Urea transport in MDCK cells that are stably transfected with UT-A1

Otto Fröhlich,1 Janet D. Klein,2 Pauline M. Smith,1 Jeff M. Sands,1,2 and Robert B. Gunn1

1Department of Physiology and 2Renal Division, Department of Medicine, Emory University School of Medicine, Atlanta, Georgia 30322

Submitted 29 December 2003 ; accepted in final form 18 January 2004


    ABSTRACT
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Progress in understanding the cell biology of urea transporter proteins has been hampered by the lack of an appropriate cell culture system. The goal of this study was to create a polarized epithelial cell line that stably expresses the largest of the rat renal urea transporter UT-A isoforms, UT-A1. The gene for UT-A1 was cloned into pcDNA5/FRT and transfected into Madin-Darby canine kidney (MDCK) cells with an integrated Flp recombination target site. The cells from a single clone were grown to confluence on collagen-coated membranes until the resistance was >1,500 {Omega}·cm2. Transepithelial [14C]urea fluxes were measured at 37°C in a HCO3/CO2 buffer, pH 7.4, with 5 mM urea. The baseline fluxes were not different between unstimulated UT-A1-transfected MDCK cells and nontransfected or sham-transfected MDCK cells. However, only in the UT-A1-transfected cells was UT-A1 protein expressed (as measured by Western blot analysis) and urea transport stimulated by forskolin or arginine vasopressin. Forskolin and arginine vasopressin also increased the phosphorylation of UT-A1. Thionicotinamide, dimethylurea, and phloretin inhibited the forskolin-stimulated [14C]urea fluxes in the UT-A1-transfected MDCK cells. These characteristics mimic those seen in rat terminal inner medullary collecting ducts. This new polarized epithelial cell line stably expresses UT-A1 and reproduces several of the physiological responses observed in rat terminal inner medullary collecting ducts.

urea transporter-A1; arginine vasopressin; collecting duct; Madin-Darby canine kidney cells


DURING THE PAST DECADE, substantial progress has been made in understanding the regulation of urea transport proteins in kidney. Major advances include the cloning of two urea transporter genes and several cDNA isoforms as well as the creation of polyclonal antibodies to different protein isoforms (reviewed in Ref. 27). The UT-A family of urea transporters currently consists of five isoforms plus three cDNA variants that differ in the 3'-untranslated region. UT-A1, UT-A2, UT-A3, and UT-A4 are expressed in kidney, but UT-A5 is found only in the testes (6). UT-A1 is the largest UT-A protein, with its 97- and 117-kDa glycosylated forms. It is expressed in the apical membrane of the inner medullary collecting duct (IMCD) in humans (17) and rodents (23, 30). Stimulation of protein kinase A increases urea transport in the rat terminal IMCD (28, 31), increases the phosphorylation of UT-A1 in IMCD suspensions (32), and increases urea flux in Xenopus oocytes that heterologously express human or rodent UT-A1 (7, 24, 30).

The primary method for investigating the rapid regulation of urea transport has been perfusion of rat IMCDs. This method provides physiologically relevant functional data, although it cannot determine which urea transporter isoform is responsible for a specific functional effect, because both UT-A1 and UT-A3 are expressed in this nephron segment. Progress in understanding the cell biology of urea transporters and the functions of the separate isoforms has been hampered by the lack of an appropriate cell culture system. The goal of the present study was to create a polarized epithelial cell line that stably expresses the UT-A1 urea transporter and reproduces many of the functional properties of urea transport in the IMCD.

In early studies of type I, high-resistance Madin-Darby canine kidney (MDCK) cells, it was shown that they responded to arginine vasopressin (AVP, antidiuretic hormone) by increasing the adenylyl cyclase activity, cAMP levels, and the synthesis of prostaglandins (reviewed in Ref. 8). Addition of apical AVP (110 nM = 50 mU/ml) also increased the rate of tracer Na+ efflux from the cells, but the effect was smaller and delayed compared with the increased rate of efflux seen with 1 mM cAMP (8).


