ATP Depletion Induces a Loss of Respiratory Epithelium Functional Integrity and Down-regulates CFTR (Cystic Fibrosis Transmembrane Conductance Regulator) Expression
1997; Elsevier BV; Volume: 272; Issue: 44 Linguagem: Inglês
10.1074/jbc.272.44.27830
ISSN1083-351X
AutoresStéphane Brézillon, Jean‐Marie Zahm, Denis Pierrot, Dominique Gaillard, Jocelyne Hinnrasky, Hervé Millart, Jean‐Michel Klossek, Burkhard Tümmler, Édith Puchelle,
Tópico(s)Neuroscience of respiration and sleep
ResumoTo mimic the effect of ischemia on the integrity of airway epithelium and expression of cystic fibrosis transmembrane conductance regulator (CFTR), we induced an ATP depletion of the respiratory epithelium from upper airway cells (nasal tissue) and human bronchial epithelial 16HBE14o− cell line. Histological analysis showed that 2 h of ATP depletion led to a loss of the epithelium integrity at the interface between basal cells and columnar cells. The expression of connexin 43 (Cx43, subunit of the gap junctions) and desmoplakins 1 and 2 (DPs 1 and 2, major components of the desmosomes) proteins was inhibited. After 90 min of ATP depletion, a significant decrease of the transepithelial resistance (25%) was observed but was reversible. Similar results were obtained with the 16HBE14o− human bronchial epithelial cell line. ATP depletion led to actin filaments depolymerization. The expression of the mature CFTR (170 kDa) and fodrin proteins at the apical domain of the ciliated cells was down-regulated. The steady-state levels of CFTR, Cx43, DPs 1 and 2 mRNAs, semiquantified by RT-polymerase chain reaction kinetics, remained constant throughout ATP depletion in nasal tissue as in the homogeneous cell population of 16HBE14o− human bronchial epithelial cell line. This suggests that the down-regulation of these proteins might be posttranscriptional. The intercellular diffusion through gap junctions of Lucifer dye was completely inhibited after 90 min of ATP depletion but was reversible. The volume-dependent and the cAMP-dependent chloride secretion were inhibited in a nonreversible way. Taken together, these results suggest that an ATP depletion in human airway epithelium, mimicking ischemia, may induce a marked alteration in the junctional complexes and cytoskeleton structure concomitantly with a loss of apical CFTR expression and chloride secretion function. To mimic the effect of ischemia on the integrity of airway epithelium and expression of cystic fibrosis transmembrane conductance regulator (CFTR), we induced an ATP depletion of the respiratory epithelium from upper airway cells (nasal tissue) and human bronchial epithelial 16HBE14o− cell line. Histological analysis showed that 2 h of ATP depletion led to a loss of the epithelium integrity at the interface between basal cells and columnar cells. The expression of connexin 43 (Cx43, subunit of the gap junctions) and desmoplakins 1 and 2 (DPs 1 and 2, major components of the desmosomes) proteins was inhibited. After 90 min of ATP depletion, a significant decrease of the transepithelial resistance (25%) was observed but was reversible. Similar results were obtained with the 16HBE14o− human bronchial epithelial cell line. ATP depletion led to actin filaments depolymerization. The expression of the mature CFTR (170 kDa) and fodrin proteins at the apical domain of the ciliated cells was down-regulated. The steady-state levels of CFTR, Cx43, DPs 1 and 2 mRNAs, semiquantified by RT-polymerase chain reaction kinetics, remained constant throughout ATP depletion in nasal tissue as in the homogeneous cell population of 16HBE14o− human bronchial epithelial cell line. This suggests that the down-regulation of these proteins might be posttranscriptional. The intercellular diffusion through gap junctions of Lucifer dye was completely