Characterization of CFTR expression and chloride channel activity in human endothelia

Albert Tousson1,2, Brian A. Van Tine3,4, Anjaparavanda P. Naren1,5, George M. Shaw4,6, and Lisa M. Schwiebert1,2,5

1 Gregory Fleming James Cystic Fibrosis Research Center, Departments of 2 Cell Biology, 3 Pathology, 4 Medicine, and 5 Physiology and Biophysics, and 6 Howard Hughes Medical Institute, University of Alabama at Birmingham, Birmingham, Alabama 35294

    ABSTRACT
Top
Abstract
Introduction
Methods
Results
Discussion
References

The cystic fibrosis transmembrane conductance regulator (CFTR) functions as a low-conductance, cAMP-regulated chloride (Cl-) channel in a variety of cell types, such as exocrine epithelial cells. Our results demonstrate that human primary endothelial cells isolated from umbilical vein (HUVEC) and lung microvasculature (HLMVEC) also express CFTR as determined via RT-PCR and immunohistochemical and immunoprecipitation analyses. Moreover, Cl- efflux and whole cell patch-clamp analyses reveal that HUVEC (n = 6 samples, P < 0.05) and HLMVEC (n = 5 samples, P < 0.05) display cyclic nucleotide-stimulated Cl- transport that is inhibited by the CFTR selective Cl- channel blocker glibenclamide but not by the blocker DIDS, indicative of CFTR Cl- channel activity. Taken together, these findings demonstrate that human endothelial cells derived from multiple organ systems express CFTR and that CFTR functions as a cyclic nucleotide-regulated Cl- channel in human endothelia.

cystic fibrosis; cystic fibrosis transmembrane conductance regulator

    INTRODUCTION
Top
Abstract
Introduction
Methods
Results
Discussion
References

THE CYSTIC FIBROSIS transmembrane conductance regulator (CFTR) gene encodes a 180-kDa glycosylated protein that functions as a low-conductance, cyclic nucleotide-regulated Cl- channel (reviewed in Ref. 23); mutations within this gene cause cystic fibrosis (20). Structurally, the CFTR protein is comprised of a domain of six transmembrane-spanning alpha -helices, a first nucleotide-binding domain (NBD-1) that binds ATP, a large regulatory domain that is rich in cAMP-dependent kinase and protein kinase C phosphorylation sites, a second domain of six transmembrane-spanning alpha -helices, and a second nucleotide-binding domain that binds ATP (23). CFTR is expressed in a variety of epithelial cells, including cells isolated from large and small airway, sweat duct, and kidney, as well as in other cell types such as lymphocytes and cardiac myocytes (23). In addition, recent studies have demonstrated that CFTR functions as a Cl- channel within intracellular compartments, such as the endoplasmic reticulum (18, 19).

Endothelial cells have been regarded traditionally as barrier cells that regulate vascular tone through expression of vasoactive substances, such as bradykinin, and vasoactive autacoids, including nitric oxide and prostacyclin. Multiple types of Cl- channels have been characterized in endothelia (1, 16); however, direct evidence that CFTR is expressed and functioning in this cell type has not been reported previously. The objective of this study was to examine CFTR expression and function in human endothelial cells. Results presented herein demonstrate that human endothelial cells isolated from umbilical vein and lung microvasculature express CFTR and that CFTR functions as a cyclic nucleotide-regulated Cl- channel in these cells.

    METHODS
Top
Abstract
Introduction
Methods
Results
Discussion
References

Cell culture. Human endothelial cells isolated from umbilical vein (HUVEC) and lung microvasculature (HLMVEC) were purchased from Clonetics (Walkersville, MD) and were cultured according to the manufacturer's instructions. Briefly, cells were grown in endothelial cell basal media (Clonetics) containing 10 ng/ml human epidermal growth factor, 1 mg/ml hydrocortisone, 50 mg/ml gentamicin, 50 µg/ml amphotericin B, 3 mg/ml bovine brain extract, 1% penicillin/streptomycin, and 5% FBS. Cells were grown on plastic tissue culture ware coated with 2% gelatin.

A variety of cell lines known to express CFTR were included in the analyses described herein as positive controls. These cell lines included the following: 16HBE14o-, a bronchial epithelial cell line [a gift from Dr. Dieter Gruenert, Stanford University, Palo Alto, CA (6)]; 9HTEo-, a human tracheal epithelial cell line [a gift of Dr. Dieter Gruenert (5)]; PANC-1, a human pancreatic epithelial cell line [American Type Culture Collection, Rockville, MD (3)]; and 3T3-WT-H7 fibroblasts (a gift of Dr. Michael Welsh, University of Iowa, Iowa City, IA). 16HBE14o-, 9HTEo-, and PANC-1 cells were cultured in MEM-D-valine (GIBCO-BRL/Life Technologies, Grand Island, NY) containing 5% FBS and 1% penicillin/streptomycin and were grown on plastic tissue culture ware coated with Vitrogen (Collagen, Palo Alto, CA). 3T3-WT-H7 fibroblasts were cultured in MEM-high glucose containing 10% FBS and 1% penicillin/streptomycin.

