Adenovirus stimulates choline efflux by increasing expression of organic cation transporter-2
Olga L. Miakotina,1
Marianna Agassandian,1
Lei Shi,1
Dwight C. Look,1 and
Rama K. Mallampalli1,2,3
Departments of 1Internal Medicine and 2Biochemistry; and 3Department of Veterans Affairs Medical Center, Roy J. and Lucille A. Carver College of Medicine, The University of Iowa, Iowa City, Iowa
Submitted 19 May 2004
; accepted in final form 13 September 2004
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ABSTRACT
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We examined the effect of wild-type human adenovirus (Ad5) on choline transport in murine lung epithelia (MLE) and in rodent primary alveolar type II cells. Cells were active in pH-sensitive, reversible transport of choline, a process blocked pharmacologically with phenoxybenzamine, an inhibitor of organic cation transporters (OCT). PCR products for the choline transporters, OCT-1 and OCT-2, were detected, but only OCT-2 protein was robustly expressed within MLE and primary alveolar epithelial cells. Ad5 produced a two- to threefold increase in choline efflux from cells, resulting in a significant reduction in intracellular choline content and its major product, phosphatidylcholine. Effects of Ad5 on choline efflux were inhibited with phenoxybenzamine, and choline efflux was attenuated by OCT-2 small interfering RNA. Adenovirus also produced a dose-dependent increase in immunoreactive OCT-2 levels concomitant with increased cellular OCT-2 steady-state mRNA. These results indicate that adenoviruses can significantly disrupt choline trafficking in lung epithelia by upregulating expression of an alveolar protein involved in organic cation transport.
phenoxybenzamine; organic cation transporter
HUMAN ADENOVIRUSES are nonenveloped DNA viruses that have reemerged as important pathogens of acute respiratory illness (35). Adenoviral infections typically produce a wide array of respiratory illness including mild upper respiratory tract infection, bronchopneumonia, and the acute respiratory distress syndrome (ARDS). Adenoviral infection also predisposes patients to severe bacterial sepsis both in humans and in animal models (18, 22, 34). Severe adenoviral infection is characterized pathologically by a multifocal necrotizing alveolitis, alveolar hemorrhage, viral inclusions within alveolar epithelial cells, and hyaline membranes typical of diffuse alveolar damage or ARDS (27). The observation that alveolar epithelia are a cellular target for this pathogen led us to investigate whether adenoviral infection might alter various aspects of alveolar epithelial function.
It has been demonstrated that alveolar cells, via specific transport systems, are actively involved in the uptake and internalization of essential cationic elements used for the maintenance of lung fluid and phospholipid homeostasis. For example, alveolar epithelia highly express transporters such as the Na+/K+ ATPase, aquaporins, and epithelial sodium channels involved in clearance of lung edema fluid (1, 3, 5, 33, 41). Expression of some of these systems appears to be dysregulated by adenoviruses (44). Likewise, tight homeostatic regulation of transport of other cationic molecules, such as choline, is essential in the lung as choline serves as a substrate for phosphatidylcholine (PC), the major phospholipid constituent of eukaryotic cell membranes and of pulmonary surfactant. Lung epithelia express a potential-dependent, low-affinity, Na+-independent choline transport system localized to the basolateral alveolar membrane, an apical Na+-dependent, choline+/Na+ cotransporter, and an electroneutral choline/H+ exchanger against a concentration gradient (7, 11, 17, 30). Although the electrophysiological properties of these choline systems are well described, the identity and molecular regulation of pulmonary choline transport proteins remain relatively unexplored.
The organic cation transporters (OCTs) are a superfamily of transporters recently described that mediate the bidirectional, electrogenic transport of small organic cations independently of sodium (4, 6, 20, 21, 46). OCT members from several species have been cloned and sequenced; to date three isoforms, OCT-1, OCT-2, and OCT-3, sharing a high degree of sequence identity and yet differentially expressed in various tissues, have been described (4, 21). OCT-1 is detected in kidney, liver, and small intestine, whereas OCT-2 has been identified primarily within the kidney and brain (4, 21). Immunohistochemical analysis identifies OCTs within the basolateral membranes of polarized epithelia (29, 45). Structurally, OCT proteins have twelve transmembrane
-helices, four highly conserved intracellular sequences, and consensus phosphorylation sites for several kinases including protein kinase A, casein kinase, and tyrosine kinase (4, 21). All OCTs share broad overlap in substrate requirements, and overexpression of OCT-1 and OCT-2 regulates choline uptake and efflux (2, 4, 42). Mutations within the primary sequence of OCT-1 result in loss of function of the transporter with regard to choline transport, and targeted disruption of the transporter in mice impairs hepatic accumulation and intestinal and renal excretion of cations (12, 1416). Interestingly, adenovirus triggers efflux of choline from plasma membranes, but whether these effects are mediated by OCT proteins is unknown (37). Thus the aims of this study were to 1) examine OCT expression within distal lung epithelia, 2) determine their role in choline transport, and 3) determine whether OCT expression might be modulated by human adenoviruses. We hypothesize that OCT proteins partake in alveolar choline transport and that their expression will be upregulated by human adenoviruses.
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MATERIALS AND METHODS
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Materials.
The murine lung epithelial (MLE-12) cell line was purchased from American Type Culture Collection (ATCC, Manassas, VA). All culture medium was obtained from the University of Iowa Tissue Culture and Hybridoma Facility (Iowa City, IA). Wild-type human adenovirus type 5 (Ad5) or E1A/B-deficient adenovirus (Ad5-d312) was obtained from Dr. S. Brody (Washington University, St. Louis, MO). [Methyl-3H]choline chloride (specific activity of 85 Ci/mmol) was obtained from American Radiolabeled Chemicals (St. Louis, MO). Tri-Reagent, tetraethylammonium (TEA), N-methyl-D-glucamine, quinine, hemicholinium-3, and phenoxybenzamine were obtained from Sigma (St. Louis, MO). Guanidine was purchased from Amresco (Solon, OH). The protease inhibitor cocktail and murine monoclonal antiadenovirus 2 EIA antibody was obtained from CalBioChem (Darmstadt, Germany). Choline mass was assayed by liquid chromatography/electrospray ionization-isotope dilution mass spectrometry at the University of North Carolina Core facility. Rabbit anti-rat OCT-1 and OCT-2 antiserum and rat OCT-2 control peptide were obtained from Alpha Diagnostic International (San Antonio, TX). The chemiluminescent reagent SuperSignal was purchased from Pierce (Rockford, IL). Nitrocellulose membranes (0.45 µm) were obtained from Bio-Rad Laboratories (Hercules, CA). Assays-on-Demand mouse primers, GAPDH primers, Taqman reverse transcription reagents, SYBR Green PCR master mix, and the probe for solute carrier family 22, member 2 (OCT-2) (ID Mm00457295 m1) were obtained from Applied Biosystems (Foster City, CA). Cell numbers were determined using a Coulter Z1 dual particle counter (Coulter, Miami, FL). The silencer small interfering (si) RNA cocktail kit (RNase III) was from Ambion (Austin, TX). SuperScript III RNase H reverse transcriptase, scrambled siRNA was obtained from Invitrogen (Carlsbad, CA), and FuGENE 6 transfection reagent were purchased from Roche Diagnostics (Indianapolis, IN). The Messenger RNA isolation kit was from Stratagene (La Jolla, CA).
Cell culture and adenoviral infection.