    METHODS
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
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MDCK cells, the kind gift of Dr. James Schafer (University of Alabama at Birmingham), were originally selected in Rossier's laboratory (University of Lausanne) for low epithelial sodium channel expression. They were then used and passaged in Rotin's laboratory (University of Toronto) and then in Schafer's laboratory (19) and are a type I, high-resistance cell line (2, 25). These cells were cultured in T-75 Corning Costar flasks (Corning, Acton, MA) in a humidified incubator at 37°C in the presence of 5% CO2. The medium was DMEM (GIBCO/Invitrogen Life Technologies, Carlsbad, CA) supplemented with 25 mM HEPES (pH 7.4), 100 IU/ml each of penicillin and streptomycin, and 10% fetal calf serum (Atlanta Biological). The cells were fed every other day, trypsinized (0.05% trypsin; GIBCO/Invitrogen) when confluent, and split.

A new cell line of MDCK cells was constructed by using the Flp-In system (Invitrogen). This system creates isogenic cell lines with one or more Flp recombination target (FRT) sites (5, 29). To create the new MDCK-FRT cell line, MDCK cells were first stably transfected with pFRT/lacZeo by using Lipofectamine 2000 (Invitrogen) and were selected for 10 days in 100 µg/ml Zeocin. Single cell colonies were grown, their {beta}-galactosidase activity was measured, and Southern blot analysis was performed to determine the number of FRT sites inserted into the genome. Since the MDCK-UT-A1 cells are sensitive to Zeocin, all of the FRT sites have UT-A1 inserted into them by homologous recombination.

A clonally selected MDCK-FRT cell line was then cotransfected with pOG44, a vector for transiently expressing the Flp recombinase, and with pcDNA5/FRT/UT-A1, an expression vector that contains the coding region for UT-A1 and possesses an FRT site for homologous recombination. After insertion of the UT-A1 gene into the FRT site, UT-A1 transcription was driven by the human cytomegalovirus immediate-early enhancer/promoter. The latter vector also contains the hygromycin resistance gene under the control of the SV40 promoter to permit selection for recombinant clones in 800 µg/ml hygromycin. The homologous recombination inactivates the lacZ-Zeocin fusion gene. These cells were then grown and passaged in T-75 flasks by using 500 µg/ml hygromycin in DMEM to maintain selection. The final MDCK-UT-A1 cells, which express UT-A1 by functional assay and Western blot analysis, are sensitive to Zeocin, resistant to hygromycin, and lack {beta}-galactosidase activity. The cells were grown and passaged in DMEM containing HEPES and bicarbonate buffers by using standard techniques.

Collagen-coated Costar Transwell inserts (1 cm2 growth surface area; Corning) were used to grow an epithelial layer of MDCK cells. After 1-h incubation of the Transwells in DMEM at 37°C, a suspension of selected, trypsinized cells was prepared, and 7.5 x 104 cells were loaded onto each Transwell and fed after 1 h without hygromycin. These cells grew to confluence over 5–7 days. The transmembrane resistance was measured daily by using an epithelial resistance meter (EVOMX-G; World Precision Instruments, Sarasota, FL). Inserts in which there were not orderly increases in the transepithelial electrical resistance from <0.1 k{Omega} to >1 k{Omega} were discarded. We used only membranes with 1.5 k{Omega} or higher resistance for flux measurements. We transferred the membranes from the CO2 incubator to the flux plate in the following manner to minimize changes in ionic conditions. The urea flux medium contained Hanks' balanced salt solution with bicarbonate (HBBS; GIBCO/Invitrogen) supplemented with 20 mM HEPES from a 1-M stock (GIBCO/Invitrogen). Flux medium made of HBBS-HEPES containing 5 mM urea was aliquoted (1.5 ml) into the wells of a 12-well plate and placed with the lid open for ~1 h in the humidified tissue culture incubator (Pco2 = 40 mmHg, 37°C). We replaced the lid before we moved the covered plates to the experimental bench, where we temporarily transferred them to a 37°C water bath and then placed them on top of a thermostated aluminum block.