inhibited after 90 min of ATP depletion but was reversible. The volume-dependent and the cAMP-dependent chloride secretion were inhibited in a nonreversible way. Taken together, these results suggest that an ATP depletion in human airway epithelium, mimicking ischemia, may induce a marked alteration in the junctional complexes and cytoskeleton structure concomitantly with a loss of apical CFTR expression and chloride secretion function. In renal grafts, ischemia via cellular ATP depletion induces a series of structural, biochemical, and functional alterations, which lead to a loss of epithelial cell surface membrane polarity (1Molitoris B.A. Wagner M.C. Kidney Int. 1996; 49: 1592-1597Abstract Full Text PDF PubMed Scopus (37) Google Scholar). Evidence accumulated so far indicates that the dissociation of the actin cytoskeleton and associated surface membrane structures leads to numerous cellular alterations including loss of cell-cell contact, cell extracellular matrix adhesion, and surface membrane polarity of renal proximal tubule cells (1Molitoris B.A. Wagner M.C. Kidney Int. 1996; 49: 1592-1597Abstract Full Text PDF PubMed Scopus (37) Google Scholar, 2Molitoris B.A. Am. J. Physiol. 1993; 265: F693-F697PubMed Google Scholar, 3Bacallao R. Garfinkel A. Monke S. Zampighi G. Mandel L.J. J. Cell Sci. 1994; 107: 3301-3313Crossref PubMed Google Scholar, 4Mandel L.J. Doctor R.B. Bacallao R. J. Cell Sci. 1994; 107: 3315-3324Crossref PubMed Google Scholar). During renal ischemia, the disruption of junctional complexes of proximal tubule induces a redistribution of the Na/K-ATPase from the basolateral membrane domain to the apical membrane domain (1Molitoris B.A. Wagner M.C. Kidney Int. 1996; 49: 1592-1597Abstract Full Text PDF PubMed Scopus (37) Google Scholar, 2Molitoris B.A. Am. J. Physiol. 1993; 265: F693-F697PubMed Google Scholar). The lung is the only solid organ that is transplanted without restoration of systemic arterial supply and where blood flow is reduced to levels insufficient to maintain cellular energy levels (5Egan T.M. Cooper J.D. Crystal R.G. West J.B. Barnes P.J. Cherniack N.S. Weibel E.R. The Lung: Scientific Foundations. Raven Press, New York1991: 2205-2215Google Scholar). Therefore, it is essential to determine the effects of ischemia in airway epithelial structure and function to adequately restore lung function. Lung transplantation is indicated for patients with end-stage respiratory failure (6Brézillon S. Hamm H. Heilmann M. Schäfers H.-J. Hinnrasky J. Wagner T.O.F. Puchelle E. Tümmler B. Hum. Pathol. 1997; 28: 944-952Crossref PubMed Scopus (19) Google Scholar), such as patients with cystic fibrosis (CF). 1The abbreviations used are: CF, cystic fibrosis; CFTR, cystic fibrosis transmembrane conductance regulator; NBF1, nucleotide binding fold domain 1; PBS, phosphate-buffered saline; FITC, streptavidin-fluorescein isothiocyanate; RT, reverse transcriptase; RT-PCR, reverse transcription-polymerase chain reaction; LY, Lucifer Yellow; SPQ, 6methoxy-N-(3-sulfopropyl)quinolinium; ZO-1, zonula occludens 1; CK, cytokeratin; DP, desmoplakin; Cx, connexin; ER, endoplasmic reticulum; bp, base pair(s); Ω, ohm(s). 1The abbreviations used are: CF, cystic fibrosis; CFTR, cystic fibrosis transmembrane conductance regulator; NBF1, nucleotide binding fold domain 1; PBS, phosphate-buffered saline; FITC, streptavidin-fluorescein isothiocyanate; RT, reverse transcriptase; RT-PCR, reverse transcription-polymerase chain reaction; LY, Lucifer Yellow; SPQ, 6methoxy-N-(3-sulfopropyl)quinolinium; ZO-1, zonula occludens 1; CK, cytokeratin; DP, desmoplakin; Cx, connexin; ER, endoplasmic reticulum; bp, base pair(s); Ω, ohm(s). Cystic fibrosis, the most common and severe autosomal recessive disease among the northern American and European populations, is characterized by a defect in cyclic AMP-dependent chloride channel activity in a number of tissues, in particular the respiratory tract tissue (7Li M. McCann J.D. Liedtke C.M. Nairn A.C. Greengard P. Welsh M.J. Nature. 