membrane paper was reprobed for tubulin. Exposed bands were quantitated by densitometry and data were normalized as the ratio of (GST signal)/(tubulin signal) for each slot. Data are reported as means ± SE for the number of replicates. PDZ1 domain, but not PDZ2 domain, binds to M- and P-tubulin. B: dose-dependent binding of the PDZ1 domain to 0.5 µg immobilized P-tublin was performed as described in A. P-tubulin was overlaid with the indicated amount of recombinant GST-tagged PDZ1 in 50 µl total volume. Bound protein was detected by immunoblot analysis for the GST tag and exposed bands quantitated by optical density (OD) scan using a VersaDoc Imaging System. Immunoblots were reprobed for tubulin and quantitated by densitometry. The GST densitometric value was normalized to the tubulin signal and the normalized ratio used to determine binding constants. Binding constants, EC50s, were calculated from %maximal binding as 1.5 µg, or a nominal 0.03 nmole. Results are representative of three independent experiments, each performed in triplicate or quadruplicate.

 
Binding of endogenous tubulin to NHERF1 in Calu-3 cells. To determine whether tubulin interacts with endogenous NHERF1, we performed coimmunoprecipitation and pulldown experiments using total cell lysates prepared from Calu-3 cells. Typical results are shown in Fig. 5. Tubulin was detected in immunoprecipitates of NHERF1 (Fig. 5A), CFTR (Fig. 5B), and RACK1 (Fig. 5C). We also probed immunoprecipitates of NHERF1 for syntaxin1A, a protein shown to bind specifically to the N-terminus of CFTR (16). As expected, we did not detect syntaxin1A. We probed pulldowns of tubulin for RACK1, NHERF1, PKC{varepsilon}, and CFTR and observed protein bands corresponding to NHERF1, CFTR, and PKC{varepsilon} (Fig. 5D). We did not detect a protein band corresponding to RACK1. One explanation for this is that RACK1 only indirectly interacts with tubulin via NHERF1. These results indicate an interaction between endogenous tubulin and NHERF1.


Figure 5
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  		Fig. 5. Interaction of endogenous proteins with tubulin. Calu-3 cells were grown to confluence, serum-deprived overnight, rapidly washed in ice-cold PBS, and solubilized in lysis buffer. NHERF1 (A), cystic fibrosis transmembrane regulator (CFTR) (B), or RACK1 (C) were immunoprecipitated from 1 ml aliquots of TCL. Tubulin (D) was pulled down using anti-tubulin antibody coupled to agarose beads, as described in MATERIALS AND METHODS. Proteins recovered in immunoprecipitates (IPs) were probed by immunoblot analysis (IB) using antibodies directed against the protein of interest. Each protein was detected in TCL. A: tubulin, but not syntaxin 1A, was detected in IPs of NHERF1. B and C: tubulin was detected in IPs of CFTR and RACK1. D: NHERF1, PKC-{varepsilon}, and CFTR were detected in pulldowns of tubulin. RACK1 was not detected. Data are representative of two experiments in which each condition was replicated at least three times.

 
Evidence for involvement of the PDZ1 domain in the binding of NHERF1 to tubulin. To determine whether the PDZ1 domain of NHERF1 interacts with endogenous tubulin, we used recombinant GST-PDZ1 domain as a competitive inhibitor of coimmunoprecipitation of endogenous NHERF1 and tubulin from Calu-3 total cell lysates. Total cell lysates was preincubated with 50 µg GST-PDZ1, followed by immunoprecipitation of NHERF1. Proteins recovered in immunoprecipitates were probed for tubulin and reprobed for NHERF1. As a control, separate aliquots of total cell lysates were preincubated with 50 µg GST-PDZ2 domain or with GST alone. The results, shown in Fig. 6, demonstrate that the PDZ1 domain, but not the PDZ2 domain, blunts coimmunoprecipitation of tubulin with NHERF1.