Urea flux measurements. Immediately before initiating the flux measurements, we removed four epithelial membrane inserts (Costar) from their culture plates and placed them into empty wells of a 12-well plate. We carefully removed the 500–600 µl of culture medium over each epithelial layer and added 400 µl of prewarmed flux medium containing 0.4 µCi of [14C]urea. The four inserts were then transferred to the first row of wells, each containing the 1.5 ml of flux medium to initiate the transepithelial flux. We removed the lid only to add the inserts and then three more times during the flux as the four inserts were moved at designated times from row to row, ending one and initiating the next flux period (Fig. 1). We always measured the [14C]urea flux from the cis solution (insert or apical membrane bathing solution or upper solution) to the trans solution (well or basolateral membrane bathing solution or the lower solution) because it was difficult to rapidly and carefully remove the upper solution without damaging the cells growing on the filter that forms the base of the insert. Forskolin (10 mM stock in DMSO), AVP (10–4 M stock in H2O), phloretin (100 mM stock in ethanol), dimethylurea, and thionicotinamide (Sigma) were certified grade and were added to the trans solution.



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Fig. 1. Schematics of flux experiments. The insert containing the cultured Madin-Darby canine kidney (MDCK) cells that form a confluent, high-resistance epithelial membrane is moved at each time point from 1 row of a 12-well plate to the next. Here the insert is located in the well of row 2. The insert contains the radioactive cis solution; the bottom well contains the trans solution into which the [14C]urea is transported during each flux period.

 
Technical control studies. Before placing the flux plates onto the thermostated aluminum block, we added a small amount of water onto the block to enhance heat transfer from the block to the culture dish. The difference in temperature was <0.5°C between the circulated water and the efflux medium in the culture dish wells. Furthermore, the hydrostatic pressure difference due to unequal levels within the (upper) insert and the (lower) well was small (<3 mmH2O) compared with the total osmotic pressure (19 mmHg/mM x 150 mM x 13.6 mmH2O/mmHg = 38,700 mmH2O) of the solutions. In control experiments we saw no difference in the flux when we made the levels different by 8 mm, thus excluding significant solute drag of urea with bulk water transport. There was always residual fluid in the upper chamber after the culture medium was removed, which diluted the isotope in the [14C]urea flux solution that was added. We therefore sampled the upper chamber for specific activity at the beginning and end of the flux measurements. The dilution of the specific activity from the stock was appropriate for a residual of ~20 µl in the upper chamber above the epithelial layer. There was fluid attached to the lower surface when the insert was moved from well to well. The residual that was inadvertently carried over from well to well on the underside of the insert during the flux measurements was minimized by dipping the insert three times and tipping it at an angle as it was removed and transferred to the next well. The carried-over counts did not alter the calculated flux more than a few percent, since in most cases the amount carried forward from the last well was not too different from the amount carried forward to the next well. Having consistent residual volumes transferred was a matter of practice and of consistency of tilting and dipping of the insert. We took all of the lower well solution to be counted. In control experiments, there was 27 µl residual in the well from a total of 1,500 µl. Triplicate 10-µl samples of the upper insert solution supplemented with 1.5 ml nonradioactive flux solution, to give equal volumes, were each counted with 3 ml of OptiFluor (Packard) in a Packard Tricarb liquid scintillation counter. Each time period was an independent flux measurement. The total counts transferred across the epithelial layer into the well divided by the specific activity of the upper solution and the time interval was the calculated flux.

A near-saturated solution of thionicotinamide (5 mM) was prepared in HBBS-HEPES. Thionicotinamide color quenches the [14C]urea counting rate when present in the scintillation fluid. We corrected the counts (using a quench curve derived from data obtained at a fixed [14C]urea amount and variable thionicotinamide concentrations) when thionicotinamide was present in the trans well.