Rat and murine primary alveolar type II epithelial cells were isolated and characterized as described (23, 32). The purity of type II cells from each species was >95% in our laboratory as determined by tannic acid staining (23). Adenoviruses were propagated and titered as described previously using human 293 cells (ATCC) (24). Cells were isolated and infected with Ad5 at various multiplicities of infection (MOIs) in serum-free Dulbecco's modified Eagle's medium (DMEM) and cultured in DMEM with 10% heat-inactivated fetal bovine serum (FBS) for up to 24 h. MLE cells were maintained in Hite's medium with 2% heat-inactivated FBS at 37°C in atmosphere containing 5% CO2. After reaching confluence, the cells were collected using 0.25% trypsin with 0.04% EDTA and plated into 24-well plates (
100,000 cells/well) or 60-mm (
1,400,000/dish) or 100-mm tissue culture dishes (
2,500,000 cells/dish) for 24-h culturing before each experiment. After reaching subconfluence (5060%), cells were incubated with Hite's medium with 2% FBS alone (control medium) or in combination with Ad5 or Ad5-d312 at various MOI for 624 h.
For in vivo infection, C57BL/6N mice were deeply anesthetized with ketamine (50 mg/kg ip) and xylazine (7.5 mg/kg ip), and following adequate anesthesia, the larynx was well visualized under a fiber-optic light source before endotracheal intubation with a -inch, 24-gauge plastic catheter. Human replication-competent adenovirus [Ad5, 5 x 108 plaque-forming units (pfu) in 50 µl of 10 mM Tris·HCl, pH 8.1, 100 mM NaCl, and 0.1% BSA] or diluent was instilled intratracheally on day 1, after which the animals were allowed to recover under BL2 containment for 24 h before a second intratracheal dose on day 2. Mice were killed with pentobarbital (150 mg/ip) 48 h after the initial inoculation. Murine lungs were lavaged, and surfactant pellets were isolated as described previously (36). These procedures are in accordance with the protocols approved by the University of Iowa Animal Care and Use Committee.
Choline uptake.
Cells were plated in 24-well plates for analysis of choline uptake. Internalization was measured in choline-free uptake buffer that contained 25 mM HEPES/Tris pH 7.4, 120.6 mM NaCl, 4.16 mM KCl, 1.05 mM CaCl2, 0.407 mM MgSO4, 17.51 mM D-glucose, and 10 or 20 nM [3H]choline chloride at 25°C. Medium was discarded, and cells were washed with unlabeled choline-free uptake buffer once and incubated in 250 µl of buffer with radiolabeled choline for various periods of time at 25°C. Reactions were terminated by three rinses with ice-cold 1x PBS. Cells were lysed in 0.5 ml of 1% SDS/0.2 M NaOH, and cellular incorporation of radiolabeled choline was determined by scintillation counting (10). In select experiments choline transport was assayed with uptake buffer devoid of Na+ or by using sodium thiocyanate as a chloride substitute.
To assess pH dependence for choline uptake, cells were rinsed with choline-free uptake buffer twice at varying pH (pH 5.58.5) and incubated in 250 µl of buffer at a desired pH for 10 min in the presence of 20 nM [3H]choline chloride at 25°C. Buffer used for pH dependence contained 25 mM MES/Tris at pH 5.5 or a mixture of 25 mM MES/Tris and 25 mM HEPES/Tris at a pH of 6.5 and 7.5, and 25 mM HEPES/Tris at pH of 8.5 (10, 45). Reactions were stopped by three quick rinses with ice-cold 1x PBS, cells were lysed in 0.5 ml of 1% SDS/0.2 M NaOH, and incorporation of radiolabeled choline into cells was determined by scintillation counting.
For inhibitory studies, cells were pretreated with choline-free uptake buffer with or without the following: TEA, quinine, hemicholinium-3, guanidine (at 50, 200, and 500 µM), or phenoxybenzamine (at 115 µM) for 15 min at 25°C. Cells were exposed to choline-free uptake buffer with 10 nM [3H]choline chloride for 10 min with or without inhibitors at 25°C. Uptake was stopped with ice-cold PBS, cells were lysed, and cellular [3H]choline incorporation was analyzed as above. In all uptake studies, two or three wells of the same plate were harvested in 0.5 ml of PBS/0.1% SDS for protein determination, and uptake was corrected for protein concentration. Choline uptake measured in cells at 0°C for 10 min served as a blank.
Choline efflux.
Cells grown in 100-mm dishes were washed twice and incubated with choline-free uptake buffer in the presence of 50 nM [3H]choline chloride and 4.95 µM unlabeled choline chloride (total concentration of choline was 5 µM) at 37°C for 30 min. Cells were then rinsed twice with choline-free uptake buffer and incubated with fresh buffer at 25°C. Aliquots of medium were taken at different time points to determine efflux of radiolabeled choline by scintillation counting (7, 43). Cells were rinsed twice with ice-cold PBS and scraped in 2 ml of cold MeOH/H2O (5:4), and efflux was corrected for protein concentration.
For inhibitory studies, cells were grown in 24-well plates, preloaded with 50 nM [3H]choline chloride and 4.95 µM unlabeled choline chloride in uptake buffer at 37°C for 30 min, washed, and then incubated in fresh uptake buffer in presence or absence of phenoxybenzamine (115 µM) for 10 min at 25°C. Aliquots of media were sequentially taken for scintillation counting, and values were corrected for protein concentration.
Apical and basolateral choline transport.
MLE cells were plated in Transwells (six-well plate, 4.7-cm2 membrane growth area, polyester membrane, 0.4-µm pore size; Corning, Corning, NY) and grown until 100% confluence for 34 days in Hite's medium with 2% FBS. For choline uptake measurements, cells were rinsed with uptake buffer containing 120.6 mM NaCl and incubated in uptake buffer with 20 nM [3H]choline chloride added basolaterally, apically, or in both chambers for 5 min at 25°C. The uptake reaction was stopped with cold PBS, and cells were lysed in 1 ml of 1 M NaOH followed by neutralization with 1 ml of 1 M HCl and used for analysis of cellular [3H]choline incorporation and protein levels in each well. For choline efflux, cells were rinsed with uptake buffer containing 120.6 mM NaCl at 25°C and incubated with uptake buffer in the presence of 50 nM [3H]choline chloride, and 4.95 µM unlabeled choline chloride was added basolaterally, apically, or in both chambers at 37°C for 30 min. Cells were then rinsed twice with uptake buffer in both chambers and incubated with fresh buffer at 25°C. Aliquots of media were then taken to determine radiolabeled activity as above, and choline efflux was corrected to protein level in each well.
Intracellular choline and PC analysis.
For determination of choline mass, subconfluent cells (
60%) were infected with Ad5 for 24 h. Cells were rinsed once with ice-cold 1x PBS, scraped in 1 ml of 1x PBS/1 mM phenylmethylsulfonyl fluoride (PMSF), pelleted at 500 g for 5 min, and resuspended in 1x PBS/1 mM PMSF. Cells were counted, aliquots were taken for protein determination, and samples were analyzed by liquid chromatography/electrospray ionization-isotope dilution mass spectrometry as described (19).
We also determined intracellular radiolabeled free choline pool size by washing cells with low-choline M-199 medium twice followed by incubation in M-199 medium for 1 h at 37°C. Cells were then labeled with [3H]choline (5 µCi/60-mm dish) for 15 min in M-199 at 37°C, washed with label-free medium, and incubated in the same medium for an additional 15 min. Cells were then rinsed in ice-cold PBS, harvested in MeOH/H2O (5:4), and processed for free [3H]choline by paper chromatography as described (25).