Analysis of CFTR mRNA expression. HUVEC and HLMVEC were assayed for CFTR mRNA expression via RT-PCR. Total RNA was isolated from HUVEC and HLMVEC cells with TRIzol (GIBCO-BRL) and was DNase treated with 1 unit of DNase (GIBCO-BRL) per microgram of total RNA to remove contaminating genomic DNA. One microgram of DNase-treated RNA was reverse transcribed in a reaction containing 200 units Moloney murine leukemia virus-RT (GIBCO-BRL), 0.5 µg oligo(dT) primer, 0.5 mM dNTPs, and 25 units RNasin (Promega, Madison, WI). To control for RNA degradation during DNase treatment and reverse transcription, HUVEC and HLMVEC cDNA products were amplified for the housekeeping gene beta -actin before amplification for CFTR. For amplification of beta -actin, 0.2 µg of HUVEC or HLMVEC cDNA were mixed with 1 unit Taq polymerase (Perkin-Elmer, Norwalk, CT), 200 mM dNTPs, and 20 pmol each of the PCR primers (GIBCO-BRL) 5'-TGA CGG GGT CAC CCA CAC TGT GCC CAT CTA-3' and 5'-CTA GAA GCA TTG CGG TGG ACG ATG GAG GG-3'. PCR reactions were cycled with the following parameters: initial melt at 95°C for 5 min, 30 cycles of 95°C for 30 s, 46°C for 1 min, 72°C for 1 min, and a final extension at 72°C for 10 min. The expected fragment size was 690 bp. After beta -actin amplification, HUVEC and HLMVEC cDNA samples were amplified for CFTR in a reaction containing 0.2 µg of cDNA, 1 unit Taq polymerase, 200 µM dNTPs, and 20 pmol each of the following PCR primers (GIBCO-BRL): 5'-GAG GAC ACT GCT CCT ACA C-3' and 5'-CAG ATT AGC CCC ATG AGG AG-3' (spanning the region between nucleotides 531 and 778). Reactions were cycled as follows: initial melt at 95°C for 5 min, 40 cycles of 95°C for 1 min, 58°C for 1 min, 72°C for 2 min, and a final extension at 72°C for 10 min. For some endothelial samples, two rounds of CFTR amplification were performed to visualize the CFTR cDNA product. The expected fragment size was 248 bp.

To ensure that the sequence of the cDNA products derived from CFTR RT-PCR analyses corresponded to the published CFTR sequence (20), the cDNA bands were purified, subcloned, and prepared for dideoxy DNA sequencing. Briefly, cDNA bands were excised from a 1% agarose gel and purified with the Qiaquick gel extraction kit (5'-3', Santa Clarita, CA). The cDNA products were then subcloned into the pGEM-T vector (Promega) and were used to transform JM109 high-competency cells (Promega). Transformed cells were grown on Luria broth-agar plates containing ampicillin (100 µg/ml), X-gal (80 µg/ml), and isopropyl beta -D-thiogalactopyranoside (0.5 mM). Positive colonies were obtained via blue-white selection. For sequence analysis, plasmid DNA was extracted from the bacteria using the Perfect Prep kit (5'-3'), denatured, precipitated, and then sequenced using the Sequenase version 2.0 kit (Amersham, Arlington Heights, IL) according to the manufacturer's protocol.

Analysis of CFTR protein expression. HUVEC and HLMVEC were analyzed for CFTR protein expression via immunohistochemical and immunoprecipitation analyses. For immunohistochemical analysis, HUVEC and HLMVEC were grown to ~80% confluency on glass coverslips coated with 2% gelatin, rinsed briefly in PBS, and then permeabilized and fixed using several different methods to maximize the specific immunoreactivity of each CFTR-specific antibody employed, including antibodies directed against the R-domain [monoclonal (9); Genzyme, Cambridge, MA], the COOH terminus [polyclonal, alpha-1468 (14); a generous gift from Dr. Jonathan A. Cohn, Duke University, Durham, NC], NBD-1 (polyclonal; provided by the Gregory Fleming James Cystic Fibrosis Research Center at the University of Alabama at Birmingham), or the first extracellular loop sequence [monoclonal, MATG 1031 (7); a kind gift of Dr. D. Escande, Hopital G & R Laennec, Nautes, France]. For R-domain immunolocalization, cells were fixed with 3:1 (vol/vol) absolute methanol (Optima Grade; Fisher Scientific, Pittsburgh, PA)-acetic acid solution for 30 min at -20°C. Samples were then rinsed in PBS and postfixed with 3% formaldehyde (Tousimis Research, Rockville, MD) in PBS for 15 min at room temperature. For COOH terminus and NBD-1 detection, cells were fixed with absolute methanol only and postfixed with formaldehyde. For extracellular loop immunostaining, samples were fixed with 3% formaldehyde in PBS for 45 min at room temperature. Samples were then rinsed in PBS, permeabilized with 0.5% Triton X-100 (Sigma Chemical, St. Louis, MO) in PBS for 2.5 min, and again rinsed with PBS. For negative controls, cells were fixed accordingly. After fixation, nonspecific binding sites on HUVEC and HLMVEC were blocked with PBS containing 1% bovine serum albumin (BSA) for 30 min at room temperature. Cells were then stained with the CFTR-specific antibodies listed above (each at 10 µg/ml diluted in PBS-1% BSA) or the appropriate isotype-matched control (mouse IgG or rabbit IgG; 10 µg/ml) for 30 min at 37°C and rinsed in PBS. Samples were next stained with rat anti-mouse or goat anti-rabbit fluorescein isothiocyanate (FITC)-conjugated secondary antibody (Boeringer Mannheim, Indianapolis, IN) for 30 min at 37°C. Signal enhancement of the monoclonal antibodies (R-domain and extracellular loop) was performed with goat anti-rat IgG-FITC for 30 min at 37°C. After staining for CFTR, HUVEC and HLMVEC samples were again rinsed with PBS and then counterstained with Hoechst 33258 (Sigma Chemical) for 3 min at 20 µg/ml in PBS to visualize nuclei. After final brief rinse in PBS, samples were mounted in a solution containing 9:1 glycerol-PBS and 0.1% paraphenylenediamine and then stored at -20°C until analyzed. Samples were analyzed via digital confocal microscopy. Specifically, samples were examined on an Olympus IX70 inverted epifluorescence microscope equipped with step motor, filter wheel assembly (Ludl Electronics Products, Hawthorne, NY), and filter set 83000 (Omega Optical, Brattleboro, VT). Images were captured with a SenSys cooled charge-coupled device, high-resolution, monochromatic, digital camera (Photometrics, Tucson, AZ). Partial deconvolution of images was done with a PowerMac 9500/132 computer supplied with IP Lab Spectrum software (Scanalytics, Fairfax, VA) and PowerMac Microtome software (VayTek, Fairfield, IA).