During a long series of flux measurements, the specific activity in the cis (apical) compartment decreases. We calculated this by using the total counts that were transported to the preceding trans compartments, and we used this corrected specific activity to calculate the flux. The validity of this calculation was determined by directly measuring the specific activity of the cis compartment at the beginning and end of the flux measurements. When necessary, we therefore calculated the change in specific activity by interpolation between initial and final measurements and corrected the flux calculations accordingly.

Western blot and phosphorylation methods. We radiolabeled cells using our previously published methods (13, 14). Briefly, we washed confluent cell layers with phosphate-free DMEM and then incubated them with 1 ml of phosphate-free DMEM containing 0.1 mCi/ml of [32P]orthophosphate for 3 h, 37°C, 5% CO2, and 100% humidity. At the end of the 3-h loading period, inhibitors or activators were added for further incubation. We then washed and solubilized the cells in RIPA buffer. All cellular material was collected. Each phosphorylation immunoprecipitation sample contained the contents of two wells from a six-well plate. We sheared the cells in the samples with a 26-gauge needle, centrifuged the samples for 15 min at 14,000 g, and removed the top 80% of each supernatant fraction to a fresh tube containing 10 µl of antibody. We removed an additional 50 µl, which we added to an equal volume of Laemmli sample buffer and boiled before using it as a preimmunoprecipitation control sample in Western blots.

For immunoprecipitation, we incubated samples with UT-A1 antibody at 4°C overnight with gentle agitation. Protein A-agarose beads (25 µl) were added, and cold incubation continued for a further 2 h. We pelleted the beads in a microcentrifuge and then washed the pellet seven times with RIPA buffer. We verified the completeness of the washing protocol by counting all of the washes as well as the supernatant and sample. On the basis of the counts present on the protein A-agarose beads, we added an amount of Laemmli buffer to the beads and boiled the samples for 1–3 min.

For electrophoretic analysis, proteins were size separated by SDS-PAGE on Laemmli gels and then either stained with Coomassie blue and dried for autoradiography or electroblotted to polyvinylidene difluoride membranes for Western blot analysis as described previously (12–15, 32). Western blots were incubated with our antibody to the COOH terminus of UT-A1 (5% milk, Tris-buffered saline, 0.05% Tween-20) overnight at 4°C (20). The secondary antibody we used depended on the method of analysis. For enhanced chemiluminescence detection, we further incubated the blot with horseradish peroxidase-linked goat anti-rabbit IgG at a dilution of 1:5,000 (2 h, room temperature) and then washed and analyzed it. For infrared detection, we incubated the blots with goat anti-rabbit IgG fluorescently labeled with Alexa 680 (1:4,000, 2 h, room temperature), and then we washed and visualized the blot by using the LI-COR Odyssey gel scanning system (LI-COR Biotechnology, Lincoln, NE). We stained gels in parallel with Coomassie blue to verify the uniformity of gel loading.

UT-A1 protein was immunoprecipitated from equal amounts of the whole cell lysates by using our previously described method (13, 14, 32). Proteins were size separated on two identical SDS-polyacrylamide gels containing an equal portion of the total immunoprecipitated protein per lane. The proteins on one gel were transferred to a polyvinylidene difluoride membrane, and the amount of immunoprecipitated UT-A1 was assayed by Western blot. The other gel was dried, and 32P incorporation into UT-A1 was analyzed by autoradiography.


    RESULTS
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Expression of UT-A1 protein in MDCK-UT-A1 cells. We tested whether the newly derived cell line expresses the desired urea transporter protein by Western blot analysis. Figure 2 demonstrates that these cells express significant amounts of newly expressed UT-A1 protein. The protein is predominantly present as the lower-glycosylation form of 97 kDa.



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Fig. 2. Western blot analysis of urea transporter UT-A1 protein expression in MDCK-Flp recombinant target (FRT) cells without UT-A1 and MDCK-UT-A1 cells with UT-A1. The glycosylated 97-kDa UT-A1 protein is stably expressed in transfected cells. Either 5 or 15 µg of total protein was loaded on the gel before electrophoresis.