We assayed PC by extracting lipids, resolving individual phospholipids by thin-layer chromatography, and by quantitative analysis using the phosphorus assay as described (36). Disaturated PC (DSPC) was analyzed similarly but after treatment of lipids with osmium tetroxide as described (36).
Detection of OCT transcripts.
RT-PCR reactions were performed using poly(A)+ RNA samples isolated from MLE cells (Messenger RNA isolation kit) using two gene-specific primer sets for OCT-1 and OCT-2. Total RNA from mouse kidney served as a positive control. The sense and antisense primers for OCT-1 were 5'-gttggttgccttccagatgt-3' and 5'-ggctgtcgttctcctgtagc-3', respectively, resulting in generation of an 861-bp PCR product, and 5'-gctacaggagaacgacagcc-3' and 5'-gcctttgatttcctccttcc-3', respectively, with an expected PCR product of 922 bp. The sense and antisense primers for OCT-2 were 5'-aacccttcgttcctggactt-3' and 5'-gttgaccaggcagaccattt-3', respectively, resulting in a 369-bp PCR product and 5'-acggcagatgaggatactgg-3' and 5'-aggccaaccacagcaaatac-3', respectively, with a PCR product of 551 bp. PCR was performed using an Ampli Taq Gold cDNA polymerase with the following profile: 95°C for 10 min, then 95°C for 15 s, and 60°C for 3 min for a total of 50 cycles.
Quantitative detection of murine type II cell OCT-2 transcripts.
Cells were harvested in Tri-Reagent. After total RNA isolation and DNA digestion with DNase I, cDNA was generated using random hexamers and SuperScript III RNase H at 50°C for 4 h (Invitrogen) (47). Real-time PCR was then performed on cDNA using the Applied Biosystems 7000 real-time PCR instrument. Universal PCR master mix, Assays-on-Demand mouse primers, and probe for solute carrier family 22, member 2 (OCT-2), were used to detect OCT-2 mRNA, whereas the SYBR Green PCR master mix and rodent GAPDH primers were used for GAPDH detection. Validation of amplification rates for OCT-2 and GAPDH, an internal control, showed that the relative quantitation of OCT-2 transcript, 
CT method, could be applied.
Synthesis of siRNA.
RNA from mouse liver and kidney were isolated using Tri-Reagent and transcribed into cDNA with SuperScript III RNase H reverse transcriptase. RT reactions from the two tissues were pooled and used for PCR. Primers containing the T7 promoter and mouse OCT-2 (accession no. BC015250, shown as underlined text) sequences were used to synthesize a 467-bp PCR product (sense primer 5'-taa tac gac tca cta tag gga gac aac tgg agg tgg ctg caa t-3', anti-sense primer 5'-taa tac gac tca cta tag gga gat gac aca gcc cag gga tag c-3'). We used 100 nM of each primer in the PCR mixture. PCR conditions were 95°C for 10 min followed by 40 cycles of 94°C for 30 s, 65°C for 1 min, 75°C for 2 min, and a final extension step at 75°C for 10 min. This product was then used to synthesize double-stranded RNA followed by siRNA using the silencer siRNA cocktail kit.
Immunoblot analysis.
Cells were collected in PBS, spun at 500 g for 5 min, and resuspended in lysis buffer (10 mM Tris·HCl, pH 7.4, 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1.5 mM MgCl2, 50 mM NaF, 5 mM sodium pyrophosphate, 0.2 mM sodium orthovanadate, 10% glycerol, 1% Triton X-100, 0.5% Nonidet P-40, and 1 mM PMSF plus 1:50 protease inhibitors cocktail). Cells were sonicated, and cellular debris was pelleted at 12,000 g for 10 min at 4°C. Equal amount of protein cell lysate from each sample was separated using a 7.5% polyacrylamide gel and probed for OCT-1 or OCT-2 (1:4001:1,000 dilution) or
-actin (1:5,000 dilution). Reactive bands were visualized by chemiluminescence.
Transfectional analysis.
Cells were transfected with OCT-2 siRNA. MLE cells grown on six-well plates were transfected by adding 0.5 ml of serum-free Hite's medium containing 100 nM siRNA and 2.7 µL FuGENE 6 reagent for 24 h. After 4 h, the medium was diluted with 1 ml of Hite's medium with 2% FBS. Cells were preloaded with [3H]choline chloride, and aliquots of medium were collected and analyzed for choline efflux as above.
Protein analysis.
Protein concentrations were determined using a Lowry method or Bio-Rad protein assay.
Statistical analysis.
All data were analyzed by a paired t-test or one-way ANOVA with a Bonferroni test for multiple group analysis. Data are presented as means ± SE. Statistical significance was accepted at the P < 0.05 level.
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RESULTS
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Effect of adenovirus on PC and DSPC levels.
Human adenovirus infection in C57BL/6N mice decreased the levels of PC in murine lung lavage by 45% [378 ± 33 nmol phospholipid phosphorus/mg protein (control mice) to 209 ± 39 nmol phospholipid phosphorus/mg protein (Ad5-treated mice), P < 0.05, n = 6 mice/group]. These effects on PC levels in vivo were achieved with a second inoculum of virus as a single inoculum was insufficient to reduce phospholipid levels (data not shown). Preliminary studies also revealed that MLE cells infected with Ad5 contained
30% lower levels of DSPC, the major phospholipid component of surfactant, compared with uninfected control cells. These in vivo and in vitro results suggest that limited availability of choline might be one explanation for reduced PC levels in response to adenoviral infection.
Choline transport in MLE.
Choline levels in cells are actively regulated by uptake and efflux. We characterized choline uptake in MLE cells in the presence or absence of Na+ (Fig. 1A). Cells accumulated choline over a 60-min period in a linear manner. At initial time points of analysis, choline uptake was observed to be Na+ dependent (Fig. 1A; e.g., at the 10-min time point values with Na+ were 1.47 ± 0.29 vs. 1.88 ± 0.29 pmol/mg without Na+; P < 0.05). In experiments performed where Na+ was replaced by N-methyl-D-glucamine, this linear pattern of choline transport did not differ significantly (n = 4).