For immunohistochemical analysis of CFTR protein expression in intact tissue, paraffin-embedded tissue samples derived from human lung microvasculature (obtained through the Tissue Procurement Facility at the University of Alabama at Birmingham) were heated at 57°C for 1 h. The samples were then deparaffinized with xylene three times and rehydrated with decreasing dilutions of ethanol (100, 100, 85, 70, and 50%) for 5 min each; this was followed by three rinses with distilled water for 2 min each. To block endogenous peroxidase activity, the tissue samples were incubated in 3% H2O2 for 15 min and then washed with distilled water three times at 2 min each. Next, samples were treated with a proteinase K solution (DAKO, Carpinteria, CA) for 15 min at 37°C and washed three times in distilled water for 2 min each, followed by three rinses in PBS for 2 min each. Samples were then blocked in 50% goat serum, diluted in 2× saline-sodium citrate (SSC) blocking buffer (1× SSC: 0.15 M NaCl, 0.015 M sodium citrate), for 1 h at 37°C. Once blocked, the samples were incubated with CFTR-specific antibodies, including the anti-NBD-1 or alpha-1486 (each at 100 µg/ml) antibodies described above, or the appropriate isotype-matched control (rabbit IgG, Sigma Chemical; rabbit Ig fraction, DAKO), diluted in 2× SSC blocking buffer, for 50 min at 37°C. Samples were washed in PBS for three times at 2 min each and then examined using the DAKO LSAB2 detection kit (DAKO). In brief, the tissue samples were incubated with a biotinylated anti-rabbit antibody for 20 min at 37°C, washed in PBS for three times at 2 min each, and then incubated in a streptavidin-horseradish peroxidase solution for 20 min at 37°C. Samples were again washed in PBS for three times at 2 min each and then incubated with fluorescein-tyramide (1:75 dilution in the provided amplification diluent; TSA-Direct kit; NEN Life Science Products, Boston, MA) for 10 min. Last, samples were washed in PBS, as described above, and then mounted in a 4,6-diamino-2-phenylindole antifade solution (Oncor, Gaithersburg, MD). Samples were analyzed via digital confocal epifluorescence microscopy as detailed above.

Immunoprecipitation analysis was performed as described previously (15). Briefly, HLMVEC were grown to confluence on 100-mm plastic petri dishes coated with 2% gelatin and then lysed in a solution containing 1.0% Triton X-100 in PBS (pH 7.4). HT-29CL19A cells, an intestinal epithelial cell line that expresses CFTR (15), and COS-7 cells that had been transfected transiently with CFTR were included in these experiments as positive controls; COS-7 mock controls and HLMVEC immunoprecipitated with an isotype-matched immunoglobulin (IgG) were included as negative controls. Immunoprecipitation was performed by incubating cell lysates with protein A/G beads (Santa Cruz Biotechnology, Santa Cruz, CA) to which anti-CFTR IgG (anti-COOH terminus; a generous gift from the Gregory Fleming James Cystic Fibrosis Research Center, University of Alabama at Birmingham) or nonimmune IgG was covalently bound. Bound proteins were then eluted and subjected to Western blot analysis and chemiluminescence detection (Amersham).

Measurement of CFTR Cl- channel function. To examine the Cl- channel function of CFTR in HUVEC and HLMVEC, Cl- efflux assays and patch-clamp analyses were performed as described previously (21, 24) with some modifications described below. For Cl- efflux analysis, cells were grown to 80-90% confluence on gelatin-coated plastic six-well dishes, washed with phospho-buffered saline, and then loaded with 36Cl- (5 µCi/dish; NEN). The Ringer solution for these experiments was a standard HCO-3-free, HEPES- and phosphate-buffered 140 mM NaCl Ringer supplemented with 5 mM glucose and titrated to pH 7.45 with 1 N NaOH. All efflux runs were paired, with each well serving as its own control; all runs were performed on a slide warmer at 37°C. At time 0, fresh Ringer solution was added and then removed at 15-s intervals to assess the rate of efflux of 36Cl- from the cells over time. At time "1 min," Ringer solution containing a cocktail of agonists including permeable cAMP analogs [8-bromo-cAMP, 8-(4-chlorophenylthio) (CPT)-cAMP each at 250 µM; Sigma Chemical], cGMP analogs (8-bromo-cGMP, CPT-cGMP each at 250 µM; Sigma Chemical), or all four cyclic nucleotide agonists together (cAMP/cGMP; 8-bromo-cAMP, CPT-cAMP each at 250 µM plus 8-bromo-cGMP, CPT-cGMP each at 250 µM) was added and removed at 15-s intervals to assess the effect of cyclic nucleotides, which stimulate CFTR, on 36Cl- efflux rate. This rate (calculated as min-1) reflects the incremental loss/efflux of 36Cl- over time from interval to interval (each interval or time point reflects 15 s of time in which 36Cl- efflux was monitored). This rate is an absolute rate and is not a rate coefficient. Pharmacological characterization of the Cl- channels that facilitate cyclic nucleotide-stimulated 36Cl- efflux was assessed in parallel experiments; these experiments utilized all four cyclic nucleotide agonists together with the Cl- channel blockers DIDS (200 µM; Sigma Chemical), glibenclamide (100 µM; Sigma Chemical), diphenylamine carboxylic acid (DPC, 500 µM; Fluka, Heidelberg, Germany), and 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB, 10 µM; Calbiochem, La Jolla, CA). After each efflux assay, cells were lysed in 0.5 N NaOH, and lysates were monitored for 36Cl- content via scintillation counting.