 
Stimulation of UT-Al-mediated urea fluxes by AVP. We tested the stimulation of trans-epithelial urea flux by six concentrations (0–10–8 M) of AVP on six separate epithelial membranes (Fig. 3). After six control flux periods, AVP was added to the basolateral side (well) at the nominal concentration stated in Fig. 3. After stimulation for 33 min, 5 mM thionicotinamide was present during the three final time periods. AVP stimulation is apparent at 10–10 M and increases to a maximum at 3 x 10–9 and 10–8 M. The membrane stimulated by 10–8 M AVP had a higher control flux and an equally higher flux in the presence of thionicotinamide. Thus the absolute increase in the flux was not different from that in 3 x 10–9 M AVP. The average flux without AVP was 1.8 ± 0.2 nmol·min–1·cm–2 (all values are means ± SD). The average flux without AVP and with 5 mM thionicotinamide was 1.6 ± 0.1 nmol·min–1·cm–2.



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Fig. 3. Effect of arginine vasopressin (AVP) on urea transport. The [14C]urea flux across 6 separate membranes was measured for 6 control time periods. At ~18 min, different concentrations of AVP were added to the basolateral wells for the remaining flux periods. At ~52 min, 5 mM thionicotinamide was added to the basolateral wells to inhibit the UT-A1-mediated urea flux. The nominal AVP concentrations were zero (x), showing that the control flux did not change in the absence of added AVP at 10–10 M ({triangleup}), 3 x 10–10 M ({square}), 10–9 M ({circ}), 3 x 10–9 M ({lozenge}), and 10–8 M (+). Stimulation was evident at a physiological 10–10 M AVP and was maximal at 3 x 10–9 M. All of the AVP-stimulated flux was thionicotinamide inhibitable.

 
Phosphorylation of UT-A1 by AVP. The increase in urea flux in response to AVP is mimicked by the increase in UT-A1 phosphorylation. Figure 4 shows both uniform protein and increased phosphorylation of UT-A1 (arrows) in cells treated with 10–8 M AVP for 15 min.



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Fig. 4. Stimulation of UT-A1 phosphorylation by AVP. MDCK-UT-A1 cells were labeled with [32P]orthophosphate for 3 h and then treated with 10–8 M AVP for 15 min. The cells were thoroughly washed, solubilized, and immunoprecipitated and were run on a gel. Detected radiophosphorylated UT-A1 (left) and total UT-A1 protein (right) in samples from separate cell cultures without and with treatment with AVP are shown. The double-headed arrow is at 97 kDa.

 
Stimulation of UT-A1-mediated urea fluxes by forskolin. In four parallel flux experiments, we compared the stimulation by basolateral forskolin (10 µM) for 30–60 min and subsequent inhibition by basolateral thionicotinamide (5 mM) of urea transport in MDCK-FRT cells transfected with the FRT site(s) alone (control) or with UT-A1 inserted in those sites (Fig. 5). In the control cells, forskolin did not stimulate the [14C]urea flux significantly and thionicotinamide essentially did not inhibit the flux. This suggests that the native MDCK cells have little, if any, intrinsic thionicotinamide-inhibitable urea transport and that it is not stimulated by forskolin. Without forskolin treatment, UT-A1-transfected cells also exhibited a low urea flux and no significant inhibition by thionicotinamide. In contrast, when the UT-A1-transfected cells were exposed to 10 µM forskolin, transepithelial urea fluxes were stimulated two- to threefold (and sometimes as much as fourfold) over the course of 30–60 min. This stimulated urea flux is nearly completely inhibited by thionicotinamide, indicating that it is mediated by UT-A1.