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Fig. 1. Choline transport in lung epithelia. A: choline uptake by murine lung epithelial (MLE) cells in the presence or absence of sodium. MLE cells were incubated in choline-free uptake buffer containing 120.6 mM sodium chloride (Na) or 120.6 mM N-methyl-D-glucamine (NMDG) in the presence of 10 nM [3H]choline chloride for 160 min. Cells were lysed, and [3H]choline was counted by scintillation counting. Choline uptake measured at 0°C for 10 min served as a blank. Data are expressed as means of 6 independent experiments ± SE; #significant difference in choline uptake measured in sodium-containing buffer compared with NMDG buffer (P < 0.05, paired t-test). B: choline uptake is pH sensitive. Cells were incubated in choline-free uptake buffer that contained 120.6 mM sodium chloride or 120.6 mM NMDG for 10 min in presence of 20 nM [3H]choline chloride at varying pH optima. Data are expressed as means of 4 independent experiments in duplicate ± SE, * and #, P < 0.05 vs. uptake at pH 5.5 by ANOVA. C: choline efflux is not sensitive to sodium or chloride. MLE cells were preloaded with 50 nM [3H]choline chloride and 4.95 µM unlabeled choline chloride in choline-free buffer at 37°C for 30 min. The buffer contained sodium and chloride (+Na, +Cl), or sodium containing thiocyanate (NaSCN), as a chloride substitute (+Na, Cl), or NMDG and sulfate to substitute sodium and chloride, respectively (Na, Cl), all at a pH of 7.5. Cells were then rinsed and incubated in the respective fresh buffers at 25°C for 5 min. Aliquots of media (100 µl) were taken for counting of labeled choline in duplicates. Cells were rinsed, lysed in 1 M NaOH, and counted for labeled choline. Data were normalized to total labeled choline contained in cells and medium and expressed as a percentage of effluxed choline. The values are expressed as means of 3 independent experiments performed in triplicate ± SE. D: choline efflux is stimulated by acidic pH. Choline efflux was measured in MLE cells in choline-free buffer, which contained sodium and chloride, at various pH as described in C. Data demonstrate the results of 3 experiments performed in triplicate and expressed as means ± SE. *Statistically significant difference in choline efflux compared with pH 5.5 (P < 0.05, paired t-test, n = 3). E: choline uptake in mouse primary type II cells is sensitive to pH but not to sodium or chloride. Primary type II cells isolated from 4 mice were incubated overnight in medium containing 10% FBS to allow cells to adhere to plastic dishes. Choline uptake in cells was assayed the next day using different uptake buffers in presence of 20 nM [3H]choline chloride for 10 min as described in A. Two uptake buffers contained sodium and chloride at pH 5.5 and 7.5, a 3rd buffer included NaSCN (+Na, Cl), and a 4th buffer contained NMDG and sulfate (Na, Cl), both at a pH of 7.5. Data represent 3 experiments performed in triplicate, and data are expressed as means ± SE. F: choline efflux in mouse primary type II cells is sensitive to pH but not to sodium or chloride. Isolated primary cells from 4 mice were isolated and used to measure choline efflux as in E. Cells were incubated in 4 choline-free buffers as in E (+Na, +Cl, pH 5.5, +Na, +Cl, pH 7.5, +Na, Cl, pH 7.5 and Na, Cl, pH 7.5) following the protocol described in C. Data are from 3 experiments in triplicate and expressed as means ± SE.
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We next evaluated pH dependence of choline uptake in MLE cells. Choline accumulation was inhibited by an extracellular pH <7.5 and stimulated by alkaline conditions in either sodium-free or sodium-containing medium (Fig. 1B). Sodium inhibited choline accumulation within a pH range of 6.58.5. These results are consistent with organic cationic transport in other systems (17, 39, 40, 45).
Additional studies were performed to assess choline efflux in MLE cells. Efflux was observed to be insensitive to either Na+ or Cl (Fig. 1C). Moreover, unlike uptake, efflux of choline was stimulated approximately eightfold from alkaline (pH 8.5) to acidic conditions (pH 5.5) (Fig. 1D). To investigate whether these observations were unique to the cell line, we assayed choline transport in primary distal lung epithelia (Fig. 1, E and F). Mouse primary alveolar epithelia exhibited similar pH dependence as observed with MLE cells with regard to both choline uptake (Fig. 1E) and efflux (Fig. 1F) and also exhibited Na+ and Cl independence. Thus MLE cells recapitulate many of the choline transport properties of primary cells.
To study polarized choline uptake, MLE cells were plated in Transwells, which separate apical and basolateral compartments. We compared choline uptake when cells were labeled basolaterally, apically, or from both sides. Figure 2A demonstrates that choline accumulated basolaterally and apically, although there was a trend for greater apical uptake.

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Fig. 2. Choline transport in Transwells. A: MLE monolayers established in Transwells were incubated in choline-free uptake buffer with 20 nM [3H]choline chloride added basolaterally (Baso), apically (Apic), or in both chambers (Both) for 5 min at 25°C. Cells were then washed and lysed in 1 ml of 1 M NaOH followed by neutralization with 1 ml of 1 M HCl. We used 1.5 ml of cell lysate for uptake determination and correction for protein. Data are expressed as means of 3 independent experiments performed in duplicate ± SE. B and C: choline efflux in MLE cells plated in Transwells. Cells were prelabeled in choline-free uptake buffer in the presence of 50 nM [3H]choline chloride and 4.95 µM unlabeled choline chloride added basolaterally, apically, or in both chambers at 37°C for 30 min. Cells were then rinsed in both chambers and incubated in a fresh buffer at 25°C. Aliquots of media (100 µl), basolateral and apical, were taken for counting of labeled choline at 5- and 30-min time points (B) or only after a 5-min incubation (C). Data were normalized for protein concentration. The values are expressed as means of 2 independent experiments performed in triplicate ± SD.
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We also examined whether choline efflux in lung epithelia occurs in apical and basolateral membrane. To assess paracellular choline transfer, we generated confluent monolayers and added labeled choline into either the basolateral or apical chamber, washed cells, and counted the changes in effluxed radiolabeled choline in each chamber with time. When the radiolabel was added into the basolateral chamber, we detected 30% of radioactivity effluxed basolaterally vs.
70% apically after either 5 or 30 min of incubation (Fig. 2B). When we added labeled choline into the apical chamber, only 7% of radioactivity was detected in basolateral chamber vs. 93% of activity apically; this ratio did not appreciably change by 30 min (Fig. 2B). Because the ratio of basolateral-apical effluxed choline remained approximately the same during 30 min of incubation, we conclude that only limited paracellular transfer of choline occurs in this model system. Next we compared choline efflux when labeled choline was added only in one chamber or in both chambers (Fig. 2C). Apical choline efflux was greater than basolateral, and efflux was additive when labeled choline was added in both chambers. Together, the data presented indicate bidirectional choline transport in murine lung epithelia with a tendency for greater apical transport compared with basolateral activity.
Adenovirus inhibits choline uptake and stimulates choline efflux.
We next determined whether bidirectional, pH-sensitive choline transport activity might be modulated by human adenovirus. MLE cells were infected with Ad5 or Ad5-d312 for 24 h, and choline uptake was assayed (Fig. 3A). Ad5 was added at MOI 3, resulting in infection of >
95% of cells as assessed by nuclear adenoviral E1A oncoprotein expression detected by immunofluorescence staining (results not shown). Unlike replication-deficient Ad5-d312, Ad5 significantly inhibited choline uptake by
60% compared with control, an effect that was maximal at 24 h after initiation of infection (Fig. 3, A and B).

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Fig. 3. Adenovirus 5 (Ad5) modulates choline transport in lung epithelia. Choline uptake in MLE cells in presence of E1A and B-deficient virus, Ad5-d312 (Delta-Ad), and wild-type Ad5. Cells were infected with Ad5-d312 or Ad5 for 24 h at a multiplicity of infection (MOI) = 3. Choline uptake was measured in uptake buffer with 10 nM [3H]choline chloride for 15 min at 25°C. Cells were lysed, and labeled choline in cells was counted using a scintillation counter. Choline uptake measured at 0°C for 15 min served as a blank. Data are expressed as means of 3 independent experiments ± SE, * and #, significant difference from controls and Ad5-d312, respectively (P < 0.05, ANOVA, n = 3). B: time course of Ad5 effects on choline uptake. Cells were exposed to Ad5 for various times, and choline uptake measured as described in Fig. 1A. Data represent means of 4 independent experiments ± SE; *P < 0.05 vs. control (n = 4, t-test). C: Ad5 effects on choline efflux. Cells were infected with or without Ad5 at an MOI = 3 for 24 h. Cells were preloaded with 50 nM [3H]choline chloride and 4.95 µM unlabeled choline chloride in choline-free uptake buffer at 37°C for 30 min, washed, and incubated in unlabeled uptake buffer at 25°C for 115 min. Aliquots of media were taken, and labeled choline was counted using a scintillation counter. Data are expressed as means of 3 experiments performed in triplicate ± SE; *P < 0.05 vs. control at respective time points (n = 3, t-test).
