1Institute for Environmental Medicine; and 2Institute for Medicine and Engineering, University of Pennsylvania, Medical Center, Philadelphia, Pennsylvania 19104; and 3Department of Cellular and Molecular Pharmacology, University of South Alabama, Mobile, Alabama 36688
Submitted 5 November 2002 ; accepted in final form 16 June 2003
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ABSTRACT |
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flow adaptation; KIR 6.2; sulfonylurea receptor; fluorescent glyburide; pulmonary microvascular endothelial cells
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MATERIALS AND METHODS |
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Exposure of cells to shear stress. Three systems for flow adaptation of cells were used because each has specific advantages. The first was an artificial capillary system that yields sufficient cells to obtain protein lysates and RNA for immunoblotting and mRNA dot blots. The use of this commercially available apparatus (CellMax Quad Artificial Capillary Cell Culture System, Cellco, Germantown, MD) for flow-adapting endothelial cells has been described previously (36, 37). The capillaries were precoated with fibronectin (Sigma, St. Louis, MO), and cells were allowed to attach during a 24-h period. Cells were then cultured under static conditions or under shear stress at 1 dyn/cm2 for 72 h or 10 dyn/cm2 for 24 h.
A parallel plate chamber was used to adapt cells to flow on coverslips. The chamber used to adapt cells for microscopy and whole cell patch-clamp studies consisted of a steel plate (7.5 cm long, 3.7 cm wide, 0.4 cm high) with a hollow slot in the center that can be fitted on top with a glass slide and on the bottom with a coverslip containing cells to create a rectangular flow channel (0.01 cm high, 2 cm wide, and 2 cm long). Inlet and outlet ports on the steel plate were connected to a pump and a reservoir to facilitate flow of medium into the system. RPMVEC were grown to confluence on either fibronectin- or gelatin-coated glass coverslips (2 x 2 cm) and then fitted into the chamber slot. With this apparatus, cells were subjected to shear stress for 72 h at 1 dyn/cm2 or 24 h at 5 or 10 dyn/cm2. The chamber used to flow adapt cells for measurements of membrane potential in real time with fluorescent dye was designed to fit in a standard fluorometer cuvette holder and has been described previously (24). Cells on plastic coverslips (1.25 x 2.5 cm) coated with fibronectin were grown under static conditions or flow adapted at 10 dyn/cm2 for 24 h.
Electrophysiological measurements of membrane potential. Cells grown on coverslips were either subjected to flow in the parallel plate chamber or kept under static conditions. Cells were removed with 0.025% trypsin and then replated on coverslips and placed on a petri dish on the modified stage of an inverted microscope. Ion currents were measured using the whole cell patch-clamp configuration (17). Cells were continuously perfused with an external bath solution that contained 156 mM KCl, 1.5 mM CaCl2, 1 mM MgCl2, 10 mM HEPES, and 1 mM EGTA, pH 7.3. The internal pipette solution contained 145 mM KCl, 1 mM MgCl2, 10 mM HEPES, and 1 mM EGTA, pH 7.3. The holding potential was -60 mV. Current was elicited by 500-ms linear voltage ramps from -160 to +60 mV at an interpulse interval of 5 s. Currents were recorded by using an EPC9 amplifier (HEKA Electronik, Lambrecht, Germany) and accompanying acquisition and analysis software (Pulse and Pulsefit, HEKA Electronik) running on a PowerCenter 150 (Mac OS) computer. For each cell, at least 100 voltage ramps were recorded and averaged by the computer. Cells that ruptured before 100 voltage ramps were excluded from the analysis. Whole cell capacitance and series resistance were monitored throughout each recording, and cell current values were normalized to the capacitance of the cell to obtain current density. Peak current density was defined as the most negative value during the voltage ramp. An experiment was defined as all of the measurements made on a particular day, and generally three to five cells were evaluated for each condition. For each experiment, measurements were made alternately from static cultured and flow-adapted cells, and the results for each condition were averaged. Data are expressed as means ± SE and were evaluated by analysis of variance with post hoc Tukey's test for paired comparisons using SigmaStat software (Jandel Scientific, San Rafael, CA). Differences between groups were considered statistically significant at P < 0.05.
Real time fluorescence measurements of membrane potential. RPMVEC grown on fibronectin coated slides under static conditions or adapted to flow of 10 dyn/cm2 for 24 h were preincubated with the membrane potential-sensitive dye bisoxonol (50 nM) for 30 min. In some experiments, the KATP channel agonist cromakalim (30 µM) was added during the preincubation period. In other experiments, cycloheximide (10 µg/mol) was added to the perfusate medium during the flow adaptation period. Detection of bisoxonol fluorescence was at 520 nm with 490 nm excitation. Fluorescence measurements for endothelial cell membrane potential measurements during flow and with stop of flow were performed with a PTI spectrofluorometer (Photon Technology International, Bricktown, NJ) equipped with single photon counting system with excitation and emission slits at 1 and 3 nm, respectively.
