Vitamin D 3 upregulates plasma membrane Ca 2 -ATPase expression and potentiates - 《American Journal of Physiology》美国生理学杂志干细胞之家



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发表于 2009-4-22 08:16 |只看该作者 |倒序浏览 |打印

作者：Sertac N. Kip and Emanuel E. Strehler作者单位：Department of Biochemistry and Molecular Biology, Mayo Clinic, Rochester, Minnesota 55905
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      【摘要】
   Plasma membrane Ca 2  -ATPases (PMCAs) are a ubiquitous system for the expulsion of Ca 2  from eukaryotic cells. In tight monolayers of polarized Madin-Darby canine kidney (MDCK) cells representing a distal kidney tubule model, PMCAs are responsible for about one-third of the vectorial Ca 2  transport under resting conditions, with the remainder being provided by the Na /Ca 2  exchanger. Vitamin D 3 (VitD) is known to increase PMCA expression and activity in Ca 2  -transporting tissues such as the intestine, as well as in osteoblasts and Madin-Darby bovine kidney epithelial cells. We found that VitD upregulated the expression of the PMCAs (mainly PMCA4b) in MDCK cell lysates at the RNA and protein level in a time- and dose-dependent manner. Interestingly, VitD caused a decrease of the PMCAs in the apical plasma membrane fraction and a concomitant increase of the pumps in the basolateral membrane. Functional studies demonstrated that transcellular 45 Ca 2  flux from the apical-to-basolateral compartment was significantly enhanced by VitD. These findings demonstrate that VitD is a positive regulator of the PMCAs in MDCK epithelial cells. The correlation of decreased apical/increased basolateral expression of the PMCAs with an increase in transcellular Ca 2  flux from the apical (urine) toward the basolateral (blood) compartment indicates the physiological relevance of VitD function in kidney tubular Ca 2  reabsorption.
      【关键词】 calcium transport kidney distal tubule MadinDarby canine kidney transcellular ion flux0 `) m% i& k; Y- U3 D" ?
   MAINTENANCE OF overall Ca 2  homeostasis is essential for proper organ function in higher eukaryotes. A delicate balance between Ca 2  loss and absorption exists in vertebrates to provide the necessary amount of Ca 2  for structural (e.g., bone, teeth) and signaling functions (e.g., muscle contraction, neuronal communication). Accordingly, uptake and excretion of Ca 2  are finely tuned to the physiological needs of the body. Ca 2  uptake primarily occurs through the intestine, whereas Ca 2  loss is associated with urinary excretion. As the major organ involved in Ca 2  excretion, the kidney plays a crucial role in the regulation of Ca 2  homeostasis ( 28 ). Although the majority of filtered Ca 2  reabsorption occurs in the proximal tubules via a passive paracellular pathway, the remaining 10-15% are reabsorbed in the distal nephron by an active transcellular and highly regulated pathway ( 16, 28 ). Ca 2  influx into epithelial cells lining the distal tubules occurs through specific channels, such as ECaC1, expressed on the apical membrane ( 14, 15 ). Transcellular Ca 2  transport may be facilitated by Ca 2  -binding proteins such as calbindin-D 28K, and Ca 2  is finally extruded against its electrochemical gradient at the basolateral membrane by plasma membrane Ca 2  pumps and Na /Ca 2  exchangers ( 16 ).
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Because of its importance for overall Ca 2  homeostasis, active transcellular Ca 2  reabsorption in the distal kidney is under tight hormonal control. The "classic" Ca 2  -mobilizing hormones 1,25-(OH) 2 vitamin D 3 (VitD), calcitonin, and parathyroid hormone are all known to act on Ca 2  reabsorption in the kidney ( 7, 28 ), as are several other hormones and ligands such as vasopressin, ATP, and nitric oxide (for a recent succinct review, see Ref. 16 ). Ca 2  influx at the apical membrane is likely the rate-limiting step for transcellular Ca 2  reabsorption, and the ECaC1 channel may thus be a primary target of regulation by the above hormones and receptor agonists ( 14, 16 ). VitD has been shown to upregulate ECaC1 both at the RNA and protein level ( 12, 13 ). For increased transcellular Ca 2  reabsorption, however, Ca 2  buffering/shuttling mechanisms as well as basolateral extrusion must be coordinately upregulated. Calbindin-D 28K, the major Ca 2  -shuttling protein in the kidney, is a well-known target of VitD upregulation, and mRNAs for both the Na /Ca 2  exchanger (NCX1) and the plasma membrane Ca 2  pump (PMCA) isoform 1b have recently been found to be upregulated in 25-hydroxyvitamin D 3 -1 -hydroxylase knockout mice following treatment with VitD ( 12 ). However, no detailed studies have yet been reported on the effects of VitD on the basolateral Ca 2  extrusion mechanisms at the protein and functional levels.; W* r$ @2 I3 L- `2 T" W2 }
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Madin-Darby canine kidney (MDCK) cells are a well-established cell culture model for distal kidney tubules and have been widely used for studies of the expression, regulation, and function of proteins involved in renal ion transport ( 10, 11, 18, 26, 27 ). We recently showed that polarized MDCK cells express primarily PMCA isoforms 1b and 4b and that these pumps account for about one-third of the basolateral Ca 2  efflux under resting conditions, with the remainder being handled by the NCX ( 19 ). Here we show that VitD upregulates the expression of PMCA1b and 4b in MDCK cells in a time- and dose-dependent manner and that this leads to enhanced apico-basal Ca 2  flux via a specific increase of the pumps (mainly PMCA4b) in the basolateral membrane with a concomitant decrease in the apical membrane. These data suggest that VitD is a physiologically relevant regulator of active Ca 2  reuptake in the distal kidney where it functions, at least in part, via increasing the expression and altering the membrane distribution of specific PMCA isoforms.  U! p) n. ^$ T, P) k