1989; 331: 358-360Crossref Scopus (257) Google Scholar). It is caused by mutations in the gene coding for the cystic fibrosis transmembrane conductance regulator (CFTR) (8Rommens J.M. Iannuzzi M.C. Kerem B.-S. Drumm M.L. Melmer G. Dean M. Rozmahel R. Cole J.L. Kennedy D. Hidaka N. Zsiga M. Buchwald M. Riordan J.R. Tsui L.-C. Collins F.S. Science. 1989; 245: 1059-1065Crossref PubMed Scopus (2479) Google Scholar, 9Riordan J.R. Rommens J.M. Kerem B.-S. Alon N. Rozmahel R. Grzelczak Z. Zielinski J. Lock S. Plasvic N. Chou J.-L. Drumm M.L. Iannuzzi M.C. Collins F.S. Science. 1989; 245: 1066-1073Crossref PubMed Scopus (5805) Google Scholar, 10Kerem B.M. Rommens J.M. Buchanan J.A. Markiewicz D. Cox T.K. Chakravarti A. Buchwald M. Tsui L.-C. Science. 1989; 245: 1073-1080Crossref PubMed Scopus (3133) Google Scholar), which is a cAMP-regulated low-conductance channel (11Riordan J.R. Annu. Rev. Physiol. 1993; 55: 609-630Crossref PubMed Scopus (301) Google Scholar). Lack or mislocalization of CFTR is regarded as being specific for CF (12Denning G.M. Ostedgaard L.S. Welsh M.J. J. Cell. Biol. 1992; 118: 551-559Crossref PubMed Scopus (156) Google Scholar, 13Puchelle E. Gaillard D. Ploton D. Hinnrasky J. Fuchey C. Boutterin M.-C. Jacquot J. Dreyer D. Pavirani A. Dalemans W. Am. J. Respir. Cell. Mol. Biol. 1992; 7: 485-491Crossref PubMed Scopus (122) Google Scholar). In normal airway surface respiratory epithelium, CFTR is restricted to the apical domain of the ciliated cells (13Puchelle E. Gaillard D. Ploton D. Hinnrasky J. Fuchey C. Boutterin M.-C. Jacquot J. Dreyer D. Pavirani A. Dalemans W. Am. J. Respir. Cell. Mol. Biol. 1992; 7: 485-491Crossref PubMed Scopus (122) Google Scholar). Whether ischemia in lung transplants may induce a redistribution of CFTR protein from the apical membrane domain remains unknown. The aim of this study was to investigate the effects of ATP depletion, simulating lung graft ischemia, on epithelium functional integrity and on CFTR expression. Our data demonstrate that ATP depletion induces an alteration of the respiratory epithelium at the interface between basal cells and columnar cells with a disruption of desmosomes and gap junction complexes. The expression of the mature CFTR protein is down-regulated, and the cAMP-mediated chloride secretion is inhibited. Our results suggest that the respiratory epithelial cells may lose polarity and CFTR chloride channel function during the ischemia of the lung transplants. Fresh nasal polyps were obtained from non-CF patients undergoing nasal polypectomy due to nasal obstruction. Immediately after the removal, the tissue material was transferred to the laboratory in Hanks' medium, which contained 20 mm HEPES and antibiotics (100 units/ml penicillin and 100 μg/ml streptomycin). Pieces of nasal polyp (approximately 1 cm2 in size) were directly snap-frozen in liquid nitrogen. Explants (1–2 mm2 in size) of human respiratory nasal epithelial tissue from nasal polyps were seeded on 35-mm tissue culture dishes coated with type I collagen. Cells were incubated in RPMI 1640 culture medium supplemented with 2 mml-glutamine, 1 μg/ml insulin, 1 μg/ml transferrin, 10 ng/ml epidermal growth factor, 0.5 mg/ml hydrocortisone, 10 ng/ml retinoic acid, and 100 units/ml penicillin/100 μg/ml streptomycin, at 37 °C. After 3 days in culture, explants were surrounded by a cell outgrowth resulting from both cell migration and cell proliferation (14Chevillard M. Hinnrasky J. Zahm J.