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Because choline transfer in cells is bidirectional, we preloaded control and Ad5-infected cells with radiolabeled choline and measured choline efflux. Indeed, efflux of choline increased over time in a linear manner up to 15 min (Fig. 3C). Moreover, Ad5 produced a twofold increase in cellular choline efflux above values seen with uninfected cells. In contrast, Ad5-d312 did not produce similar effects (data not shown). Collectively, these data indicate the existence of a bidirectional choline transporter(s) in MLE cells that is adenovirally regulated.
Adenovirus limits choline availability in lung epithelia.
The above data demonstrate that adenovirus might limit choline availability within alveolar epithelia because of increased cellular export coupled with reduced choline uptake. To confirm this, we first measured the intracellular pool of radiolabeled choline in cells. Ad5 compared with control only produced a modest (13%) decrease in the detectable pool of radiolabeled free choline (data not shown). This method potentially detects a small pool of choline distinct from a reservoir involved in phospholipid synthesis and may not accurately reflect total cellular choline mass. Furthermore, there is a rapid flux of radiolabeled free choline to other intermediates within the PC biosynthetic pathway (38). We therefore analyzed choline mass directly. Ad5 significantly reduced choline levels from 1.155 ± 0.080 nmol/mg protein (control) to 0.619 ± 0.017 nmol/mg protein (Ad5) after 24 h of exposure in MLE cells. Thus adenoviral modulation of choline transport results in a net decrease in cellular choline levels.
Adenovirus upregulates OCT-2 gene expression.
The existence of a bidirectional, sodium-independent choline efflux process in alveolar epithelia led us to investigate the OCT family of genes. We first examined gene expression of OCT-1 and OCT-2 (Fig. 4). RT-PCR using two sets of primers for OCT-1 revealed the presence of 922-bp and 861-bp products in mouse kidney and MLE cells (Fig. 4A). RT-PCR also showed ampliers for OCT-2 of 369 and 551 bp in mouse kidney and within MLE cells (Fig. 4B). To assess effects of adenovirus on steady-state OCT mRNA expression, we infected primary murine type II cells with Ad5 for 24 h. Analysis conducted on primary cells revealed that Ad5 produced a twofold increase in mouse OCT-2 mRNA expression, which was corrected to the internal control, the expression of the GAPDH transcript in the same samples (Fig. 4C). The relative amount of cellular OCT-2 mRNA in primary murine type II epithelia was detected at only 0.0010.002% of that seen in mouse kidney. The results suggest that adenovirus increases OCT-2 gene expression in lung epithelia.

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Fig. 4. Ad5 increases organic cation transporter (OCT)-1 and OCT-2 gene expression in lung epithelia. Poly(A)+ RNA isolated from MLE cells and mouse kidney (M. kidney) total RNA were used for RT-PCR for OCT-1 (A) and OCT-2 (B) with 2 sets of gene-specific primers. RT-PCR products were separated on an agarose gel with a molecular size marker (100-bp ladder). The negative control (Neg. C) contained no template. C: relative amount of OCT-2 mRNA in mouse primary type II cells in presence of Ad5. Isolated mouse primary type II cells were infected with Ad5 for 24 h and harvested, and total RNA was isolated. We used 3.6 µg of total RNA for RT reactions followed by quantitative real-time PCR. Total RNA of mouse kidney served as a positive control. The data represent RNA isolations from 10 mice (means ± SD).
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Adenovirus induces OCT-2 levels.
We next examined whether OCT-1 and OCT-2 proteins are adenovirally regulated in lung epithelial cells. To assess the specificity of our antibodies for immunoblotting, protein lysates from freshly isolated primary mouse and rat type II cells were resolved by SDS-PAGE, transferred to nitrocellulose, and immunoblotted with OCT-2 affinity-purified antiserum in presence or absence of blocking OCT-2 control peptide (100 µg/5 µg of antibody). In presence of blocking peptide, the intensity of the OCT-2 (
60 kDa) product was remarkably reduced (Fig. 5A).
OCT-1 was not detectable in either primary mouse type II epithelia or MLE cells but was present in mouse kidney (Fig. 5B). In contrast to OCT-1, OCT-2 was robustly detected in mouse primary type II epithelial cells cultured for 24 h, but relative to cultured primary cells, more limited expression was observed in MLE cells. Rodent fibroblasts and alveolar macrophages did not express much OCT-1, but they expressed OCT-2 (Fig. 5C). Furthermore, two faster migrating products and one predominant band reacted with the OCT-2 antibody in rat and mouse lung cells, respectively (Fig. 5C). In rat tissue, OCT-2 appeared as a primary band at
5866 kDa and a secondary product at
5255 kDa; in mouse tissue, it was visible mostly as a
70-kDa single band. This pattern resembles expression of OCT-2 that appears either as a double band from posttranslational processing or alternative splicing when expressed in oocytes or as a single 74-kDa product (26, 45). Overall, OCT-2, unlike OCT-1, is detectable and appears to be the predominant isoform expressed within MLE and alveolar epithelial cells.
We used rat primary type II cells to assess Ad5 effects as these cells were isolated in greater quantities for in vitro analysis. Interestingly, cells tended to rapidly lose OCT-2 expression after 24 h in culture (Fig. 5D). In a representative experiment, when rat primary type II cells were infected with Ad5 for 24 h at different MOI, OCT-2 expression was significantly increased compared with uninfected cells (Fig. 5D). E1A protein expression generally correlated with MOI after viral exposure in cultured primary epithelia (Fig. 5D, bottom). Note that adenovirus upregulated only the expression of the upper OCT-2 band without altering the lower band (Fig. 5D). Similar to cultured rat primary epithelia, MLE cells constitutively expressed low levels of OCT-2; Ad5, however, produced a robust increase in immunoreactive OCT-2 levels after 24-h infection (Fig. 5E).
Manipulation of OCT alters choline efflux.
Because lung epithelia expressed OCT, we assessed the capacity of cells to partake in choline uptake in the presence of other known organic substrates/inhibitors for this class of transporters: TEA, quinine, guanidine, and hemicolinium-3 (Fig. 6A). Choline uptake was inhibited by all OCT substrates at concentrations of 50500 µM in the following order: hemicolinium-3 > quinine > guanidine > TEA. TEA only slightly inhibited choline uptake. These results demonstrate existence of a polyspecific cation transporter(s) in lung epithelia.