RT-PCR and RNA dot blots. Total RNA was isolated from static and flow-adapted RPMVEC by using an RNEasy kit (Qiagen, Los Angeles, CA). Total RNA was also isolated from blood-free rat lung homogenates. For RT-PCR, RNA (1 µg) from flow-adapted cells was reverse transcribed into cDNA and subjected to PCR by using a GeneAmp RNA-PCR core kit (Roche, Nutley, NJ) and GeneAmp 2400 (Perkin Elmer, Boston, MA). For inwardly rectifying K+ channels (KIR) 6.1, the forward and reverse primers were TGCGCAAACCGCGCATCCGCGACC and TCAGTGGCTGACTACGCTTATCAATCAC, respectively. For KIR 6.2, the forward and reverse primers were CTACAGAGCCCAGGTACCGTACTC and GGTGCAGGTCACTAGGAGCCTGTTGGAG. The PCR program consisted of initial denaturation at 95°C for 2 min followed by 35 cycles for 45 s, annealing at 60°C for 1 min with final extension step at 72°C for 10 min. The PCR products were separated by electrophoresis on a 8% SDS-polyacrylamide gel. The primers used in these reactions should yield a PCR product of 770 bp for KIR 6.1 and 780 bp for KIR 6.2.
For RNA dot blots, total RNA was absorbed on a nylon membrane by using a dot blot microfiltration apparatus (Bio-Rad) and probed with KIR 6.2 cDNA (entire coding sequence, a gift from F. Ashcroft, Oxford, UK) labeled with 32P. The blots were serially washed with 2x SSC-1% SDS and 0.2x SSC-0.1% SDS to a final temperature of 42°C for 30 min, followed by autoradiography with overnight exposure.
Immunoblotting. Flow-adapted and static cultured RPMVEC or BPAEC
were removed from the flow chamber by trypsinization. Trypsinized and pelleted
cells were suspended in ice-cold buffer consisting of 145 mM NaCl, 0.1 mM
MgCl2, 15 mM HEPES, 10 mM EGTA, 1 mM Na3VO4,
1% Triton X-100, and protease inhibitor cocktail (0.5 mM phenylmethylsulfonyl
fluoride, 50 mg/l leupeptin, 25 mg/l aprotinin, and 25 mg/l pepstatin, all
from Sigma). The cells were sonicated, and protein extracts were obtained by
denaturation in boiling sample buffer. Pancreatic -cells grown in static
culture and lung homogenate obtained from lungs perfused until blood free
(2) were used as a control.
Cell sonicate protein concentration was determined by Coomassie blue assay
(Bio-Rad) by using bovine IgG as the standard, and equal amounts of
experimental sample protein were loaded in the wells. After electrophoresis on
12% SDS-PAGE gel, proteins were transferred onto nitrocellulose membranes by
electroblotting. After being blocked with milk, the membranes were incubated
with polyclonal anti-KIR 6.2 peptide antibodies. An antibody to a
COOH terminus peptide, KAKPKFSISPDSLS (KIR 6.2-C Ab) was obtained
from S. Seino (Chiba University, Chiba, Japan). An antibody to a
NH2 terminus peptide, EEYVLTRLAEDPAEPRYRC (KIR 6.2-N
Ab), was obtained from A. Tinker (University College, London, UK).
Peroxidase-conjugated goat anti-rabbit IgG (Jackson ImmunoResearch Labs, West
Grove, PA) was used as the secondary antibody at 1:10,000 dilution. The signal
was detected by chemiluminescence by using an ECL kit (Amersham, Piscataway,
NJ). Reactivity of the antibodies was confirmed with pancreatic
-cell
extracts. The bands obtained were scanned using a Bio-Rad Fluor-S MultImager
equipped with Quant-I software for quantitation. Data are expressed as means
± SE and evaluated by t-test using SigmaStat software.
Expression of the SUR was evaluated by cellular binding of fluorescently labeled glyburide (BODIPY-FL glyburide). A 100 mM stock of BODIPY-FL glyburide (Molecular Probes, Eugene, OR) was prepared in DMSO. For RPMVEC, cells in the parallel plate chamber were incubated for 20 min with 50 nM BODIPY-FL glyburide, washed three times, fixed on the coverslips with 4% paraformaldehyde in 0.1 M Na-cacodylate buffer, and coverslipped in Mowiol. For BPAEC, cells were removed from attachment in artificial capillaries by trypsinization, placed in petri dishes, incubated with BODIPY-FL glyburide, and processed as for RPMVEC. Fluorescence images at 480 nm were obtained by using Meta-morph imaging software (Universal Imaging, West Chester, PA) and a Nikon Optiphot microscope (Melville, NY) (4). All images were processed by using a preset scale on the Meta-morph image analysis software to facilitate quantitative comparisons.