MATERIALS AND METHODS, I3 i, X. O) B$ i) B) K( I
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Materials. MDCK type I cells were obtained from the American Type Culture Collection (Manassas, VA). Trypsin-EDTA, DMEM, FBS, L -glutamine, sodium pyruvate, and antibiotics/antimycotics were from Invitrogen (Carlsbad, CA). RT-PCR reagents and enzymes were purchased from Roche-Boehringer Mannheim (Indianapolis, IN). All other chemicals were from Sigma (St. Louis, MO). Monoclonal (5F10, JA9) and polyclonal (NR-1, NR-2) antibodies against PMCAs were provided by Dr. J. T. Penniston and A. G. Filoteo (Mayo Clinic, Rochester, MN). These antibodies have been described previously ( 4, 6 ). Primary antibodies against calbindin-D 28K, occludin, and -actin, and all secondary antibodies were from Sigma, whereas an antibody against Na -K -ATPase was purchased from Affinity Bioreagents (Golden, CO). X-ray films were from Eastman Kodak (Rochester, NY) and 45 Ca 2  was obtained from New England Nuclear Life Science Products (Boston, MA). VitD was a generous gift from Dr. R. Kumar (Mayo Clinic).$ G7 U! _, z  W, [( W' Y2 y  e
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Cell culture. MDCK cells were propagated in DMEM containing 10% (vol/vol) FBS, 2 mM L -glutamine, 1 mM sodium pyruvate, 50 µg/ml gentamicin sulfate, 100 U/ml penicillin, and 100 U/ml streptomycin at 37°C in a humidified atmosphere containing 5% CO 2. MDCKs grown to 70-80% confluency were treated with 10 nM (1 x ), 50 nM (5 x ), or 100 nM (10 x ) doses of VitD in ethanol for 1-4 days. Control MDCKs were subjected to equal amounts of the carrier without VitD.

Semiquantitative RT-PCR. Total RNA was isolated from control and VitD-treated MDCK cells using the TRIzol reagent (Invitrogen), as specified by the manufacturer and as described ( 19 ). Reverse transcription was carried out as described ( 19 ), using PMCA4-specific primers that amplify a 352-nt fragment of PMCA4b. In addition, RT-PCR of a housekeeping transcript (GAPDH) was performed on the same first-strand cDNAs yielding an 220-nt fragment. After RT-PCR, 10% of the amplicons were electrophoresed on a 1.8% agarose gel along with a molecular size standard (100-bp ladder, Bio-Rad, Hercules, CA) and the intensities of the PMCA4-derived fragments were then normalized to those derived from the GAPDH internal standard.