CFTR function in HLMVEC was analyzed via whole cell patch-clamp recording. Patch-clamp electrodes (VWR borosilicate capillary glass) were pulled with a Narishige microelectrode puller (PP-83) to a tip resistance of 5 MOmega . Patch pipettes were manipulated with a Burleigh/Newport motor-driven macro- and micromanipulator system mounted on an inverted Nikon Eclipse TE-200 microscope equipped with epifluorescence. Patch pipettes were mounted in an Axon Instruments pipette holder attached to an Axopatch CV 203BU head stage amplifier interfaced with an Axopatch 200B integrated patch-clamp amplifier, an Axon Instruments DigiDate 1200 series interface, an Axon Instruments mother board, and PClamp 6.0 software loaded on a Gateway Pentium (IBM-PC) model P5-166 computer. In the whole cell recordings, the bath solution (extracellular solution) contained (in mM) 110 N-methyl-D-glucamine chloride, 1 CaCl2, 1 MgCl2, 5 glucose, 100 sucrose, and 5 HEPES (pH 7.45). The pipette solution (intracellular solution) contained (in mM) 145 Tris · Cl, 5 HEPES, 2 EGTA to chelate Ca2+ and Mg2+ to ~100 nM, 1 MgGTP, and 5 MgATP. Sucrose was included in the bath solution to eliminate swelling-activated Cl- currents. Cl- is the only major permeant ion in these solutions; thus, only Cl- currents are measured in these recordings. Asymmetry in the Cl- between pipette and bath solutions also allows for a slight but significant shift in reversal potential (+5 to 10 mV, +7 mM predicted by the Nernst equation), which helps distinguish between leak and CFTR-dependent current. As described for efflux assays above, similar cocktails of permeable cAMP and cGMP analogs or all four cyclic nucleotide analogs together were used to stimulate CFTR whole cell Cl- currents; addition of the blockers DIDS (100 µM) followed by glibenclamide (100 µM) was utilized to characterize CFTR whole cell Cl- currents pharmacologically.

    RESULTS
Top
Abstract
Introduction
Methods
Results
Discussion
References

Analysis of CFTR mRNA expression in human endothelial cells. CFTR expression has been observed in a variety of human cell types, including epithelial cells, lymphocytes, and cardiac myocytes (23); therefore, it was hypothesized that CFTR may also be expressed in human endothelial cells. To examine CFTR mRNA expression in human endothelia, total RNA isolated from HUVEC and HLMVEC was analyzed via RT-PCR with CFTR-specific primers. cDNA derived from the human bronchial epithelial cell line 16HBE14o- was included as a positive control, since CFTR expression and Cl- channel activity have been well characterized in this cell line (6, 11). As shown in Fig. 1A, HUVEC from five different donor samples (lanes 3-7) expressed the expected 248-bp CFTR fragment as did the positive control, 16HBE14o-. Likewise, Figure 1B demonstrates that HLMVEC from three independent donors expressed the expected 248-bp CFTR fragment. These findings suggest that human endothelial cells derived from multiple organ systems express CFTR mRNA.


View larger version (41K):
[in this window]
[in a new window]
 
Fig. 1.   Human umbilical vein endothelial cells (HUVEC) and human lung microvasculature endothelial cells (HLMVEC) express cystic fibrosis transmembrane conductance regulator (CFTR) mRNA as determined via RT-PCR. cDNA derived from 5 different HUVEC donors (A, lanes 3-7) and 3 independent HLMVEC donors (B, lanes 3-5) were amplified for CFTR with primers spanning the region between nucleotides 531 and 778 (248-bp fragment). cDNA derived from 16HBE14o- was amplified for CFTR as a positive control (A and B, lane 2); a reaction containing no template was included as a negative control (A and B, lane 1). Samples were also amplified for beta -actin (690-bp fragment) to control for RNA degradation during DNase treatment and reverse transcription. M represents lanes containing 100-bp markers.

To confirm that the cDNA products generated via RT-PCR represented CFTR amplification, all cDNA products were subjected to DNA sequencing analysis. CFTR fragments amplified from HUVEC, HLMVEC, and the positive control 16HBE14o- corresponded exactly to the published CFTR sequence [Fig. 2 (20)] and, therefore, support the conclusion that HUVEC and HLMVEC express CFTR mRNA.


View larger version (37K):
[in this window]
[in a new window]
 
Fig. 2.   DNA sequencing of cDNA products derived from HUVEC and HLMVEC confirm CFTR identity. cDNA products amplified from 5 different HUVEC and 3 independent HLMVEC donor samples were sequenced using the DNA dideoxy method. Resulting sequences were analyzed for homology with a published sequence of CFTR derived from human epithelium (hCFTR; see Ref. 20). A representative HUVEC and HLMVEC CFTR sequence (huENDO) is shown. 16HBE, positive control 16HBE14o-.

Detection of CFTR protein in human endothelial cells. CFTR protein is localized on the plasma membrane of CFTR-expressing cells; however, recent evidence indicates that CFTR protein may also be inserted into the membrane of intracellular organelles, including the endoplasmic reticulum (18, 19). To examine CFTR protein expression in human endothelia, HUVEC and HLMVEC were analyzed via immunohistochemistry. Specifically, endothelial cells were permeabilized and stained with antibodies directed against various domains of human CFTR, including the R domain, the COOH terminus, NBD-1, or the first extracellular loop sequence. As shown in Fig. 3, positive staining for CFTR expression was observed with each of these antibodies. Cells analyzed with antibodies directed against the R-domain or the first extracellular loop displayed a punctate staining pattern around the nucleus and along cellular processes (Fig. 3). In addition, the antibody against the first extracellular loop of CFTR stained the plasma membrane positively and exhibited intense perinuclear staining (Fig. 3). Endothelial cells analyzed with antibodies that recognized NBD-1 or the COOH terminus of CFTR yielded a diffuse staining pattern throughout the cytoplasm and also stained the plasma membrane (Fig. 3). HUVEC and HLMVEC incubated with the appropriate negative control antibodies displayed no background detection (Fig. 3).


View larger version (100K):
[in this window]
[in a new window]
 
Fig. 3.   HUVEC and HLMVEC express CFTR protein as determined via immunohistochemical analysis. HUVEC and HLMVEC were fixed, permeabilized, and then stained with anti-CFTR antibodies directed against the R-domain, COOH terminus, nucleotide binding domain (NBD)-1, the first extracellular loop, or the negative controls, mouse or rabbit IgG, followed by the appropriate secondary antibody as described in METHODS. Cells were also stained with Hoechst 33258 to visualize nuclei. Representative results of 2 different HUVEC donors and 3 independent HLMVEC are shown.

To determine if human endothelial cells in intact lung tissue also express the CFTR protein, tissue samples derived from human lung microvasculature were analyzed immunohistochemically with antibodies directed against the COOH terminus or NBD-1. As shown in Fig. 4, these antibodies positively stained the lining of a small blood vessel within the tissue sample; such a pattern of staining most likely reflects expression of the CFTR protein in endothelial cells that line the blood vessel. Tissue stained with appropriate negative control antibodies revealed no cross-reactivity with the vessel lining (Fig. 4).