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Fig. 5. Activation of urea flux by forskolin. We measured sequential [14C]urea fluxes on 4 epithelial membranes in parallel. After 4 control flux periods, cells were treated (closed symbols) or not (open symbols) with 10 µM forskolin. MDCK-FRT cells transfected with the Flp site alone and treated with 10 µM forskolin (control cells, diamonds) were not activated. Cells transfected with UT-A1 but not treated with forskolin (triangles) also did not change their flux rate. Cells transfected with UT-A1 and treated with forskolin until 73 min (squares) or until 50 min (circles) had increased urea fluxes up to 3-fold. Two controls and one treated membrane were finally treated with 5 mM thionicotinamide (shaded symbols), which inhibited all of the increased flux. The forskolin-stimulated flux remained elevated for up to 30 min after forskolin removal. The membranes shown by the squares and circles demonstrate the range of differences in stimulation by forskolin.

 
Phosphorylation of UT-A1 by forskolin. A 97-kDa UT-A1 protein is expressed in the stably transfected MDCK-UT-A1 cells (Fig. 2). This 97-kDa UT-A1 protein is basally phosphorylated in unstimulated MDCK-UT-A1 cells, but phosphorylation is increased 2- to 10-fold within 2 min of stimulation with 10 µM forskolin. This is shown in Fig. 6, where, despite equal amounts of immunoprecipitated UT-A1 protein, the phosphorylation level was substantially increased within 2 min of forskolin treatment.



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Fig. 6. Stimulation of UT-A1 phosphorylation by forskolin in immunoprecipitated UT-A1. Cells were incubated with [32P]orthophosphate for 3 h and then treated with forskolin, and the UT-A1 was immunoprecipitated. Left: treatment by forskolin for up to 5 min did not alter the total amount of UT-A1 protein in the cells. (Adventitious bands of IgG are also shown.) Right: treatment by forskolin for 2.5 or 5 min greatly increased the phosphorylation of UT-A1.

 
Persistence and reversal of forskolin activation. To determine how rapidly the activation by forskolin was reversed by its withdrawal and how long it would persist in the presence of forskolin, we measured the time course of the activation of the [14C]urea flux in five inserts (Fig. 7). In the four inserts where 10 µM forskolin was applied, the flux was activated over ~20–30 min and then appeared to reach a plateau value. When after 42 min of stimulation the forskolin was removed from two of the inserts, the flux in the absence of forskolin remained stimulated for ~20 min and then declined ~40% over the next 35 min before 5 mM thionicotinamide was included on the basolateral side to inhibit the flux. In the continued presence of forskolin, the urea flux remained stimulated for a significantly longer period and then decayed more slowly. The control fluxes in the absence of forskolin were nearly constant (average 2.6 ± 0.3 nmol·min–1·cm–2) over the same time period.



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Fig. 7. Persistence of forskolin stimulation. We made sequential [14C]urea flux measurements in 5 membranes of MDCK-UT-A1 cells. Four membranes were treated with 10 µM forskolin for 20–40 min, and one membrane was not treated with forskolin (triangles). After reaching a plateau, the activated flux remained high after forskolin was removed from two membranes (circles). It then declined 40% over 0.5 h. At the end, the stimulated fluxes were inhibited by 5 mM thionicotinamide for 4 flux periods.

 
Concentration dependence of thionicotinamide inhibition on the basolateral surface. We tested the concentration dependence and reversibility of thionicotinamide inhibition by measuring the forskolin-simulated [14C]urea flux on four membranes with either increasing or decreasing concentrations of thionicotinamide in the bottom well solution (Fig. 8). There were no significant differences in the fluxes with raising or lowering the thionicotinamide concentration. The data were fit to the equation: VTN = Vinf + (V0 Vinf)/(1+ [TN]/Ki), where Vinf is the extrapolated flux at infinite thionicotinamide concentration ([TN]), V0 is the flux at zero thionicotinamide concentration, and VTN is the flux at a given thionicotinamide concentration. Ki, the apparent inhibitor constant of thionicotinamide on the basolateral membrane, was 1.2 mM, V0 was 10.8 nmol·min–1·cm–2, and Vinf was 2.4 nmol·min–1·cm–2.