-M. Plotkowski M.C. Puchelle E. Cell Tissue Res. 1991; 264: 49-55Crossref PubMed Scopus (42) Google Scholar). Dissociated human nasal respiratory epithelial cells were grown on coated type I collagen-carbodiimide (Sigma) Petri dishes in RPMI 1640-supplemented culture medium at 37 °C. Human bronchial epithelial cells (15Haws C. Krouse M.E. Xia Y. Gruenert D.C. Wine J.J. Am. J. Physiol. 1992; 263: L692-L707PubMed Google Scholar) were grown in Dulbeccos' modified Eagle's medium plus 10% fetal bovine serum-supplemented medium on coated type I collagen-carbodiimide (Sigma) Petri dishes. All cell cultures were performed at 37 °C in an atmosphere of 5% CO2 and 95% air before ATP depletion procedure. Inhibition of glycolysis was accomplished by washing the cells in a glucose-free modified Ringer's buffer supplemented with 2 mm glutamine, followed by a 3-h incubation in this buffer at 37 °C to deplete the tissue of endogenous metabolic substrates (3Bacallao R. Garfinkel A. Monke S. Zampighi G. Mandel L.J. J. Cell Sci. 1994; 107: 3301-3313Crossref PubMed Google Scholar). The composition of the modified Ringer's buffer was 115 mm NaCl, 20 mm Hepes, 5 mmK2HPO4, 2 mm MgSO4, 1 mm CaCl2, and 2 mm glutamine. ATP depletion was achieved by adding in the modified Ringer's buffer the respiratory inhibitor antimycin A (10 μm) and the glycolytic inhibitor 2-deoxyglucose (10 mm) (3Bacallao R. Garfinkel A. Monke S. Zampighi G. Mandel L.J. J. Cell Sci. 1994; 107: 3301-3313Crossref PubMed Google Scholar, 4Mandel L.J. Doctor R.B. Bacallao R. J. Cell Sci. 1994; 107: 3315-3324Crossref PubMed Google Scholar). Nucleoside contents were analyzed by high performance liquid chromatography (16Hull-Ryde E.A. Cummings R.G . Lowe J.E. J. Chromatogr. 1983; 275: 411-417Crossref PubMed Scopus (48) Google Scholar) from primary culture of nasal respiratory epithelial cells and from 16HBE14o− human bronchial epithelial cells. The cells were rinsed, harvested, and placed in 0.5 ml of 0.6 n perchloric acid at 4 °C. The cells were lysed using an Ultraturrax homogenizer by two pulses of 2 s. The lysates were neutralized with Tris-ethanol-amine carbonate-potassium hydroxide to a pH of 5.0–6.0. The samples were placed at 4 °C for 15 min in order to de-gas. The cell lysates were centrifuged (15 min, 4 °C, 4500 rpm), and 20 μl of the supernatant were injected on the column (C18 μBondapack, Waters, St. Quentin en Yvelines, France). The height of each nucleoside peak was measured and compared with the height of the standard peaks. Nucleoside contents were calculated and normalized according to the protein concentration of the samples. Samples designed as controls were obtained from primary culture of nasal respiratory epithelial cells and from 16HBE14o−human bronchial epithelial cells incubated for 5 h in the modified Ringer's buffer. To test the efficiency of the protocol of ATP depletion on the inhibition of cell activity, we analyzed the ciliary beating frequency. The culture dishes were placed on the heated stage of a phase-contrast inverted microscope (Nikon TMS-F) equipped with a CCD video camera (Panasonic WV CD50). The variation in light intensity, resulting from the ciliary beating, was detected by a photodetector placed on a video screen. The signal of the detector was converted into frequency spectrum by fast Fourier transform software. The mean ciliary beating frequency was calculated from this spectrum. Ciliary beating frequency was measured on 50 different ciliated cells/outgrowth (n = 3) at 0, 30, 60, and 90 min of ATP depletion. Reversibility of ATP depletion was checked by incubating the cell culture for 12 h in a fresh RPMI 1640-supplemented medium. Viability of the cells was controlled by trypan blue exclusion staining. For histological observations, nasal explants (n = 2) were fixed in formalin (15%) for 60 min at room temperature, dehydrated, and embedded in paraffin. Five-μm-thick sections were