Figure 6
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  		Fig. 6. Inhibition of coimmunoprecipitation of tubulin with NHERF1 by the PDZ1 domain. One milliliter (2.8 mg protein) total cell lysate was prepared from Calu-3 cells and preincubated with 50 µg GST-PDZ1 domain for 20 min at 30°C immediately before addition of anti-EBP50 antibody. In one control, GST-PDZ2 was substituted for GST-PDZ1. In a second control, pulldown with GST alone was performed. Immune complexes were recovered and subjected to gel electrophoresis on 4–15% gradient slab gels and probed for tubulin then were reprobed for NHERF1. Exposed bands were quantitated by densitometry and the value for tubulin normalized to NHERF1. GST alone did not immunoreact with antibody to tubulin or NHERF1. In the absence of added PDZ domain or after preincubation with PDZ2 domain, the ratio was 0.14 and 0.16, respectively. In the presence of PDZ1 domain, the ratio decreased to 0.06, indicating a 57% inhibition of tubulin pulldown with NHERF1. The results are representative of two experiments in which each condition was replicated twice.

 
Evidence for involvement of the PDZ2 domain in the binding to NHERF1. Previously, we (20) reported binding of RACK1 with NHERF1 at the PDZ1 domain of NHERF1. One model for the interaction of a PKC{varepsilon}-RACK1-NHERF1 protein complex with tubulin involves, minimally, the binding of two NHERF1 molecules at the PDZ2 domain. Recombinant PDZ1 and PDZ2 domains have been reported to bind directly to NHERF1 (9, 12). We examined direct binding of recombinant PDZ domains and NHERF1 in solid phase binding studies. We cleaved off the GST-tag from NHERF1 using thrombin. The GST-cleaved NHERF1 was immobilized on PVDF paper then overlaid with solutions containing varying amounts of a GST-PDZ domain. As expected, PDZ1 and PDZ2 domains each bind to NHERF1 (Fig. 7A). To determine whether PDZ2 domains interact, we performed competitive binding experiments using PDZ2 domain from which the GST tag was cleaved off to inhibit binding of the GST-PDZ2 domain. As a control, we tested a PDZ1 domain from which the GST tag was cleaved. Direct binding of GST-PDZ2 domain to NHERF1, calculated as a ratio of GST/NHERF1 based on densitometric values, was 1.45 ± 0.3 (n = 4). The addition of PDZ1 domain reduced the binding of the PDZ2 domain by 7.6% to 1.34 ± 0.09 (n = 4), as shown in Fig. 7B. This ratio was not significantly different than direct binding. PDZ2 domain, however, reduced binding of GST-PDZ2 to NHERF1 by 48.9% to 0.74 ± 0.02 (n = 4), a ratio significantly different than direct binding (P < 0.01). These results indicate direct binding of PDZ2 domains of NHERF1. We next asked whether recombinant GST-PDZ2 domain interacts with endogenous NHERF1. Pulldown assays were performed by adding 50 µg GST-PDZ2 domain to 1 ml Calu-3 total cell lysate and probing recovered proteins for NHERF1. As seen in Fig. 7C, NHERF1 was detected in pulldowns, indicating an interaction between GST-PDZ2 domain and endogenous NHERF1. These results provide support for direct interaction of PDZ2 domains, as depicted in our model of interacting NHERF1 molecules.