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RESULTS |
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The whole cell configuration of the patch-clamp technique was used to study the function of the KATP channel in RPMVEC that were grown under static conditions or flow adapted at 1, 5, or 10 dyn/cm2. A schematic of the voltage ramp that was used and typical patch-clamp currents is shown in Fig. 2. Cells cultured under static conditions showed a low-magnitude current that was appreciably greater in cells that were adapted to flow at 10 dyn/cm2 for 24 h. The currents show an inwardly rectifying current-voltage relationship typical for KIR. The reversal potential for both static and flow-adapted cells was approximately -2 mV, as expected under symmetrical K+ conditions. The addition of glyburide (50 nM) to the external bath solution significantly blocked the inward rectifier current, indicating a KATP channel. Peak current density for cells that were flow adapted at 1 dyn/cm2 was not different from cells cultured under static conditions, but flow adaptation at 5 dyn/cm2 resulted in a significant increase with a further increase at 10 dyn/cm2 (Table 1). These cells were grown and adapted to shear on a fibronectin matrix. Cells grown on gelatin-coated coverslips also showed a significant (P < 0.05) increase in channel activity with flow adaptation at 5 dyn/cm2 for 24 h compared with static cells.
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The inward rectifier subunit of KATP channels is a member of the KIR 6 family (15). Because there is high homology between the two currently known members, KIR 6.1 and KIR 6.2, RT-PCR was used to determine which of these inward rectifier subunits is present in RPMVEC. RT-PCR showed the 780-bp band predicted for KIR 6.2 but not the predicted band (770 bp) for KIR 6.1 (Fig. 3).
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The presence of KIR 6.2 mRNA in RPMVEC was detected by RNA dot
blot (Fig. 4A). The
KIR message was present in static cells and was increased
2-fold in cells adapted to shear stress at 1 dyn/cm2 and
slightly more at 10 dyn/cm2. GAPDH mRNA used as a standard for
loading efficiency showed similar density for all three conditions. RNA
extracted from the whole lung demonstrated a signal intermediate between that
for static and flow-adapted cells (not shown); this result is compatible with
the observation that endothelial cells (presumably flow adapted) represent
30% of total lung cells.
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Immunoblot for KIR 6.2 protein expression (polyclonal antibodies directed against both COOH and NH2 terminus) showed a 50-kDa band consistent with that reported for the KIR 6.2 protein (15). The antibody to the COOH terminus peptide gave a relatively weak reaction with static cells but showed a significant increase in cells adapted to 1 dyn/cm2. Cells adapted to the higher shear stress were not analyzed with this antibody. The antibody to the NH2 terminus gave a more vigorous reaction; densitometric scanning of the bands revealed that immunoreactivity increased 2.2 ± 0.5-fold (n = 3) in cells exposed to 1 dyn/cm2, 2.5 ± 0.3-fold (n = 3) in cells exposed to 5 dyn/cm2, and 2.6 ± 0.5-fold (n = 3) in cells exposed to 10 dyn/cm2 (Fig. 4A). All values are statistically significant (P < 0.05) compared with cells under static culture conditions.
Expression of the SUR subunit of the KATP channel in RPMVEC was evaluated in response to shear stress of 1 dyn/cm2 for varying periods up to 72 h and 10 dyn/cm2 for periods up to 24 h. Analysis of BODIPY-FL glyburide binding demonstrated a time-dependent increase with a significantly more rapid response by cells exposed to the higher shear stress (Fig. 4, B and C). At 10 dyn/cm2 shear stress, glyburide binding reached maximum at 24 h of adaptation, whereas a similar increase in glyburide binding required 72 h of flow adaptation at 1 dyn/cm2.
Because endothelial cell membrane depolarization was observed with rat lung ischemia in situ, we studied this response in cells adapted to flow. Flow-adapted RPMVEC showed a rapid increase in bisoxonol fluorescence with stop of flow indicating plasma membrane depolarization; a return to baseline was observed with restart of flow compatible with repolarization (Fig. 5A). There was no change in bisoxonol fluorescence with stop of flow in RPMVEC that were subjected to flow for only 30 min (to load the dye) and, therefore, were not flow adapted. Cells treated with the KATP channel opener cromakalim showed no change in fluorescence upon stop of flow, providing evidence that KATP closure is required for the depolarization response. The depolarization response was studied in cells that had been treated with cycloheximide during the period of adaptation to flow. RPMVEC adapted to flow (10 dyn/cm2 for 24 h) in the presence of 10 µg/ml cycloheximide did not show the increase of bisoxonol fluorescence associated with membrane depolarization with stop of flow, indicating that this response required protein synthesis during the flow adaptation period (Fig. 5B).