stained with hematoxylin/phloxin/safran. Two explants were fixed in 4% paraformaldehyde in 0.1 mphosphate-buffered saline (PBS), pH 7.2, for 2 h and rinsed several times (3 × 10 min) in 0.1 m PBS before being dehydrated through graded concentrations of ethanol and embedded in Epon. Semithin sections (1 μm) were cut on a Reichert-Jung Ultramicrotome (Ultracut E, Leica, Rueil-Malmaison, France). Sections were stained with toluidine blue and observed under an Axiophot microscope (Zeiss, Le Pecq, France) at a magnification of ×40. Ultrathin sections (0.07 μm) of a nasal explant were realized with a Reichert-Jung Ultramicrotome (Ultracut E, Leica) and observed using a Hitachi H300 transmission electron microscope (Elexience, Verrières-le-buisson, France) at 75 kV. The epithelial integrity of nasal outgrowth cell cultures (n = 3) and 16HBE14o− cell line cultures (n = 3) was quantified by transepithelial resistance measurements using a Millicell-ERS Resistance System (Millipore Co., Bedford, MA). Nasal outgrowth cell cultures and 16HBE14o− cell line cultures were grown on type I collagen-coated 12-mm Transwell porous cell culture inserts, 0.4-μm pore size transparent collagen membrane (Costar, Cambridge, MA). Transepithelial resistance was measured after a period of 90 min of ATP depletion and compared with that evaluated for cells incubated before ATP depletion (control) or after a 12-h incubation in cell culture control medium following the 90 min of ATP depletion. Data were expressed as means ± standard error (S.E.) of triplicate measurements. Nasal explants (n = 7) were fixed in situ in cold methanol for 10 min at −20 °C, dried, embedded in OCT (Tissue-Tek, Miles Inc.), cryofixed in liquid nitrogen, and stored at −80 °C. Cryosections of 5 μm thickness were placed onto gelatin-coated glass slides, further air-dried, and stored at −20 °C until used for immunofluorescence microscopy, as described previously (17Brézillon S. Dupuit F. Hinnrasky J. Marchand V. Kälin N. Tümmler B. Puchelle E. Lab. Invest. 1995; 72: 191-200PubMed Google Scholar, 18Dupuit F. Kälin N. Brézillon S. Hinnrasky J. Tümmler B. Puchelle E. J. Clin. Invest. 1995; 96: 1601-1611Crossref PubMed Scopus (99) Google Scholar). 16HBE14o− human bronchial epithelial cells were grown on glass coverslips coated with 200 μl of type I collagen associated with carbodiimide (Sigma). Cell cultures were fixedin situ in cold methanol for 10 min at −20 °C and analyzed for immunofluorescence microscopy.The following mouse monoclonal primary antibodies were used: mouse anti-CFTR (MATG 1061; Transgene, Strasbourg, France, raised against a synthetic peptide corresponding to the amino acid sequence 503–515 without the residue 508, thus equivalent to the ΔF 508 epitope in the NBF1 domain of the human CFTR protein, dilution 1:200; MATG 1104, Transgene, raised against a synthetic peptide corresponding to the amino acid sequence 722–734 of the R domain of the human CFTR protein, dilution 1:200; mAb 24–1 (Genzyme Corporation, Cambridge, MA) raised against the amino sequence 1377–1480 of the COOH-terminal domain of CFTR, dilution 1:200). To test the specificity of the two CFTR antibodies (MATG 1061 and 1104) and to more accurately assess the location of the CFTR epitopes, we performed peptide competition assays. We used the synthetic peptides corresponding to the epitopes in the NBF1 domain (amino acid positions 503–515 without 508) and in the R domain (amino acid positions 722–734). The blockage was complete at a peptide/IgG ratio of 8 for MATG 1061 and a peptide/IgG ratio of 2 for MATG 1104. Fodrin was detected with mouse anti-fodrin (dilution 1:10, Sigma). Mouse anti-desmoplakins 1 and 2 (clone DP 1 and 2–2.15, dilution 1:10; Boehringer Mannheim France S. A., Meylan, France), rabbit anti-connexin 43 (dilution 