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Fig. 6. Pharmacological and genetic inhibition of OCT and choline efflux. A: inhibition of choline uptake by organic cations/inhibitors. Cells were preincubated in choline-free uptake buffer in presence of TEA, quinine, hemicholinium-3 (HC), or guanidine (0500 µM) for 15 min, and choline uptake was determined after a 10-min incubation as in Fig. 1A. A representative experiment of 3 independent experiments is shown, data are expressed as means of 3 wells ± SE. B: phenoxybenzaminc (PbA) does not alter choline uptake. Cells were preincubated with PbA at various concentrations for 15 min at 25°C and then pulsed with 20 nM labeled uptake buffer for 10 min in the presence of the inhibitor, washed, harvested with 0.2 M NaOH, and counted for [3H]choline chloride. Data summarize 3 independent experiments performed in triplicate and are expressed as means ± SE. *P < 0.05, ANOVA. C: PbA inhibits choline efflux in a dose-dependent manner. Cells were preloaded with 50 nM [3H]choline chloride and 4.95 µM unlabeled choline chloride in uptake buffer at 37°C for 30 min, washed, and incubated in unlabeled uptake buffer in presence or absence of PbA for 10 min at 25°C. Aliquots of media were taken, and choline efflux was determined as in Fig. 2C. Data are expressed as means of 25 experiments performed in triplicate. D: inhibitory effects of PbA on choline efflux are reversible. Cells were preloaded with labeled choline chloride, rinsed, and exposed to 10 µM PbA for 10 min as in C. Aliquots of media were taken to determine choline efflux. In some samples, cells then were incubated in fresh medium for 30 min at 25°C without inhibitor (30 min wash), medium was changed again, and aliquots of media were taken after a 10-min incubation for labeled choline counting. Results were corrected to protein level. Data are expressed as means of 3 experiments performed in triplicate ± SE. *P = 0.08 vs. control (no wash); n = 3, t-test. E: effect of OCT-2 small interfering (si) RNA on choline efflux in MLE cells. MLE cells were transfected with FuGENE 6 alone (Control), FuGENE 6 plus OCT-2 siRNA, or FuGENE 6 plus scrambled RNA (scrRNA) for 24 h. Cells were then incubated with labeled choline chloride and washed, and aliquots (100 µl) of media were taken after 5-min incubation for analysis of choline efflux. Cells were also harvested for protein determination and immunoblotting for OCT-2. Data on effluxed choline normalized to protein are presented as means of 2 independent experiments performed in duplicate ± SD as a percentage of control values equal to 100% (graph). Inset: representative immunoblot for OCT-2 protein levels in transfected cells in presence or absence of siOCT-2 or scrRNA. Each lane contained 120 µg of protein lysate. F: Ad5 stimulation of choline efflux in MLE cells is blocked with PbA. Uninfected cells (control, left set of bars) or cells infected with Ad5 for 24 h (right set of bars) were incubated with labeled choline chloride and treated with 0, 5, or 10 µM PbA as described in C, and aliquots of medium were taken after 5 min of incubation. Data are expressed as a percent efflux of radiolabeled choline relative to control or Ad5-infected cells not exposed to PbA (set as 100%, n = 24, means ± SD). When compared with the control group, 10 µM PbA effectively reduced choline efflux regardless of Ad5 exposure.
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Phenoxybenzamine is a relatively specific inhibitor of OCT transporters (13). Phenoxybenzamine inhibited choline efflux in a dose-dependent manner with maximal effects achieved at 5 µM concentration (Fig. 6B). In contrast, cellular choline uptake was not significantly altered by phenoxybenzamine (Fig. 6C). To examine reversibility, cells were preloaded with radiolabeled choline, treated with phenoxybenzamine, and then exposed to fresh media without the inhibitor for 30 min. After washing, choline efflux in control and phenoxybenzamine-treated cells was not significantly different (Fig. 6D). Thus phenoxybenzamine selectively inhibits choline efflux in lung epithelia in a reversible manner.
As an alternative approach to assess the role of OCT-2 as a choline transporter, we employed genetic strategies. Cells were transfected with transfectant reagent alone or in the presence of siOCT-2 or control scrambled RNA for 24 h. Unlike our scrambled RNA control, OCT-2 siRNA decreased immunoreactive OCT-2 levels significantly (Fig. 6E, top). Moreover, OCT-2 siRNA inhibited choline efflux by 3040% compared with control cells or cells transfected with scrambled RNA (Fig. 6E). These data indicate that OCT-2, in part, contributes to choline efflux in lung epithelia. Because optimal inhibition of OCT-2 activity was achieved pharmacologically, we examined whether phenoxybenzamine might counteract adenoviral stimulation of choline efflux. Cells were exposed to Ad5 for 24 h, and choline efflux was measured as in Fig. 6B. Phenoxybenzamine decreased choline efflux in both Ad5-infected and uninfected cells with similar potency (Fig. 6F).
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DISCUSSION
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Our data reveal that mice infected with replication-competent Ad5 exhibited significantly reduced bronchoalveolar PC levels compared with control mice. Because adenoviruses alter PC substrate availability by increasing choline release from biomembranes, we pursued studies examining effects of these viruses on alveolar choline transport. Our studies uncover a new mechanism whereby human adenoviruses disrupt transport of choline in lung epithelia. Our studies identify for the first time that members of the OCT family are differentially expressed and partake in pulmonary choline transport. Specifically, OCT-2 is highly expressed in alveolar cells and catalyzes the transport of choline. Choline transport was bidirectional and pH dependent, all features of OCT-2 activity. Finally, human adenovirus coordinately inhibit choline uptake and stimulates its export unlike replication-defective (E1A and E1B deleted) virus, thereby depleting cells of choline. These deleterious effects of adenovirus were attenuated by pharmacological inhibition of OCT activity. The pathophysiological significance of these results is that in the setting of acute viral lung injury, adenoviruses might reduce the availability of choline-containing metabolites, such as PC, as observed in the present study, that are critical for maintenance of alveolar structure and function.
Choline is an essential endogenous substrate acquired primarily from the diet. Indeed, plasma levels of choline do not exceed 25 µM (20), with a Km for choline uptake by alveolar type II cells of
18 µM (7). Thus the transfer of choline into cells is a critical event for the biosynthesis of end products including acetylcholine, sphingomyelin, and PC. Prior electrophysiological studies indicate possibly several functional alveolar transport systems: a potential-dependent, hemicholinium-sensitive transporter and both Na+-dependent and Na+-independent carriers that exhibit pH sensitivity (7, 8, 17). In studies of membrane vesicles isolated from type II cell plasma membranes, choline uptake occurred basolaterally by an Na+-independent, HC-3-sensitive transporter accompanied by electroneutral choline/H+ exchange (30). The purpose of the present study, however, was not to recapitulate elegant electrophysiological studies characterizing these alveolar transport systems. Rather, in the process of investigating adenoviral pathogenesis on lung PC metabolism, we observed that wild-type replication-competent adenovirus significantly depleted cells of choline and of alveolar PC. Our goal was to examine the metabolic target(s) for adenovirus on choline transport and investigate the molecular basis whereby adenovirus might alter expression of these proteins.
Our observation that choline transport in MLE cells was bidirectional and pH sensitive and inhibited by a gamut of monoamine substrates and phenoxybenzamine led us to investigate OCT proteins in alveolar epithelia and whether their expression might be modulated by adenovirus. OCT members are highly expressed and functional in the brain, liver, and kidney, but little is known about their regulation in the lung (21). Among OCTs, only OCT-1 and OCT-2 transport choline, but with differing substrate affinities (42). OCTs provide facilitated transport of organic cations in a potential-dependent, electrogenic, and sodium-independent manner (4, 20, 21). Moreover, as seen elsewhere and in lung epithelia, OCTs provide facilitated transport of organic cations bidirectionally (4, 20, 21). Although OCT-1 and OCT-2 cDNAs are present in MLE cells, levels were likely very low, and steady-state OCT-2 mRNA was detectable only in primary murine type II epithelia. Furthermore, only OCT-2 protein was highly expressed in freshly isolated rodent primary type II cells, but expression waned with time in culture. Thus OCT-2 may be an in vivo marker of type II cells, and its loss of expression might signal conversion to a type I cell phenotype in vitro. These observations also suggest that there may be a regulation of OCT protein stability or at the pretranslational level. More importantly, it is likely that OCT-2 only partly contributes to alveolar choline transport and that a multitude of yet unidentified cationic systems exists. In support of this, OCT-2 siRNA only partly attenuated choline efflux, whereas pharmacological inhibition was much more effective (Fig. 6).