View larger version (53K):
[in this window]
[in a new window]
 
Fig. 4.   CFTR protein is expressed in endothelial cells within human lung tissue. Human tissue samples derived from lung microvasculature were incubated with anti-CFTR antibodies directed against the COOH terminus, NBD-1, or with negative control rabbit IgG followed by amplification using a fluorescein-tyramide scheme as described in METHODS. Representative results of 3 different human lung tissue samples are shown.

The CFTR protein may exist in either the core glycosylated form (Fig. 5, band B) and/or the mature, fully glycosylated form (Fig. 5, band C). To determine if CFTR expressed in human endothelial cells was fully glycosylated into the mature form, immunoprecipitation analysis was performed utilizing an antibody directed against the COOH terminus; analyses of COS-7 cells that had been transfected with CFTR and HT-29CL19A cells, an intestinal epithelial cell line that expresses CFTR (15), were included as positive controls. As shown in Fig. 5, COS-7-CFTR expressed both the core glycosylated (band B) and mature (band C) forms of CFTR protein; HT-29CL19A cells expressed primarily the mature form. COS-7 mock controls and the nonimmune IgG negative control displayed no cross-reactivity. Interestingly, the CFTR protein detected in HLMVEC was observed primarily in the mature form of the protein (Fig. 5, band C), suggesting that CFTR is processed normally as a membrane glycoprotein in endothelia as has been observed in epithelia. These findings, together with the results presented above, demonstrate clearly that human endothelia express CFTR.


View larger version (41K):
[in this window]
[in a new window]
 
Fig. 5.   Immunoprecipitation analysis reveals that HLMVEC express the mature form of CFTR protein. HLMVEC, COS-7 cells transfected with CFTR, COS-7 mock controls, and HT-29CL19A cells were lysed in a solution containing 1.0% Triton X-100 in PBS (pH 7.4) and then immunoprecipitated with anti-CFTR antibody (directed against the COOH terminus) or nonimmune IgG. Bound proteins were then eluted and subjected to Western blot analysis. Representative results of 3 independent HLMVEC donor samples are shown. Band B, core glycosylated form of CFTR; band C, mature, fully glycosylated form of CFTR. rCFTR, recombinant CFTR.

Analysis of CFTR function in human endothelia. CFTR functions as a cyclic nucleotide-regulated Cl- channel in a variety of cell types, including airway epithelial cells; specifically, CFTR can be regulated by the cyclic nucleotides cAMP (reviewed in Ref. 23) and cGMP (4, 12, 26). To determine if CFTR expressed in human endothelial cells functions as a cyclic nucleotide-regulated Cl- channel, HUVEC and HLMVEC were analyzed for Cl- transport stimulated in the presence of cAMP or cGMP analogs via Cl- efflux and whole cell patch clamp analyses. 36Cl- efflux in HUVEC was compared with cyclic nucleotide-stimulated 36Cl- efflux in a panel of epithelial and heterologous cells known to express wild-type CFTR; these results are summarized in Table 1. 36Cl- efflux stimulated in HUVEC by cAMP or cGMP analogs was comparable to that stimulated in 9HTEo- cells, a human tracheal epithelial cell line, and PANC-1 cells, a human pancreatic epithelial cell line; both cell lines express low but detectable levels of CFTR mRNA and protein (E. M. Schwiebert, unpublished observations; Table 1). 16HBE14o- bronchial epithelial cells and 3T3-WT-H7 fibroblasts stably transfected with CFTR (a generous gift of Dr. Michael Welsh, University of Iowa), both of which express readily detectable CFTR (E. M. Schwiebert, unpublished observations), exhibited a greater amount of cAMP-stimulated 36Cl- efflux than HUVEC (Table 1). Interestingly, 16HBE14o- cells failed to respond to cGMP, whereas cGMP stimulated 36Cl- efflux in HUVEC significantly (Table 1).

                              
View this table:
[in this window]
[in a new window]
 
Table 1.   Results of 36Cl- efflux in HUVEC

Several reports suggest that cross-talk between cAMP- and cGMP-mediated signaling pathways activates CFTR (4, 12). To examine the effects of cAMP combined with cGMP on CFTR activation in human endothelial cells, Cl- efflux was measured in endothelial cells stimulated with a cocktail containing cAMP and/or cGMP analogs in the presence and absence of various Cl- channel blockers. As shown in Fig. 6, cAMP or cGMP alone each stimulated a modest increase in 36Cl- efflux from HUVEC; however, cAMP in combination with cGMP stimulated 36Cl- efflux further. Glibenclamide, a selective inhibitor of CFTR (24), abolished this response (Fig. 6D). DPC and NPPB, blockers that inhibit a broader range of Cl- channel subtypes, also blocked this response (Fig. 6D). In sharp contrast, DIDS, a Cl- channel blocker that does not affect CFTR when added extracellularly (24), failed to inhibit cAMP/cGMP stimulation of 36Cl- efflux in HUVEC (Fig. 6D). Taken together, these results suggest that cyclic nucleotides stimulate 36Cl- efflux in HUVEC via the activation of CFTR in the plasma membrane.


View larger version (21K):
[in this window]
[in a new window]
 
Fig. 6.   HUVEC express functional CFTR as suggested by Cl- efflux assays. All efflux runs were paired, with each sample serving as its own control. Ringer solution containing agonist was added to the cells and removed at 15-s intervals to assess loss of Cl- over time. Agonists included a cAMP cocktail (A), a cGMP cocktail (B), and a cocktail of cAMP and cGMP analogs (C) as described in METHODS. bullet , Agonists; , time controls. Arrows indicate addition of agonists. Results with the blockers glibenclamide (Glib), DIDS, diphenylamine carboxylic acid (DPC), and 5-nitro-2-(3-phenylpropyl-amino)-benzoic acid (NPPB) at 1 min postaddition of cAMP combined with cGMP agonists are summarized in D. After the efflux run, cells were lysed and monitored for 36Cl- content via scintillation counting. Results are reported as percent Cl- lost per minute (n = 6 independent HUVEC samples; * P < 0.05 of paired experiments).