1:50, Dr. Gros, University of Sciences of Luminy, Marseille, France), mouse anti-β1-integrin (dilution 1:20, Dr. Sheppard, Lung Biology Center, University of California, San Francisco, CA), rabbit anti-β-catenin (dilution 1:500, Dr. Van Roy, Laboratory of Molecular Biology, University of Ghent, Ghent, Belgium), mouse anti-E-cadherin (dilution 1:100, Dr. Van Roy), rat anti-zonula occludens 1 (ZO-1) (clone 6A1, dilution 1:20, Valbiotech Genesis, France). Negative controls were performed by using nonimmune mouse IgG fraction (ref. M7769; Sigma) or by omitting the primary antibody. Secondary antibodies: goat biotinylated anti-mouse IgG fractions (Boehringer Mannheim), donkey biotinylated anti-rabbit IgG fractions (Amersham, Amersham International, UK), goat anti-rat IgG fractions (Boehringer Mannheim), and streptavidin-fluorescein isothiocyanate (FITC) (Amersham) were used at a 1:50 dilution. The sections were counterstained with Harris hematoxylin solution for 10 s, mounted in Citifluor antifading solution (Agar Scientific, Stansted, UK), and observed with a Zeiss Axiophot microscope using epifluorescence and Nomarski differential interference illumination. Nasal outgrowth cell cultures (n = 2) were grown on a glass coverslip coated with 200 μl of type I collagen associated with carbodiimide (Sigma). Cell cultures were fixed in situin cold methanol for 10 min at −20 °C and used for immunofluorescence microscopy, as described previously. The labeling of the microtubule network was realized with the mouse anti-β-tubulin (N357, dilution 1:50, Amersham). F-actin filaments were immunodetected as follows; outgrowth were fixed in 3.7% paraformaldehyde for 10 min, permeabilized in 0.5% Triton X-100 for 10 min, incubated in 1% bovine serum albumin-PBS for 5 min, labeled with phalloidin-FITC (dilution 1:50, Sigma) for 1 h, and rinsed twice in PBS. Cell outgrowths were examined with an MRC-600 Bio-Rad confocal system mounted on a Zeiss Axioplan microscope (Ivry/Seine, France). CFTR labeling in 16HBE14o− cell line was also examined by scanning laser confocal microscopy. Sequential serial sections were collected at 0.3 μm pitch. RNA extraction was performed from frozen human nasal polyps incubated in control medium (n = 3) and from frozen human nasal polyps subjected to 2 h of ATP depletion (n = 3). In parallel, RNA extraction was performed from 16HBE14o− human bronchial epithelial cells incubated in control medium and from 16HBE14o− human bronchial epithelial cells subjected to 2 h of ATP depletion. Before RNA extraction, glassware was sterilized overnight at 180 °C, and all solutions and plasticware were treated for at least 12 h with 0.1% (v/v) aqueous diethylpyrocarbonate solution (Sigma) to inactivate RNases. Frozen nasal tissue was disrupted with an Ultraturrax homogenizer by two pulses of 2 s, lysed in 7–9-fold excess volume of 6 m guanidinum isothiocyanate, 5 mm sodium citrate, pH 7.0, 0.1 m 2-mercaptoethanol, and 0.5% laurylsarcosine. The total RNA was pelleted by ultracentrifugation (15 h, 20 °C; 35,000 rpm, SW55Ti rotor; Beckman) through a CsCl cushion (18Dupuit F. Kälin N. Brézillon S. Hinnrasky J. Tümmler B. Puchelle E. J. Clin. Invest. 1995; 96: 1601-1611Crossref PubMed Scopus (99) Google Scholar). The amounts of mRNA transcripts of CFTR (exons 3–6A), Cx43, and DP 1 were semiquantified by reverse transcription (RT) and PCR kinetic assays (18Dupuit F. Kälin N. Brézillon S. Hinnrasky J. Tümmler B. Puchelle E. J. Clin. Invest. 