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
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ABSTRACT |
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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
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INTRODUCTION |
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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
-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
-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.
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METHODS |
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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 -actin before amplification for CFTR. For amplification of
-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
-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
-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.
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RESULTS |
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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.
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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.
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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).
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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).
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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.
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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).
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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.
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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.
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DISCUSSION |
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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
-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.
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ACKNOWLEDGEMENTS |
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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.
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FOOTNOTES |
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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.
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