CFTR function in HLMVEC was examined via whole cell patch-clamp analysis. This analysis permitted the recording of the entire population of CFTR channels on the plasma membrane of the HLMVEC and facilitated quantification of the relative number of CFTR channels in HLMVEC versus the epithelial positive control, 16HBE14o- cells. As reported in Fig. 7, cAMP together with cGMP stimulated a two- to threefold increase in Cl- current in HLMVEC; cAMP or cGMP alone stimulated an equivalent level of Cl- current (data not shown). In 16HBE14o- cells, cAMP in combination with cGMP induced a fourfold increase in the Cl- current over basal levels (Fig. 7). In both cell types, the current-voltage relationship of the cyclic nucleotide-stimulated current was linear and independent of time or voltage, indicative of CFTR Cl- currents (Fig. 7A). The cyclic nucleotide-stimulated Cl- current in both cell types was inhibited to basal levels or less-than-basal levels by the selective CFTR inhibitor glibenclamide but not by DIDS (Fig. 7B). These data suggest that the Cl- current observed was carried via CFTR and demonstrate further that HLMVEC express functional CFTR Cl- channels.


View larger version (23K):
[in this window]
[in a new window]
 
Fig. 7.   CFTR functions as a cyclic nucleotide-regulated Cl- channel in HLMVEC. HLMVEC and 16HBE14o- cells were analyzed via whole cell patch-clamp recording for CFTR-mediated Cl- current. HLMVEC were analyzed in the presence of cAMP analogs combined with cGMP analogs followed by the addition of the blocker DIDS first followed subsequently by glibenclamide. Results are reported as current (I)-voltage (V) plot (A) and degree of difference in Cl- current at -100 mV holding potential (B) (n = 5 independent HLMVEC samples; * P < 0.05 by Student's paired t-test). Basal Cl- current densities were 42 ± 10 pA/pF for 16HBE14o- cells and 10 ± 1 pA/pF for HLMVEC primary cells. Current densities for Cl- currents stimulated with cAMP analogs combined with cGMP analogs were 134 ± 62 pA/pF for 16HBE14o- cells and 35 ± 2 pA/pF for HLMVEC cells at the -100 mV holding potential [membrane capacitance for the 16HBE14o- cells was 18 ± 2 pF and for the HLMVEC cells was 20 ± 4 pF (n = 3 samples each)].

    DISCUSSION
Top
Abstract
Introduction
Methods
Results
Discussion
References

The findings presented herein demonstrate that primary human endothelial cells derived from umbilical vein and lung microvasculature express CFTR mRNA and protein. To our knowledge, this is the first molecular and biochemical analysis of CFTR mRNA and protein expression in human endothelial cells. Through DNA sequencing, CFTR PCR products amplified from human endothelial cells were shown to have 100% identity with the published cDNA sequence (20). The CFTR-specific PCR primers utilized in this analysis corresponded to exons 3 through 6 of the CFTR cDNA; therefore, all PCR products from human endothelial cell mRNA samples contained exon 5. This finding suggests that CFTR expressed in human endothelial cells exhibits greater similarity to CFTR expressed in human epithelia than that expressed in other cell types, such as cardiac myocytes. CFTR expressed in cardiac myocytes was shown to lack exon 5 and display modestly different Cl- channel biophysical characteristics from CFTR in human epithelial cells (27). CFTR protein expression was documented in primary human endothelial cells as well as in endothelia that line small blood vessels in human lung via immunohistochemical analysis with an extensive panel of CFTR antibodies that recognized independent domains of CFTR, including NBD-1, R-domain, first extracellular loop, and COOH terminus. Staining likely reveals both immature and mature forms of CFTR due to punctate localization in the endoplasmic reticulum adjacent to the nucleus and clear plasma membrane staining. Detection of the fully glycosylated form of CFTR in HLMVEC suggests that these cells can express the full-length CFTR protein.

Functional analyses revealed that human endothelial cells contain CFTR Cl- channel activity. Specifically, parallel 36Cl- efflux assays and whole cell patch-clamp recordings using cyclic nucleotide agonists and Cl- channel antagonists demonstrated CFTR Cl- channel activity in these cells. Both cAMP and cGMP stimulated 36Cl- efflux and whole cell Cl- currents that were consistent with CFTR. In both assays, DIDS failed to inhibit cyclic nucleotide-stimulated Cl- transport, whereas subsequent addition of glibenclamide abolished Cl- transport. Biophysical parameters such as a linear current-voltage relationship and whole cell currents independent of time or voltage were also consistent with CFTR Cl- channel activity in human endothelial cells. Moreover, these results were consistent with results from a panel of several independent CFTR-positive epithelial cell lines, suggesting that primary human endothelial cells express functional CFTR in amounts equivalent to many epithelial cell models of CFTR.

In the study presented herein, cAMP combined with cGMP analogs did not enhance CFTR whole cell currents in endothelial cells above the levels observed with either cyclic nucleotide alone; such results suggest that cAMP and cGMP share signaling pathways to activate CFTR in these cells. Interestingly, cGMP stimulated CFTR-dependent Cl- transport in endothelial cells in a manner similar to that observed in colonic epithelial cells. In colonic epithelial cells, cGMP appears to "cross-stimulate" cAMP-dependent protein kinase and, subsequently, activate CFTR (4, 25). In contrast, Vaandrager et al. (26) have reported that, in an intestinal epithelial cell line, cGMP stimulates CFTR via cGMP-dependent kinase II (cGK II). These authors demonstrated that neither cGK I nor cAMP-dependent kinase could substitute for cGK II in the cGMP-mediated activation of CFTR in these cells (26). Moreover, Kelley et al. (12) have demonstrated that cAMP- and cGMP-mediated signaling pathways synergize to induce Cl- secretion across the nasal epithelium of CFTR knockout mice. These studies do not rule out the possibility that cyclic nucleotides may regulate CFTR Cl- channels via more than one signaling pathway. It is possible that agonists that stimulate cAMP- and cGMP-mediated signaling pathways specifically, such as beta -adrenergic agonists for cAMP and nitric oxide for cGMP, may show different stimulatory effects on CFTR.