1995; 96: 1601-1611Crossref PubMed Scopus (99) Google Scholar). Aldolase mRNA was chosen as the internal standard of a constitutively expressed housekeeping gene, to assess any degradation of the RNA and to allow a semiquantitative sample-to-sample comparison. Aldolase and the cDNA of interest (CFTR, Cx43, DP 1) were amplified separately. Purified oligodeoxynucleotides were synthesized by Transgène (Strasbourg, France). The following primers were made for the detection of mRNA transcripts: aldolase sense primer oald 141N (5′-GGCAAGGGCATCCTGGCTGCAGA), aldolase antisense primer oald 583T (5′-TAACGGGCCAGAACATTGGCATT), CFTR sense primer (exon3) (5′AGAATGGGATAGAGAGCTGGCTTC), CFTR antisense primer (exon5/exon6A boundary) for reverse transcription (5′-GTGCCAATGCAAGTCCTTCATCAA), CFTR antisense primer (exon 5) for PCR (5′-TTCATCAAATTTGTTCAGGTTGTTG), desmoplakin 1 sense primer (5′-CCGACTGACTTATGAGATTGAAG), desmoplakin 1 antisense primer (5′-GATTTTCACCAGAAGGCTCTCTC). Designed connexin 43 sense primer 98T (5′-CCTCCAAGGAGTTCAATCACTTG) and connexin 43 antisense primer 515N (5′-CCACATTGACACCATCAGTTTGG) were validated on heart tissue known to express high level of Cx43 transcripts and proteins (19Fishman G.I. Eddy R.L. Shows T.B. Rosenthal L. Leinwand L.A. Genomics. 1991; 10: 250-256Crossref PubMed Scopus (110) Google Scholar). The sizes of the expected DNA bands were: 443 base pairs (bp), aldolase; 430 bp, CFTR; 391 bp, Cx43; 324 bp, DP 1. The temperature of annealing of the primers (T°) were: 60 °C, CFTR; 58 °C, aldolase; 56 °C, Cx43; 56 °C, DP 1. First strand cDNA synthesis was performed with the Superscript preamplification system (Life Technologies, Inc.); an RNA/primer mixture of 3 μg of total RNA, 1.6 mm oligonucleotide primers, and diethylpyrocarbonate-treated water was incubated at 70 °C for 10 min and placed in ice for at least 1 min. Seven μl of a solution of 2.8 × PCR buffer, 7.14 mmMgCl2, 1.4 mm dNTP, and 0.03 mdithiothreitol were added to the volume (12 μl) of RNA/primers mixture. Two hundred units of Superscript II RT were added to the volume reaction and incubated for 50 min at 42 °C. Reverse transcription was terminated at 70 °C for 15 min. RNase H (2 units) was added to destroy the RNA strand during 20 min at 37 °C. The first strand cDNA was subjected to PCR in a thermal cycler (Biometra, Trio-Thermoblock, Göttingen, Germany). The PCR assay (50 μl of reaction volume covered with 50 μl of mineral oil) contained 0.5 mm of each oligonucleotide primer, 0.15 mm of dNTP, 1.15 mm MgCl2, 1.02 × PCR buffer, 3 μl of the RT reaction mix. After RNA/cDNA denaturation at 94 °C for 2 min, PCR (cycling parameters: 94 °C/2 min 30 s; T°/2 min; 72 °C/3 min) was run for 18–34 cycles. Aliquots (7 μl) were withdrawn in intervals of four cycles and subjected to 2% agarose gel electrophoresis. We titrated for the first reaction cycle when the cDNA became visible by ethidium bromide fluorescence during the late exponential phase of PCR and could be identified by a benchtop scanner (model GS-690 Imaging Densitometer, Bio-Rad). The amount of PCR product increased by 1 order of magnitude within four reaction cycles. Lucifer Yellow (LY) CH (Sigma) was microinjected (pressure of injection: 100 hPa) through a microcapillary (diameter of opening of tip: 0.5 μm) continuously for 2 min in the cytosol of 3 epithelial ciliated cells/outgrowth. Injections were performed on culture dishes placed in a temperature-controlled chamber (37 °C) on a stage of an inverted microscope (Zeiss IM35). The diffusion of the probe in the neighboring cells was followed by using epifluorescence microscopy (Lucifer Yellow: 400–440 nm excitation, emitted light >470 nm). Lucifer Yellow diffusion was video-recorded during the 2 min of microinjection with a low level video camera (Lhesa SIT 4036, St. Ouen l'Aumone, France). Intercellular LY diffusion was evaluated after a period of 90 min of ATP depletion and compared with that measured for cells incubated before ATP depletion (control) or after a 12-h incubation in a RPMI 1640 culture medium following the 90-min period of ATP depletion. Images were digitized as a 512 × 512 pixel, 8-bit array using a Sparc-Classic workstation equipped with a video card (Parallax Graphics, Mountain View, CA). Variation of fluorescence intensity was analyzed by a multivariate statistical technique from the temporal image series. 