Preparation of total cell extracts and plasma membrane subfractions. Confluent cell cultures were rinsed with PBS, trypsinized, pelleted, and stored at -80°C until further use. Cell extracts and plasma membranes were prepared as previously described ( 19 ). Briefly, total cell lysates were prepared by resuspending the cells in lysis buffer [50 mM HEPES (pH 7.5), 0.1% Nonidet P-40, 0.5% deoxycholate, 1 mM EDTA, 150 mM NaCl, 0.1 mM Na 3 VO 4 ] containing protease inhibitors (aprotinin, leupeptin, pefabloc, and pepstatin), followed by sonication, precipitation with ice-cold 5% TCA, and centrifugation at 4°C for 15 min at 12,000 g. The pellets were then homogenized in Krebs-Ringer-HEPES (KRH) solution containing (in mM) 130 NaCl, 5 KCl, 20 HEPES, 1.2 KH 2 PO 4, 1 CaCl 2, 1 MgSO 4, 10 glucose, as well as 1 ml/l DMEM, pH 7.4.

To generate a purified total plasma membrane fraction, the thawed cell pellets were sonicated (2 bursts, 7 s each) in 10 vol/wt of 0.3 M sucrose containing protease inhibitors; 1.43 vol of 2 M sucrose was added and the mixture was transferred to a TI70 ultracentrifuge tube (Beckman Instruments), overlaid with 0.3 M sucrose and subjected to isopycnic centrifugation for 1 h at 240,000 g. The membrane band was removed, diluted with ice-cold dH 2 O, and spun at 240,000 g for 30 min. The resulting pellet was resuspended in KRH solution, layered onto a 9-60% linear sucrose gradient, and centrifuged at 90,000 g for 3 h in a SW 28 rotor (Beckman Instruments) to obtain the mixed total plasma membranes ( 30 ).
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Apical and basolateral plasma membrane fractions were prepared from the mixed total plasma membranes as described ( 19 ). Briefly, the total plasma membrane band was diluted with 4 vol of 1 mM NaHCO 3, pH 7.5, and sedimented at 7,500 g for 30 min. The resulting pellet was washed with 10 vol of the bicarbonate buffer, centrifuged at 7,500 g for 15 min, resuspended in 0.25 M sucrose, and homogenized with a tight type B glass Dounce homogenizer by 50 up and down strokes. This suspension was layered on top of a three-step sucrose gradient consisting of 38, 34, and 31% sucrose (wt/wt) followed by centrifugation at 20,000 g for 3 h. The bands on top of the 31% sucrose layer and at the interface between the 34 and 38% layer were collected as apical and basolateral plasma membranes, respectively ( 19, 24, 29 ). The apical and basolateral membrane fractions were diluted to 10 ml with 0.125 M sucrose, pelleted at 40,000 rpm for 1 h, and the pellets were resuspended in KRH solution and stored at -80°C until use. To determine the enrichment of the respective membrane fractions, alkaline phosphatase, a common marker for apical plasma membranes ( 25 ), was assayed biochemically using a commercially available kit (Sigma), and immunoblotting for Na -K -ATPase was performed to confirm enrichment of the basolateral plasma membrane domain." |# z; P, ^' M! j