Figure 7
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  		Fig. 7. Interaction of PDZ domains with NHERF1. A: binding of GST-tagged PDZ1 and GST-tagged PDZ2 domains to NHERF1 was determined using an in vitro solid phase slot blot assay. GST-NHERF1 was incubated overnight with thrombin at 30°C with constant rotation to cleave off the GST tag. NHERF1 was recovered and immobilized on PVDF paper, overlaid with solutions containing the indicated amounts of GST-PDZ1 or GST-PDZ2 domain, and incubated at RT for 25 min. Unbound material was removed by washing and bound PDZ domain detected by immunoblot analysis for the GST-tag. Membrane paper was reprobed for NHERF1. PDZ1 and PDZ2 domains each bind to NHERF1. B: to determine specificity of binding of PDZ2 domain, competitive inhibition of binding was tested using PDZ2 domain from which the GST tag was cleaved off using PreScission enzyme (Clontech). Six micrograms of PDZ2 domain was mixed with 3 µg GST-tagged PDZ2 domain immediately before application to immobilized NHERF1. PDZ1 domain from which a GST tag was cleaved using PreScission enzyme served as a control. Bound PDZ2 domain was detected by immunoblot analysis for the GST tag. Exposed bands were quantitated by densitometry and data for the PDZ2 domain were normalized to NHERF1. Cleaved PDZ2 domain reduced normalized GST-PDZ2 domain binding from 1.45 ± 0.03 (n = 4) to 0.74 ± 0.02 (n = 4), a loss of 48.9% bound GST-PDZ2 domain. Cleaved PDZ1 domain did not affect binding of GST-PDZ2 domain to NHERF1. C: GST-tagged PDZ2 domain (50 µg) was added to 1 ml TCL in pulldown assays. Proteins bound to the PDZ2 domain were recovered using anti-GST antibody coupled to agarose beads and probed by immunoblot analysis for NHERF1. Endogenous NHERF1 was detected as a 50-kDa protein band. Results represent three independent experiments.

 
Functional role of a RACK1 proteome. The results above indicate the expression of a proteome involving two scaffold proteins, RACK1 and NHERF1 and PKC-{varepsilon} and tubulin; we termed the proteome a RACK1 proteome to differentiate it from NHERF1-related proteomes. To ascertain the functional role of the RACK1 proteome, we downregulated RACK1 using double stranded siRNA delivered into cells by electroporation. We first performed experiments to optimize the amount of siRACK1 delivered into cells and the time after electroporation to achieve maximal loss of RACK1 protein. As a control for delivery of nucleic acid into the cell nucleus, we also delivered a reporter vector pmaxGFP which encoded the full length cDNA for GFP. GFP expressing cells were detected by fluorescence microscopy after 24 (Fig. 8A), 48, and 72 h incubation following electroporation. Maximal loss of RACK1 was detected at 48 h after electroporation with 1.5 pmol siRACK1 per 1 x 106 cells (Fig. 8B). Expression of CFTR (Fig. 8B) was not affected by electroporation alone or by treatment with siRACK1.


Figure 8
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  		Fig. 8. Functional consequences of downregulation of RACK1. A: expression of GFP 24 h after delivery of pmaxGFP into Calu-3 cells by electroporation. GFP is detected in multiple cells in this micrograph as white fluorescent signal (white arrowhead). B: downregulation of endogenous RACK1. Small interfering (si)RNA directed against RACK1 (siRACK1) was delivered into Calu-3 cells by electroporation as described in MATERIALS AND METHODS. Cells were incubated for 48 h after electroporation with 1.5 pmol siRACK1 per 6 x 106 cells and TCL prepared. As controls, equal aliquots of cells were not electroporated, or electroporated without siRACK1. Immunoblot analysis for RACK1 was performed on a 15-µg aliquot of TCL. Maximal loss of RACK1 protein was 77.5% after 48-h incubation immediately following electroporation. Blots were stripped and reprobed for CFTR using a monoclonal antibody directed against the COOH-terminus of CFTR. Densitometry of exposed protein bands showed no loss of CFTR after treatment with siRACK1. Typical results from 5 experiments are illustrated. C: surface expression of CFTR in Calu-3 cells. Calu-3 cells electroporated without or with 1.5 pmol siRACK1 per 6 x 106 cells were incubated for 48 h following electroporation. Cells were biotinylated on the apical surface, as described in MATERIALS AND METHODS. As a control for detection of CFTR, immunoblot analysis for CFTR was performed on a 60 µg aliquot of TCL followed by densitometry. The ratio of biotinylated CFTR (lane 1) to CFTR detected in TCL (lane 4) was 0.39 ± .01 (n = 4). Electroporation of Calu-3 cells in the absence of siRACK1 did not alter this ratio (lane 2). Detection of biotinylated CFTR was reduced by 87.4% after delivery of siRACK1 by electroporation (lane 3). Typical results from 3 experiments are shown. D: nontargeting control RNA. Calu-3 cells electroporated without (Veh) or with 1.5 pmole nontargeting siCONTROL RNA (siCTRL), or with siRACK1 per 1 x 106 cells were incubated for 48 h following electroporation and TCL was prepared. Immunoblot analysis for proteins was performed on 50 µg TCL for CFTR immunoblot and on 15 µg TCL for the remaining immunoblots. Data are calculated as a ratio of laser densitometry values for the protein of interest divided by actin. Typical results from 3 experiments are illustrated. Electroporation of siCONTROL RNA did not affect the protein expression of CFTR, RACK1, NHERF1, and actin. E: surface expression of RACK1. Immunoblots of biotinylated samples and supernatants from streptavidin pulldowns from D were reprobed for RACK1. RACK1 was detected in biotinylated samples and supernatants, indicating the expression of RACK1 in the apical plasma membrane of Calu-3 cells.