3.5 mm of 6-methoxy-N-(3-sulfopropyl) quinolinium (SPQ) (Sigma), was microinjected through a microcapillary for 5 s in the cytosol of 10 ciliated cells/outgrowth (n = 3). Confluent 16HBE14o− human bronchial epithelial cells were loaded with SPQ in a hypotonic chloride buffer solution (65 mm NaCl, 1.2 mmK2HPO4, 0.5 mm MgSO4, 5 mm Hepes) for 10 min at 37 °C. SPQ-loaded cells were then incubated in a nitrate buffer (103 mmNaNO3, 2.4 mm K2HPO4, 1 mm MgSO4, 10 mm Hepes) in the presence or absence of 25 μm forskolin (Sigma) at 37 °C on the stage of the inverted microscope. Intracellular SPQ was excited at 365 nm through a ×32 planachromat objective, and emission light at >395 nm was recorded for 2 s every min for 15 min with a low level video camera (Lhesa). Cell chloride efflux was evaluated after a period of 90 min of ATP depletion and compared with that measured for cells incubated before ATP depletion (control) or after a 12-h incubation in a RPMI 1640 culture medium following the 90-min period of ATP depletion. Images were digitized and analyzed as described for intercellular Lucifer Yellow diffusion. Results were expressed as means ± S.E. Student'st test was used to test the differences between the 90-min ATP-depleted cell cultures and the control cell cultures. A value ofp < 0.05 was considered to be significant. To control the ATP depletion efficiency and assess the maintenance of cell viability, nucleosides contents in one primary culture of nasal respiratory cells were measured by high performance liquid chromatography (Fig. 1). ATP depletion for 90 min induced a 93% decrease in ATP content (from 17.7 μm to 1.3 μm), a 98% decrease in ADP content (from 11 μm to 0.2 μm), and a 10-fold increase in AMP content (from 0.6 μm to 6 μm) (Fig. 1, B and C). After incubation of the cells for 12 h in fresh culture medium following the 90-min ATP depletion, a 6.6-fold increase in ATP content (from 1.3 μm to 8.6 μm), a 13.3-fold increase in ADP content (from 0.2 μm to 2.9 μm), and a 90% decrease in AMP content (from 6 μm to 0.6 μm) were observed (Fig. 1 D). Concomitantly, to control the ATP depletion efficiency and assess the maintenance of cell viability of the homogeneous 16HBE14o−human bronchial epithelial cell line, nucleoside contents were measured by high performance liquid chromatography (data not shown). ATP depletion for 90 min induced a 97% decrease in ATP content (from 63 μm to 2.1 μm), a 96% decrease in ADP content (from 23 μm to 1 μm), and a 10-fold increase in AMP content (from 1.8 μm to 18 μm). After incubation of the cells for 12 h in fresh culture medium following the 90-min ATP depletion, a reversibility of ATP depletion was observed as assessed by a 23-fold increase in ATP content (from 2.1 μm to 48.3 μm), a 28-fold increase in ADP content (from 1 μm to 28 μm), and a 50% decrease in AMP content (from 18 μm to 9 μm). Compared with control values (11.0 ± 1.6 Hz), we observed a continuous and significant (p < 0.001) decrease in ciliary beating frequency in one primary culture of nasal respiratory cells after ATP depletion (7.2 ± 1.8 Hz, 5.1 ± 1.1 Hz, 4.4 ± 1.5 Hz after 30, 60, and 90 min of ATP depletion, respectively). The ciliary beating frequency could be partially re-established after incubation of the cells for 12 h in fresh culture medium (10.8 ± 1.3 Hz, p < 0.01) (data not shown). Therefore, the ciliated cells were still viable after 90 min of ATP depletion. No significant changes could be observed in ciliary beating frequency after a 48-h incub
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