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Induction of KATP channels by flow was also evaluated in BPAEC. Flow adaptation in these cells was demonstrated by cellular alignment in the direction of flow, as reported previously (24). Interestingly, alignment of RPMVEC did not occur with flow adaptation (not shown). BPAEC adapted to 1 dyn/cm2 for 72 h showed a significant increase in immunoreactive protein detected by KIR 6.2-C Ab and in the SUR detected by BODIPY-FL glyburide binding (Fig. 6). Adaptation to higher shear stress and electrophysiological responses was not evaluated with these cells.
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DISCUSSION |
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Patch-clamp studies of RPMVEC cultured under static conditions (passage 11) showed low-level expression of an inwardly rectified K+ current that was significantly increased with flow adaptation. Channel activity was sensitive to glyburide compatible with a KATP channel. Presence of the message for the KIR 6.2 pore-forming unit of the KATP channel, and absence of that for KIR 6.1, was confirmed by RT-PCR. Induction of the KIR 6.2 channel message and protein on adaptation to shear stress was demonstrated by mRNA and immunoblot analysis, whereas increased expression of SUR was shown by ligand binding. RNA and protein extracted from rat lung showed the presence of the KIR 6.2 subunit compatible with its presence in lung endothelium, although other cell types could account for some of this activity. The results suggest that the KATP channel present in pulmonary endothelial cells is downregulated when cells are cultured under static conditions and can be induced in vitro during flow adaptation. Although previous studies of endothelial cells from rat aorta and brain and rabbit aorta and pulmonary artery have identified membrane currents compatible with KATP channels (19, 20), the present results are the first demonstration in endothelial cells of the individual protein components of the KATP channel and their induction with flow adaptation.
Induction of the KATP channel components (KIR 6.2 and SUR) in cultured endothelial cells varied with the shear stress magnitude and duration. Whereas cells sheared at 5 and 10 dyn/cm2 for 24 h showed an increase in channel activity as indicated by patch-clamp studies, the change in electrical activity with adaptation to 1 dyn/cm2 for 72 h was not significant. On the other hand, there was a significant induction of protein expression at the lower level of shear stress. Because an increase in channel protein expression, as well as activity, was observed at the higher shear values of 5 and 10 dyn/cm2, it is possible that the discrepancy between the channel expression and activity at the low shear (1 dyn/cm2) arises from inadequate channel assembly. We speculate that because KATP channel activity requires a 1:1 stoichiometry of SUR and KIR 6.2 subunits, the induction of the subunits at lower shear stress does not generate the appropriate stoichiometry, resulting in a channel with lower activity. Mismatched stoichiometry has been reported to result in loss of KATP channel activity in transgenic mice (21).
Direct fluorescence studies in real time showed that flow adaptation is required for the membrane depolarization response with stop of flow and that this response can be prevented with KATP channel agonists. These results point to a role for this channel in the depolarization response with abrupt changes in shear. Abrogation of this response when cells were flow adapted in the presence of the cycloheximide, a blocker of protein synthesis, indicates that the depolarization response does not occur in the absence of increased protein expression.
Shear stress is widely accepted as a regulator of cytoskeletal organization and of cellular components such as G protein-coupled receptors, caveolae, integrins, focal adhesion kinases, and mitogen-activated receptors (11, 12, 13, 14, 16, 22, 33, 35). Our results demonstrate that shear stress also regulates KATP channel expression and activity in RPMVEC and BPAEC. Regulation likely occurs via shear stress-sensitive elements that induce gene expression because the mRNA for KIR 6.2 was increased. Shear is known to induce the transcription of some endothelial genes (23), including platelet-derived growth factor-B (PDGF-B) (29), monocyte chemotactic protein-1 (MCP-1) (30), and intercellular adhesion molecule-1 (ICAM-1) (25). Recent evidence indicates that genes regulated by shear stress respond to a shear stress response element (SSRE) defined as GAGACC (7). Our inspection of the KIR 6.2 gene (8) indicates a GAGACC sequence in the promoter region that could be the site for attachment of transcription factors. However, the mechanism for activation of the shear stress-dependent response of the KATP channel is not clear. Possibly cytoskeletal molecules such as lipid domains (caveolae or rafts), focal adhesion complexes, integrins, or the K+ channel itself might sense the change in flow, resulting in transcription factor activation.
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DISCLOSURES |
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ACKNOWLEDGMENTS |
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Present address of Abu-Bakr Al-Mehdi: Dept. of Pharmacology, University of South Alabama, Mobile, AL.
This work has been presented in part at the Experimental Biology meetings in April 2000 (San Diego, CA), April 2001 (Orlando, FL), and April 2002 (New Orleans, LA), at the International Union of Physiological Sciences meeting in Christchurch, NZ, in August 2001 and at the American Thoracic Society meeting in Seattle, WA, on May 23, 2003.
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FOOTNOTES |
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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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