 
One consequence of loss of RACK1 is likely to be the loss of an intact RACK1 proteome. The highly localized expression of RACK1 at the apical region of Calu-3 cells (Fig. 1), led us to hypothesize that the RACK1 proteome may function to stabilize CFTR surface expression. We tested this possibility by assessing apical expression of biotinylated CFTR and RACK1 in cells electroporated with or, as controls, without siRACK1, or with nontargeting siCONTROL RNA. As an estimate of the amount of biotinylated CFTR, we calculated a ratio of biotinylated CFTR to CFTR detected in Calu-3 total cell lysate from densitometric values. Before electroporation, this ratio was 0.39 ± 0.01 (n = 4). As seen in Fig. 8C, after electroporation alone, biotinylated CFTR expression relative to total cell lysate CFTR was similar at 0.31, indicating that electroporation alone did not alter CFTR expression. Downregulation of RACK1 led to an 87.4% loss of apical biotinylated CFTR. Immunoblot analysis of cell lysates derived from biotinylated Calu-3 cells showed a 97% loss of RACK1 but no effect of siRACK1 treatment on CFTR, NHERF1, or actin expression (Fig. 8D). As a control for the specificity of the siRACK1, cells were electroporated with a like amount of nontargeting siCONTROL RNA. Treatment with non-targeting siCONTROL RNA did not affect the amount of biotinylated CFTR or the amount of CFTR, RACK1, NHERF1, and actin in supernatants from the biotinylation procedure, indicating selective downregulation of RACK1 by siRACK1 (Fig. 8D). Immunoblots of biotinylated samples and supernatants recovered from the streptavidin pulldown were reprobed for RACK1, as shown in Fig. 8E. RACK1 was detected in biotinylated samples and supernatants, indicating expression of RACK1 in the apical plasma membrane of Calu-3 cells. These results confirm and corroborate the immunofluorescence studies of Fig. 1. Overall, these results indicate a role for RACK1 in the surface expression of CFTR.


   DISCUSSION
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RACK1 regulates various biological and physiological functions as a scaffold/anchoring protein. Within this context, RACK1 is thought to promote efficient intracellular signaling through accurate and highly reproducible spatial and temporal compartmentalization of binding partners. We found previously that RACK1 is expressed predominantly in the apical region of airway epithelial cells, colocalizes with CFTR, and binds activated PKC{varepsilon} and NHERF1 at distinct WD repeats and that NHERF1 binds RACK1 at the PDZ1 domain (20-22). NHERF1 is a member of a family of four related PDZ-domain proteins and is concentrated in the apical membrane of Calu-3 airway epithelial cells (8). To explain the apical concentration of RACK1, we examined, by immunofluorescence and confocal microscopy, the localization of RACK1 and actin, a scaffold protein linked to efficient cAMP-dependent activation of CFTR through direct binding (6, 31) or indirectly via an ezrin-NHERF1 complex. Actin was detected at the cell boundaries and in both apical and basolateral membranes and RACK1 almost exclusively at the apical plasma membrane (Fig. 1). Although RACK1 and actin were expressed in the apical region of Calul-3 cells, merged images of fluorescent tagged proteins revealed lack of colocalization, suggesting that RACK1 anchoring involves a different protein or protein complex. Expression of RACK1 in the apical plasma membrane was confirmed from immunoblot analysis of biotinylated apical membrane proteins (Fig. 8E). Mass spectrometric analysis of endogenous NHERF1, a binding partner of RACK1, revealed the presence of beta-tubulin, a monomer of microtubules. In an in vitro assay, polymerized tubulin was found to bind NHERF1 but not RACK1 (Fig. 3). More extensive binding studies demonstrated an in vitro interaction between NHERF1 and tubulin via a PDZ1 domain (Figs. 4 and 6), direct binding of a PDZ2 domain to NHERF1 (Fig. 7B), and pull down of endogenous NHERF1 by recombinant PDZ2 domain from Calu-3 total cell lysate (Fig. 7C).