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Fig. 8. Concentration dependence and reversibility of thionicotinamide (TN) inhibition from the basolateral solution. After forskolin stimulation of the [14C]urea flux, thionicotinamide was added in increasing concentrations ({triangleup}) to the basolateral solution or treated first with 5 mM thionicotinamide and then decreasing concentrations ({circ}) on the basolateral side. The inhibition was reversible, with an apparent Ki of 1.2 mM. Only the forskolin-activated flux was inhibited.

 
Inhibition of UT-A1 [14C]urea fluxes by basolateral dimethylurea. Dimethylurea inhibited urea fluxes to the same extent as thionicotinamide (Fig. 9). In preliminary experiments, we found that 300 mM dimethylurea rapidly inhibited the flux to the maximum extent but that the flux needed >10 min to be reversed after the dimethylurea solution was removed from the bottom well. However, after treatment with <20 mM dimethylurea the flux was rapidly and fully reversible. Figure 9 shows a fit to the data that yielded an apparent inhibitor constant for dimethylurea of 4.9 mM. Dimethylurea appears to be a relatively potent, complete, and reversible inhibitor of the UT-A1-mediated flux.



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Fig. 9. Concentration dependence of dimethylurea (DMU) inhibition from the basolateral solution. The forskolin-stimulated [14C]urea flux was reversibly (not shown) and completely inhibited by dimethylurea with an apparent Ki of 4.9 mM. The scatter in the data along the ordinate shows the variability of forskolin stimulation in different membranes. The lack of scatter of the values at 50 mM reflects the relative constancy of the unstimulated as well as maximally inhibited fluxes in the different membranes.

 
Inhibition of UT-A1-mediated [14C]urea fluxes by basolateral phloretin. Phloretin also inhibits the urea flux to the same extent as thionicotinamide and dimethylurea (Fig. 10). About 300 µM phloretin is sufficient to achieve near-complete inhibition.



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Fig. 10. Concentration dependence of phloretin inhibition. The forskolin-stimulated [14C]urea transmembrane flux was maximally inhibited by 300 µM phloretin added to the basolateral solution.

 

    DISCUSSION
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The major result of the present study is that we have created a stably transfected MDCK-UT-A1 cell line. We chose MDCK cells as parent cells because they are a model for distal nephron epithelium and are most likely to possess the same sets of signaling pathways as the distal nephron in vivo (21). MDCK cells can form a high-resistance epithelium (25) with low permeabilities for nonelectrolytes and water (18, 26). The value reported by Lavelle et al. (18) for the urea permeability of type I MDCK cells (7.3 x 10–6 cm/s) is the same as what we measured: our control fluxes in the range of 1.6–2.4 nmol·min–1·cm–2 (Figs. 3, 5, and 7) correspond to 5–8 x 10–6 cm/s.

Our Western blot analysis of urea transporter protein showed no evidence for intrinsic urea transporters in the parental untransfected MDCK cells (Fig. 2). A negative Western blot could be obtained if the antibody, which was raised against a COOH-terminal peptide from rat UT-A1, does not recognize the corresponding canine peptide sequence. However, our flux experiments in untransfected cells also showed no change in basal transport after treatment with AVP or forskolin, both of which produced a strong stimulation in transfected cells. Since expression of UT-A1 without hormonal activation did not increase the [14C]urea flux above the native flux rate, all of the hormonal activation can be ascribed to the expressed rat UT-A1. It therefore appears that the MDCK cells did not express a native canine urea transporter or, if they did, that it did not react with the antibody and was not activated by forskolin or AVP.

The MDCK-UT-A1 cells expressed significant amounts of UT-A1 protein as shown by Western blot analysis (Fig. 2), and our transport experiments showed that it was functionally active. In the renal tubule in vivo, the UT-A1 protein was present in two different glycosylation states, with apparent molecular weights of 97 kDa and 117 kDa (1), which differ in their degree of glycosylation and whose relative abundance depended on physiological and pathological conditions (1, 3, 11, 16). In comparison, in the MDCK-UT-A1 cells, the UT-A1 protein was primarily present in the 97-kDa form. However, this did not appear to impair the functional status of the protein, because the MDCK-UT-A1 cells exhibited a robust urea permeability after stimulation with forskolin.