Recent studies have characterized multiple Cl- currents in endothelial cells, including Ca2+-activated and volume-activated Cl- channels, yet have not identified CFTR-like Cl- channel activity (1, 16); however, these studies were performed on bovine endothelial primary cultures. Nilius et al. (16) reported that bovine pulmonary artery endothelial cells contain Ca2+- and volume-activated Cl- channels but lack cAMP-activated CFTR-like channels and voltage-activated Cl- (CLC)-like channels (16). It is surprising that CLC channels are lacking from these cells, because these channels are expressed ubiquitously. Moreover, these physiological assays were not paralleled with molecular or biochemical analyses to verify that these cells lack CFTR or CLC channels. In contrast, Bonanno and Srinivas (1) demonstrated the presence of cAMP-activated anion transport in corneal endothelia as measured via the transport of the halide fluorophore, 6-methoxy-N-(3-sulfopropyl)-quinolinium. These cAMP-activated transport pathways were permeable to HCO-3, inhibited by 50 µM NPPB, and insensitive to 100 µM DIDS. Although such results are consistent with CFTR channel activity, cAMP-activated Cl- transport was insensitive to 200 µM DPC and 50 µM glibenclamide; higher concentrations of DPC (>= 500 µM) and glibenclamide (>= 100 µM) are usually recommended and more efficacious in the inhibition of CFTR activity. Molecular and biochemical analyses to detect CFTR expression were not performed in this study. In both of the aforementioned studies, endothelial cells isolated from other tissues or preparations were not examined (1, 16).

The role of Cl- channels, such as CFTR, in endothelial cell biology is not well understood; however, several possibilities exist. First, CFTR may be involved in transendothelial ion transport. Although endothelia form a barrier that has low and variable resistance, vectorial ion and water transport may occur. Previous studies suggest that Cl- channels in endothelial cells may play a role in pH regulation (10). Second, because cGMP-dependent nitric oxide signaling is prevalent in vasodilation of blood vessels (8), cGMP-stimulated CFTR Cl- transport may be important in the maintenance of vascular tone. Third, CFTR-mediated Cl- transport may affect signaling pathways within endothelial cells. For example, CFTR-mediated Cl- transport may alter the endothelial cell's membrane potential. A change in the cell's membrane potential could affect the regulation of Ca2+ influx and, in turn, alter Ca2+-mediated signaling pathways that may modulate vascular permeability or elaboration of vasoactive mediators, such as nitric oxide (13, 17). Moreover, Cl- channels such as CFTR (19) and CLC-6 (2) appear to reside in the endoplasmic reticulum and, therefore, may regulate the release of Ca2+ from this intracellular store. Fourth, expression of CFTR is associated with ATP transport (22). ATP and its metabolites (especially adenosine) are potent vasoactive autocrine and paracrine substances that vasodilate or vasoconstrict, depending on the capillary bed of interest. These possible effects of CFTR on endothelial biology underscore the potential importance of CFTR expression and function in endothelial cells and, therefore, require further investigation.

    ACKNOWLEDGEMENTS

We acknowledge Drs. Dale J. Benos and Eric J. Sorscher for critical review of this manuscript and Drs. Marcio vaz Sanches, Erik M. Schwiebert, and Kevin L. Kirk and Yafen Niu for technical assistance.

    FOOTNOTES

This work was supported by the Gregory Fleming James Cystic Fibrosis Research Center at the University of Alabama at Birmingham and the American Heart Association (L. M. Schwiebert).

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. §1734 solely to indicate this fact.

Address for reprint requests: L. M. Schwiebert, Dept. of Physiology and Biophysics, McCallum Bldg., Rm. 966, Univ. of Alabama at Birmingham, 1918 University Blvd., Birmingham, AL 35294.

Received 23 June 1998; accepted in final form 20 August 1998.

    REFERENCES
Top
Abstract
Introduction
Methods
Results
Discussion
References

1.   Bonanno, J. A., and S. P. Srinivas. Cyclic AMP activates anion channels in cultured bovine corneal endothelial cells. Exp. Eye Res. 64: 953-962, 1997[Medline].

2.   Buyse, G., D. Trouet, T. Voets, L. Missiaen, G. Droogmans, B. Nilius, and J. Eggermont. Evidence for the intracellular location of chloride channel (ClC)-type proteins: co-localization of ClC-6a and ClC-6c with the sarco/endoplasmic reticulum Ca2+ pump SERCA2b. Biochem. J. 330: 1015-1021, 1998[Medline].

3.   Chang, B. K. Comparison of in vitro methods for assessing cytotoxic activity against two pancreatic adenocarcinoma cell lines. Cancer Res. 43: 3147-3149, 1983[Abstract].

4.   Chao, A. C., F. J. de Sauvage, Y. J. Dong, J. A. Wagner, D. V. Goeddel, and P. Gardner. Activation of intestinal CFTR Cl- channel by heat-stable enterotoxin and guanylin via cAMP-dependent protein kinase. EMBO J. 13: 1065-1072, 1994[Abstract].

5.   Cozens, A. L., M. J. Yezzi, L. Chin, E. M. Simon, D. S. Friend, and D. C. Gruenert. Chloride ion transport in transformed normal and cystic fibrosis epithelial cells. In: The Identification of the CF Gene, edited by L.-C. Tsui, G. Romeo, R. Greger, and S. Gorini. New York: Plenum, 1991, p. 187-196.

6.   Cozens, A. L., M. J. Yezzi, K. Kunzelmann, T. Ohrui, L. Chin, K. Eng, W. E. Finkbeiner, J. H. Widdicombe, and D. C. Gruenert. CFTR expression and chloride secretion in polarized immortal human bronchial epithelial cells. Am. J. Respir. Cell Mol. Biol. 10: 38-47, 1994[Abstract].

7.   Demolombe, S., I. Baro, Z. Bebok, J. P. Clancy, E. J. Sorscher, A. Thomas-Soumarmon, A. Pavirani, and D. Escande. A method for rapid detection of recombinant CFTR during gene therapy in cystic fibrosis. Gene Ther. 3: 685-694, 1996[Medline].