The findings from binding, coimmunoprecipitation and pulldown studies point to a direct interaction between NHERF1 and tubulin. A hypothetical model to explain the experimental results is presented in Fig. 9. In this model, termed a RACK1 proteome, a minimum number of interacting proteins include two NHERF1 molecules forming a homodimer at the PDZ2 domains. One NHERF1 molecule binds to tubulin via a PDZ1 domain and the other NHERF1 molecule binds to the WD5 repeat of RACK1, thus indirectly linking RACK1 to tubulin, as shown in coimmunoprecipitation and pull-down assays of Fig. 5, C and D. Activated PKC{varepsilon} binds to the sixth WD repeat of RACK1. One feature of the RACK1 proteome is direct binding of two NHERF1 molecules via PDZ2 domains. The experimental data of Fig. 7 provides support for this interaction from in vitro binding assays which show inhibition of binding of PDZ2 to NHERF1 by recombinant PDZ2 domain and pull down of endogenous NHERF1 from Calu-3 total cell lysate by the PDZ2 domain. These results agree with published reports of self-association of EBP50 through PDZ-PDZ interactions (9).


Figure 9
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  		Fig. 9. Hypothetical model of an epithelial RACK1 proteome. RACK1 interacts with activated PKC{varepsilon} at the WD6 repeat and the PDZ1 domain of NHERF1 at the WD5 repeat. Additional proteins that may associate with other WD repeats remain to be identified. The PKC{varepsilon}-RACK1-NHERF1 complex binds to a second NHERF1 molecule via PDZ2 domains and the second NHERF1 molecule binds to tubulin via a PDZ1 domain. The scaffold protein tubulin may serve as an anchor to localize the RACK1 proteome to the apical region of epithelial cell.

 
One function of scaffold/anchor proteins is to localize proteins critical for a physiological function to a specific intracellular subdomain. The model in Fig. 9 depicts a role for a RACK1 proteome in stabilizing CFTR surface expression. RACK1 binds activated PKC{varepsilon}, which, when downregulated or inhibited, blunts cAMP-dependent activation of CFTR (17). Downregulation of RACK1, on the other hand, reduces the apical expression of biotinylated CFTR, indicating a role for a RACK1 proteome in regulating the residency of CFTR at the cell surface. How this is accomplished remains unknown. RACK1 is thought to potentially bind seven proteins, one on each WD repeat. Thus, airway epithelial RACK1 may bind proteins, in addition to PKC{varepsilon} and NHERF1, which contribute to the surface expression of CFTR. This exemplifies the scaffolding function of RACK1, which is important for assembling signaling complexes at or near their functional site. The physiological function of some intracellular signaling complexes has only partially been elucidated. One example is signaling in mammalian hippocampus, in which RACK1 plays a role in gene expression. RACK1 binds both the NR2B subunit of the N-methyl-D-aspartate (NMDA) receptor and the nonreceptor tyrosine kinase FYN and acts as an inhibitory scaffold by inhibiting Fyn phosphorylation of NR2B (39). Release of RACK1 from NR2B and Fyn is mediated by activation of the cAMP/PKA pathway with subsequent Fyn-mediated phosphorylation of NR2B and movement of RACK1 into the nuclear compartment. Nuclear RACK1 regulates expression of pituitary adenylate cyclase activating polypeptide [PACAP(1-38)], which activates cAMP/PKA (39). Another example of the physiological role of RACK1 signaling complexes involves a component of translating microsomes (5). RACK1 binds activated PKC, which leads to initiation of translation through phosphorylation of the initiation factor 6 (eIF6). Finally, cardiac phenotype and its related function are also regulated by association of activated PKC{varepsilon} and PKCbetaII to RACK proteins, which protect against ischemic injury and promote hypertrophic growth, respectively (25, 26).