In MDCK-UT-A1 cells, AVP and forskolin increased UT-A1 phosphorylation (Figs. 4 and 6) and both agonists activated [14C]urea fluxes (Figs. 3 and 5). The activated urea fluxes were inhibited by three urea transport inhibitors: thionicotinamide, dimethylurea, and phloretin (Figs. 810). Since the MDCK-UT-A1 cells increased their phosphorylation of UT-A1 and increased their flux in response to low concentrations of AVP or forskolin, which activates protein kinase A, these cells appeared to have functional V2 receptors for AVP. To our knowledge, this is the first epithelial cell model to stably express UT-A1, the urea transporter that is expressed in the IMCD.

The time courses of phosphorylation and flux activation of UT-A1 by forskolin were different. The phosphorylation was rapid (2–5 min; Fig. 6) compared with the flux activation (10–30 min; Fig. 5). There are many possible explanations for a delay in flux activation. This may be due to phosphorylation of UT-A1 at multiple sites, many of which are unrelated to activation or many of which must be phosphorylated before the one critical activating site is phosphorylated. Alternatively, phosphorylation of UT-A1 may be a parallel phenomenon unrelated directly to its activation that may result from the phosphorylation of another protein, or linked chain of proteins, which then activate UT-A1 already in the plasma membrane. The delay in MDCK cells may be due to the slow insertion of UT-A1-containing vesicles into the plasma membrane, although this mechanism has been disproved for AVP activation of urea transport in the renal IMCD of Brattleboro rats (which lack AVP) (10, 22).

AVP increased urea permeability in perfused rat terminal IMCDs (28) and increased UT-A1 phosphorylation in IMCD suspensions (32). Forskolin also increased urea permeability in perfused rat terminal IMCDs (9). Phloretin inhibited AVP-stimulated urea transport in perfused rat terminal IMCDs (4). Although neither dimethylurea nor thionicotinamide has been tested in the perfused IMCD, methylurea and acetamide did inhibit urea transport in the rat terminal IMCD (4). Thus the properties of our stably transfected MDCK-UT-A1 cells reproduced the properties of urea transport in the rat terminal IMCD. This suggests that our transfected cells will be a useful model system for studying the cell biology and signaling pathways that regulate urea transport by each of the urea transporter isoforms.

A potential advantage of the approach that we used to create this cell line is that we initially cloned an FRT site into the genome of the parental MDCK cells and created a stable line of MDCK-FRT cells. By cloning UT-A1 into the MDCK-FRT cells, the UT-A1 cDNA can only incorporate into the FRT site that we introduced into the genome of the parental MDCK cells. In future studies, we will be able to clone other urea transporter isoforms into the MDCK-FRT cells, and these UT-A cDNAs should only be incorporated into the same FRT sites. Thus any difference in function between MDCK cell lines that are stably transfected with different UT-A cDNAs should result from differences in the UT-A protein that is expressed and not from where the transgene is incorporated into the MDCK cell genome.


    GRANTS
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
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This work was supported in part by National Institute of Diabetes and Digestive and Kidney Diseases Grants R01-DK-63657 and R01-DK-41707.


    ACKNOWLEDGMENTS
 
Portions of this work have been published in abstract form (J Gen Physiol 122: 43A, 2003; and J Am Soc Nephrol 14: 74A, 2003) and presented at the Society of General Physiologists, September 4–7, 2003, Woods Hole, MA, and at the Renal Week 2003 meeting, November 14–17, 2003, San Diego, CA.


    FOOTNOTES
 

Address for reprint requests and other correspondence: R. B. Gunn, Emory Univ. School of Medicine, Dept. of Physiology, 601 Whitehead Bldg., 615 Michael St., Atlanta, GA 30322 (E-mail: rbgunn{at}emory.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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