OCT-2 was also observed to be physiologically regulated by adenovirus. Wild-type, E1A-competent adenovirus decreased choline availability, in part, by increasing steady-state OCT-2 mRNA and protein expression. Interestingly, E1A/B-deficient adenovirus (Ad5-d312) did not inhibit choline uptake or trigger choline efflux compared with Ad5 virus. neither virus replicates in mouse cells, but wild-type adenovirus expresses all of the early-region proteins. Although speculative, it is possible that early-region proteins of Ad5 might transcriptionally activate the OCT-2 promoter or directly interact with regulatory domains within the transporter's primary structure. Adenoviral early-region proteins have diverse regulatory effects on expression of genes involved in cytolysis, cytokine and chemokine production, signaling events, and transcription factors (9). Furthermore, adenoviral early gene products such as E1A also interact within similar type II transmembrane proteins such as transporter associated with antigen processing (TAP-1) (31). Likewise, E1A expression may be critical for transcriptional regulation of the OCT gene in lung epithelia. Additional studies are required to determine whether early-region proteins directly modulate OCT-2 mRNA stability, mRNA synthesis, and protein stability or also directly affect its catalytic activity in response to viral infection.
Finally, our results showing upregulation of OCT expression by human adenovirus expands on the current understanding of viral pathogenesis within distal lung epithelia. Adenovirus regulates expression of various transport pumps, including aquaporin channels and ATP-binding cassette transporters (28). These results suggest that the virus can significantly alter fluid, electrolyte, and possibly lung lipid composition. Recent studies using respiratory syncytial virus reveal a novel mechanism whereby alveolar fluid clearance is impaired due to the ability of the virus to induce 5'-nucleotide release, resulting in altered sodium transport (5a). The present results do not investigate this mechanism for adenoviral infection, but it is reasonable to speculate that viral infections exploit a complex array of processes directed at disrupting several transport systems to increase pulmonary edema.
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GRANTS
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This study was supported by a Merit Review Award from the Office of Research & Development, Department of Veterans Affairs, and by National Heart, Lung, and Blood Institute RO1 Grants HL-55584, HL-68135, and HL-71040 (to R. K. Mallampalli).
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ACKNOWLEDGMENTS
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The authors acknowledge assistance from Satya Mathur for preparation of OCT-2 siRNA and for helpful discussions.
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FOOTNOTES
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Address for reprint requests and other correspondence: R. K. Mallampalli, Pulmonary & Critical Care Div., C-33K, GH, Depts. of Internal Medicine & Biochemistry, Univ. of Iowa College of Medicine, Iowa City, IA 52242 (E-mail: rama-mallampalli{at}uiowa.edu)
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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REFERENCES
|
---|
- Aberle SW, Aberle JH, Steininger C, Matthes-Martin S, Pracher E, and Popow-Kraupp T. Adenovirus DNA in serum of children hospitalized due to an acute respiratory adenovirus infection. J Infect Dis 187: 311314, 2003.[CrossRef][ISI][Medline]
- Arndt P, Volk C, Gorboulev V, Budiman T, Popp C, Ulzheimer-Teuber I, Akhoundova A, Koppatz S, Bamberg E, Nagel G, and Koepsell H. Interaction of cations, anions, and weak base quinine with rat renal cation transporter rOCT2 compared with rOCT1. Am J Physiol Renal Physiol 281: F454F468, 2001.[Abstract/Free Full Text]
- Berthiaume Y, Folkesson HG, and Matthay MA. Lung edema clearance: 20 years of progress: invited review: alveolar edema fluid clearance in the injured lung. J Appl Physiol 93: 22072213, 2002.[Abstract/Free Full Text]
- Burckhardt G and Wolff NA. Structure of renal organic anion and cation transporters. Am J Physiol Renal Physiol 278: F853F866, 2000.[Abstract/Free Full Text]
- Dada LA and Sznajder JI. Mechanisms of pulmonary edema clearance during acute hypoxemic respiratory failure: role of the Na,K-ATPase. Crit Care Med 31: S248S252, 2003.[CrossRef][ISI][Medline]
- Davis IC, Sullender WM, Hickman-Davis JM, Lindsey JR, and Matalon S. Nucleotide-mediated inhibition of alveolar fluid clearance in BALB/c mice after respiratory syncytial virus infection. Am J Physiol Lung Cell Mol Physiol 286: L112L120, 2004.[Abstract/Free Full Text]
- Dresser MJ, Leabman MK, and Giacomini KM. Transporters involved in the elimination of drugs in the kidney: organic anion transporters and organic cation transporters. J Pharm Sci 90: 397421, 2001.[CrossRef][ISI][Medline]
- Fisher AB, Chander A, Dodia C, Reicherter J, and Kleinzeller A. Choline transport by lung epithelium. Am J Respir Cell Mol Biol 1: 455462, 1989.[ISI][Medline]
- Fisher AB, Dodia C, Chander A, and Kleinzeller A. Transport of choline by plasma membrane vesicles from lung-derived epithelial cells. Am J Physiol Cell Physiol 263: C1250C1257, 1992.[Abstract/Free Full Text]
- Friedman JM and Horwitz MS. Inhibition of tumor necrosis factor alpha-induced NF-kappa B activation by the adenovirus E310.4/145K complex. J Virol 76: 55155521, 2002.[Abstract/Free Full Text]
- Friedrich A, George RL, Bridges CC, Prasad PD, and Ganapathy V. Transport of choline and its relationship to the expression of the organic cation transporters in a rat brain microvessel endothelial cell line (RBE4). Biochim Biophys Acta 1512: 299307, 2001.[ISI][Medline]
- Fyfe GK, Kemp PJ, Cragoe EJ Jr, and Olver RE. Conductive cation transport in apical membrane vesicles prepared from fetal lung. Biochim Biophys Acta 1224: 355364, 1994.[CrossRef][ISI][Medline]
- Gorboulev V, Volk C, Arndt P, Akhoundova A, and Koepsell H. Selectivity of the polyspecific cation transporter rOCT1 is changed by mutation of aspartate 475 to glutamate. Mol Pharmacol 56: 12541261, 1999.[Abstract/Free Full Text]
- Hayer-Zillgen M, Bruss M, and Bonisch H. Expression and pharmacological profile of the human organic cation transporters hOCT1, hOCT2 and hOCT3. Br J Pharmacol 136: 829836, 2002.[Abstract/Free Full Text]
- Jonker JW, Wagenaar E, Mol CA, Buitelaar M, Koepsell H, Smit JW, and Schinkel AH. Reduced hepatic uptake and intestinal excretion of organic cations in mice with a targeted disruption of the organic cation transporter 1 (Oct1 [Slc22a1]) gene. Mol Cell Biol 21: 54715477, 2001.[Abstract/Free Full Text]
- Jonker JW, Wagenaar E, Van Eijl S, and Schinkel AH. Deficiency in the organic cation transporters 1 and 2 (Oct1/Oct2 [Slc22a1/Slc22a2]) in mice abolishes renal secretion of organic cations. Mol Cell Biol 23: 79027908, 2003.[Abstract/Free Full Text]