8.   Gaston, B., and J. S. Stamler. Nitrogen oxides. In: The Lung, edited by R. G. Crystal, J. B. West, E. R. Weibel, and P. J. Barnes. Philadelphia: Lippincott-Raven, 1997, p. 239-253.

9.   Gregory, R. J., S. H. Cheng, D. P. Rich, J. Marshall, S. Paul, K. Hehir, L. Ostedgaard, K. W. Klinger, M. J. Welsh, and A. E. Smith. Expression and characterization of the cystic fibrosis transmembrane conductance regulator. Nature 347: 382-386, 1990[Medline].

10.   Groschner, K., and W. R. Kukovetz. Voltage-sensitive chloride channels of large conductance in the membrane of pig aortic endothelial cells. Pflügers Arch. 421: 209-217, 1992[Medline].

11.   Haws, C., M. E. Krouse, Y. Xia, D. C. Gruenert, and J. J. Wine. CFTR channels in immortalized human airway cells. Am. J. Physiol. 263 (Lung Cell. Mol. Physiol. 7): L692-L707, 1992[Abstract/Free Full Text].

12.   Kelley, T. J., C. U. Cotton, and M. L. Drumm. In vivo activation of CFTR-dependent chloride transport in murine airway epithelium by CNP. Am. J. Physiol. 273 (Lung Cell. Mol. Physiol. 17): L1065-L1072, 1997[Abstract/Free Full Text].

13.   Lantoine, F., L. Iouzalen, M. A. Devynck, E. Millanvoye-Van Brussel, and M. David-Dufilho. Nitric oxide production in human endothelial cells stimulated by histamine requires Ca2+ influx. Biochem. J. 330: 695-699, 1998[Medline].

14.   Marino, C. R., L. M. Matovcik, F. S. Gorelick, and J. A. Cohn. Localization of the cystic fibrosis transmembrane conductance regulator in pancreas. J. Clin. Invest. 88: 712-716, 1991[Medline].

15.   Naren, A. P., D. J. Nelson, W. Xie, B. Jovov, J. Pevsner, M. K. Bennett, D. J. Benos, M. W. Quick, and K. L. Kirk. Regulation of CFTR chloride channels by syntaxin and Munc18 isoforms. Nature 390: 302-305, 1997[Medline].

16.   Nilius, B., G. Szucs, S. Heinke, T. Voets, and G. Droogmans. Multiple types of chloride channels in bovine pulmonary artery endothelial cells. J. Vasc. Res. 34: 220-228, 1997[Medline].

17.   Ono, K., M. Nakao, and T. Iijima. Chloride- and voltage-dependent Ca2+ transient in cultured human aortic endothelial cells. Heart Vessels 12: 50-52, 1997.

18.   Pasyk, E. A., and J. K. Foskett. Mutant (delta F508) cystic fibrosis transmembrane conductance regulator Cl- channel is functional when retained in endoplasmic reticulum of mammalian cells. J. Biol. Chem. 270: 12347-12350, 1995[Abstract/Free Full Text].

19.   Pasyk, E. A., and J. K. Foskett. Cystic fibrosis transmembrane conductance regulator-associated ATP and adenosine 3'-phosphate 5'-phosphosulfate channels in endoplasmic reticulum and plasma membranes. J. Biol. Chem. 272: 7746-7751, 1997[Abstract/Free Full Text].

20.   Riordan, J. R., J. M. Rommens, B. Kerem, N. Alon, R. Rozmahel, Z. Grzelczak, J. Zielenski, S. Lok, N. Plavsic, and J. L. Chou. Identification of the cystic fibrosis gene: cloning and characterization of complementary DNA. Science 245: 1066-1073, 1989[Medline].

21.   Schwiebert, E. M., L. P. Cid-Soto, M. Carter, D. Stafford, C. M. Murray, W. B. Guggino, and G. R. Cutting. Functional analysis of an alternative chloride channel (CLC-2) in cystic fibrosis airway epithelial cells. Proc. Natl. Acad. Sci. USA 95: 3879-3884, 1998[Abstract/Free Full Text].

22.   Schwiebert, E. M., M. E. Egan, T. H. Hwang, S. B. Fulmer, S. S. Allen, G. Cutting, and W. B. Guggino. CFTR regulates outwardly rectifying chloride channels through an autocrine mechanism involving ATP. Cell 81: 1063-1073, 1995[Medline].

23.   Schwiebert, E. M., and W. B. Guggino. Defective chloride and sodium transport in CF. In: The Lung, edited by R. G. Crystal, J. B. West, E. R. Weibel, and P. J. Barnes. Philadelphia, PA: Lippincott-Raven, 1997, p. 2555-2571.

24.   Schwiebert, E. M., M. M. Morales, S. Devidas, M. E. Egan, and W. B. Guggino. Chloride channel and chloride conductance regulator domains of CFTR. Proc. Natl. Acad. Sci. USA 95: 2674-2679, 1998[Abstract/Free Full Text].

25.   Tien, X. Y., T. A. Brasitus, M. A. Kaetzel, J. R. Dedman, and D. J. Nelson. Activation of the cystic fibrosis transmembrane conductance regulator by cGMP in the human colonic cancer cell line, Caco-2. J. Biol. Chem. 269: 51-54, 1994[Abstract/Free Full Text].

26.   Vaandrager, A. B., B. C. Tilly, A. Smolenski, S. Schneider-Rasp, A. M. Bot, M. Edixhoven, B. J. Scholte, T. Jarchau, U. Walter, S. M. Lohmann, W. C. Poller, and H. R. de Jonge. cGMP stimulation of cystic fibrosis transmembrane conductance regulator Cl- channels co-expressed with cGMP-dependent protein kinase type II but not type Ib. J. Biol. Chem. 272: 4195-4200, 1997[Abstract/Free Full Text].

27.   Xie, J., M. L. Drumm, J. Zhao, J. Ma, and P. B. Davis. Human epithelial cystic fibrosis transmembrane conductance regulator without exon 5 maintains partial chloride channel function in intracellular membranes. Biophys. J. 71: 3148-3156, 1997[Abstract].


Am J Physiol Cell Physiol 275(6):C1555-C1564
0002-9513/98 $5.00 Copyright © 1998 the American Physiological Society