The mechanism for the effects of downregulated RACK1 in airway epithelial cells remains unknown, but has considerable relevance to cystic fibrosis for providing a unique site of protein interaction for novel drugs designed and developed to correct or improve defective CFTR function in cystic fibrosis. Intensive studies have established cellular control of CFTR channel activity, of the total number of apical CFTR channels, and of stability of residency of CFTR channels in the apical membrane through multiprotein complexes which interact with N- or COOH-terminal amino acids of CFTR at discrete binding motifs, as reviewed in Refs. 4 and 11. In addition to protein interactions, modifications of CFTR associated with Cl channel activity, such as phosphorylation, involve several protein kinases, including AMPK, cAMP/PKA, and PKC. Control of CFTR surface expression and/or its stability by an epithelial RACK1 proteome may be more complicated than direct interaction with CFTR. First, although RACK1 colocalizes with CFTR in the apical plasma membrane, there is still a question of direct binding of the proteins (21). Second, the study reported here indicates an indirect interaction between RACK1 and tubulin through two NHERF1 molecules via each PDZ domain, leaving no site on NHERF1 for binding of CFTR. One mechanism for the effects of a RACK1 proteome is through microtubule-dependent regulation of CFTR surface expression. Microtubules regulate vesicular traffic in polarized epithelial cells, a process sensitive, in some cell types, to nocodazole, which blocks polymerization of tubulin monomers and thereby prevents vesicular transport. Pretreatment with nocodazole blocks a cAMP-dependent shift of CFTR from subapical to apical membrane and reduces fluid secretion in intact rat intestine (1), prevents cAMP-induced CFTR exocytosis and activation in normal airway epithelial cells (9HTEo–) (32), reduces cAMP-dependent CFTR current in oocytes expressing CFTR (38), and blocks forskolin-induced translocation of CFTR to the cell surface in T84 cells (36). The role of microtubules in CFTR trafficking in Calu-3 cells has yet to be evaluated but is necessary to understand the role of a RACK1 proteome. Finally, a model for an airway epithelial RACK1 proteome as a linear array of interacting proteins may be an oversimplification. Indeed, RACK1 may bind additional proteins which are not yet identified, but have a vital role in CFTR surface expression.

Implications of these findings to genetically altered CFTR in cystic fibrosis have yet to be determined. One possibility is that the interaction of {Delta}F508-CFTR, the most common mutant in cystic fibrosis, with a RACK1 proteome, may be compromised and thus account for a short residency time for {Delta}F508-CFTR in the apical membrane (23, 34). This type of interaction might explain how small molecule corrector drugs improve {Delta}F508-CFTR surface density might involve an interaction with a RACK1 proteome (37). In summary, the study reported here suggests a role for a RACK1 proteome in CFTR apical expression with implications for CFTR trafficking.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This research was supported by National Heart, Lung, and Blood Institute Grant HL-67190 and from a Cystic Fibrosis Foundation Research Development Program grant.


   ACKNOWLEDGMENTS
 
The authors thank Laura Smith for her technical expertise with silencing RNA. PDZ domain constructs were kindly provided by Dr. V. Raghuram (Univ. Pennsylvania) and NHERF1 construct by Dr. C. H. C. Yun (Emory University).


   FOOTNOTES
 

Address for reprint requests and other correspondence: C. M. Liedtke, Pediatric Pulmonology, Case Western Reserve Univ., BRB, Rm. 824, 2109 Adelbert Rd., Cleveland, OH 44106-4948 (e-mail: carole.liedtke{at}case.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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