- Kerb R, Brinkmann U, Chatskaia N, Gorbunov D, Gorboulev V, Mornhinweg E, Keil A, Eichelbaum M, and Koepsell H. Identification of genetic variations of the human organic cation transporter hOCT1 and their functional consequences. Pharmacogenetics 12: 591595, 2002.[CrossRef][ISI][Medline]
- Kleinzeller A, Dodia C, Chander A, and Fisher AB. Na+-dependent and Na+-independent systems of choline transport by plasma membrane vesicles of A549 cell line. Am J Physiol Cell Physiol 267: C1279C1287, 1994.[Abstract/Free Full Text]
- Klinger JR, Sanchez MP, Curtin LA, Durkin M, and Matyas B. Multiple cases of life-threatening adenovirus pneumonia in a mental health care center. Am J Respir Crit Care Med 157: 645649, 1998.[ISI][Medline]
- Koc H, Mar MH, Ranasinghe A, Swenberg JA, and Zeisel SH. Quantitation of choline and its metabolites in tissues and foods by liquid chromatography/electrospray ionization-isotope dilution mass spectrometry. Anal Chem 74: 47344740, 2002.[CrossRef][ISI][Medline]
- Koepsell H. Organic cation transporters in intestine, kidney, liver, and brain. Annu Rev Physiol 60: 243266, 1998.[CrossRef][ISI][Medline]
- Koepsell H and Endou H. The SLC22 drug transporter family. Pflügers Arch 447: 666676, 2004.[CrossRef][ISI][Medline]
- Leggiadro RJ and Darras BT. Viral and bacterial pathogens of suspected sepsis in young infants. Pediatr Infect Dis J 2: 287289, 1983.[ISI]
- Longo CA, Tyler D, and Mallampalli RK. Sphingomyelin metabolism is developmentally regulated in rat lung. Am J Respir Cell Mol Biol 16: 605612, 1997.[Abstract]
- Look DC, Roswit WT, Frick AG, Gris-Alevy Y, Dickhaus DM, Walter MJ, and Holtzman MJ. Direct suppression of Stat1 function during adenoviral infection. Immunity 9: 871880, 1998.[ISI][Medline]
- McCoy DM, Salome RG, Kusner DJ, Iyar SS, and Mallampalli RK. Identification of sex-specific differences in surfactant synthesis in rat lung. Pediatr Res 46: 722730, 1999.[Abstract]
- Meyer-Wentrup F, Karbach U, Gorboulev V, Arndt P, and Koepsell H. Membrane localization of the electrogenic cation transporter rOCT1 in rat liver. Biochem Biophys Res Commun 248: 673678, 1998.[CrossRef][ISI][Medline]
- Mistchenko AS, Robaldo JF, Rosman FC, Koch ER, and Kajon AE. Fatal adenovirus infection associated with new genome type. J Med Virol 54: 233236, 1998.[CrossRef][ISI][Medline]
- Momburg F and Hengel H. Corking the bottleneck: the transporter associated with antigen processing as a target for immune subversion by viruses. Curr Top Microbiol Immunol 269: 5774, 2002.[ISI][Medline]
- Motohashi H, Sakurai Y, Saito H, Masuda S, Urakami Y, Goto M, Fukatsu A, Ogawa O, and Inui K. Gene expression levels and immunolocalization of organic ion transporters in the human kidney. J Am Soc Nephrol 13: 866874, 2002.[Abstract/Free Full Text]
- Oelberg DG and Xu F. Conductive choline transport by alveolar epithelial plasma membrane vesicles. Mol Genet Metab 65: 220228, 1998.[CrossRef][ISI][Medline]
- Proffitt JA and Blair GE. The MHC-encoded TAP1/LMP2 bidirectional promoter is down-regulated in highly oncogenic adenovirus type 12 transformed cells. FEBS Lett 400: 141144, 1997.[CrossRef][ISI][Medline]
- Rice WR, Conkright JJ, Na CL, Ikegami M, Shannon JM, and Weaver TE. Maintenance of the mouse type II cell phenotype in vitro. Am J Physiol Lung Cell Mol Physiol 283: L256L264, 2002.[Abstract/Free Full Text]
- Ridge KM, Olivera WG, Saldias F, Azzam Z, Horowitz S, Rutschman DH, Dumasius V, Factor P, and Sznajder JI. Alveolar type 1 cells express the alpha2 Na,K-ATPase, which contributes to lung liquid clearance. Circ Res 92: 453460, 2003.[Abstract/Free Full Text]
- Rosenlew M, Stenvik M, Roivainen M, Jarvenpaa AL, and Hovi T. A population-based prospective survey of newborn infants with suspected systemic infection: occurrence of sporadic enterovirus and adenovirus infections. J Clin Virol 12: 211219, 1999.[CrossRef][ISI][Medline]
- Ryan MA, Gray GC, Smith B, McKeehan JA, Hawksworth AW, and Malasig MD. Large epidemic of respiratory illness due to adenovirus types 7 and 3 in healthy young adults. Clin Infect Dis 34: 577582, 2002.[CrossRef][ISI][Medline]
- Salome RG, McCoy DM, Ryan AJ, and Mallampalli RK. Effects of intratracheal instillation of TNF-
on surfactant metabolism. J Appl Physiol 88: 1016, 2000.[Abstract/Free Full Text]
- Seth P. Adenovirus-dependent release of choline from plasma membrane vesicles at an acidic pH is mediated by the penton base protein. J Virol 68: 12041206, 1994.[Abstract]
- Shiratori Y, Houweling M, Zha X, and Tabas I. Stimulation of CTP:phosphocholine cytidylyltransferase by free cholesterol loading of macrophages involves signaling through protein dephosphorylation. J Biol Chem 270: 2989429903, 1995.[Abstract/Free Full Text]
- Shu Y, Bello CL, Mangravite LM, Feng B, and Giacomini KM. Functional characteristics and steroid hormone-mediated regulation of an organic cation transporter in Madin-Darby canine kidney cells. J Pharmacol Exp Ther 299: 392398, 2001.[Abstract/Free Full Text]
- Sinclair CJ, Chi KD, Subramanian V, Ward KL, and Green RM. Functional expression of a high affinity mammalian hepatic choline/organic cation transporter. J Lipid Res 41: 18411848, 2000.[Abstract/Free Full Text]
- Sugita M, Ferraro P, Dagenais A, Clermont ME, Barbry P, Michel RP, and Berthiaume Y. Alveolar liquid clearance and sodium channel expression are decreased in transplanted canine lungs. Am J Respir Crit Care Med 167: 14401450, 2003.[Abstract/Free Full Text]
- Sweet DH, Miller DS, and Pritchard JB. Ventricular choline transport: a role for organic cation transporter 2 expressed in choroid plexus. J Biol Chem 276: 4161141619, 2001.[Abstract/Free Full Text]
- Sweet DH and Pritchard JB. rOCT2 is a basolateral potential-driven carrier, not an organic cation/proton exchanger. Am J Physiol Renal Physiol 277: F890F898, 1999.[Abstract/Free Full Text]
- Towne JE, Harrod KS, Krane CM, and Menon AG. Decreased expression of aquaporin (AQP)1 and AQP5 in mouse lung after acute viral infection. Am J Respir Cell Mol Biol 22: 3444, 2000.[Abstract/Free Full Text]
- Urakami Y, Okuda M, Masuda S, Saito H, and Inui KI. Functional characteristics and membrane localization of rat multispecific organic cation transporters, OCT1 and OCT2, mediating tubular secretion of cationic drugs. J Pharmacol Exp Ther 287: 800805, 1998.[Abstract/Free Full Text]
- Van Montfoort JE, Hagenbuch B, Groothuis GM, Koepsell H, Meier PJ, and Meijer DK. Drug uptake systems in liver and kidney. Curr Drug Metab 4: 185211, 2003.[ISI][Medline]
- Zhou J, Ryan AJ, Medh J, and Mallampalli RK. Oxidized lipoproteins inhibit surfactant phosphatidylcholine synthesis via calpain mediated cleavage of CTP:phosphocholine cytidylyltransferase. J Biol Chem 278: 3703237040, 2003.[Abstract/Free Full Text]
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