Immunoblotting. Protein concentrations were measured spectrophotometrically using the BCA assay (Pierce, Rockwood, IL). Approximately 30 µg of total cell lysates, 6 µg of total plasma membranes, and 1-2 µg of distinct plasma membrane domains were mixed with Nu-PAGE electrophoresis buffer in the presence of reducing agents and anti-oxidants and heated to 70°C for 15 min before separation in denaturing 4-12% Nu-PAGE gradient gels at 200 V for 50 min. After transfer onto nitrocellulose membranes (1 h, 30 V at room temperature), immunoblotting was performed using standard Western blotting techniques ( 2 ). Briefly, the membranes were blocked for 1 h at room temperature in 50 mM Tris·HCl, pH 7.4, 150 mM NaCl, and 0.05% Tween 20 plus 10% milk before exposure to primary antibodies for 1 h at room temperature. The following primary antibodies were used: 5F10 (1:2,000) and JA9 (1:400) to detect all PMCAs and PMCA4, respectively ( 4 ), NR-1 (1:200) and NR-2 (1:9,000) to detect PMCA1 and PMCA2, respectively ( 6 ), and a commercially available monoclonal antibody to detect calbindin-D 28K (1:3,000). In addition, the blots were reprobed with an anti-Na -K -ATPase -1 antibody (1:500) as a basolateral plasma membrane marker, or with an anti- -actin antibody (1:1,000) as a cytosolic housekeeping protein marker to standardize each lane and ensure equal protein loading. After exposure to primary antibodies, the blots were washed and incubated in peroxidase-conjugated anti-mouse or anti-rabbit IgG (1:5,000) as described ( 19 ). The Renaissance chemiluminescence detection system (PerkinElmer Life Sciences) was used to visualize the immunoreactive bands. Band intensities were determined on a model GS-700 imaging densitometer, and Molecular Analyst software (Bio-Rad) was used to calculate the ratio of PMCA reactivity to that of -actin and Na -K -ATPase. To quantify the relative change in PMCA expression in the different membrane fractions following VitD treatment, the optical density readings of the PMCA-immunoreactive bands were first standardized to the amount of protein loaded per lane and divided by the optical density readings of the respective Na -K -ATPase bands. The value for each control (untreated) sample was set at 1.0, and the fold change following VitD treatment was then determined in the total, apical, and basolateral membrane fraction by comparison to the corresponding control.
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Immunofluorescence confocal microscopy. MDCK cells grown to confluence on glass coverslips were treated with carrier (ethanol, control) or with 10 nM VitD for 3 days, washed with PBS plus Ca 2  and Mg 2  (PBS CM), and then fixed and permeabilized as described ( 19 ). After being blocked for 1 h at room temperature in PBS CM containing 5% normal goat serum and 1% bovine serum albumin, the coverslips were incubated for 1 h with monoclonal anti-PMCA antibody 5F10 (1:800) and polyclonal anti-occludin antibody (1:100). After being washed, the cells were incubated for 1 h at room temperature with secondary antibodies (anti-mouse Alexa 488 and anti-rabbit Alexa 594; Molecular Probes, Eugene, OR) diluted 1:600 in blocking buffer. Coverslips were mounted onto slides with Prolong mounting media (Molecular Probes) and the cells were imaged on a Zeiss LSM 510 laser confocal microscope.- A5 P# K% e9 C. ]& Y9 u: d3 y

Transcellular Ca 2  flux across monolayers of MDCK cells. Functional Ca 2  flux studies were performed on polarized MDCK cells grown on permeable inserts (Costar, Cambridge, MA) as described ( 19 ). The transepithelial transport of 45 Ca 2  was measured on day 15, when the cells were fully differentiated, polarized, and formed tight monolayers. Briefly, the inserts harboring the MDCK cells were rinsed with wash buffer and incubated for 30 min at 37°C in a nonradiolabeled transport medium containing (in mM) 140 NaCl, 5.8 KCl, 0.34 Na 2 HPO 4, 0.44 KH 2 PO 4, 0.8 MgSO 4, 20 HEPES, 4 glutamine, 0.5 CaCl 2, and 25 glucose at pH 7.4. The medium in the top chamber was then replaced with transport medium containing 0.5 mM phenol red and 1 µCi 45 Ca 2 , and the cells were incubated for an additional 30 min at 37°C. Duplicate aliquots were then removed from the bottom compartment and read in the scintillation counter to determine the total transport of 45 Ca 2  from the apical toward the basolateral compartment. To estimate the paracellular 45 Ca 2  flux and the tightness of the monolayers, phenol red transport was measured in the basolateral compartment at the end of the transport study. The percentage of phenol red transport was calculated as described ( 19 ), and an equivalent amount of 45 Ca 2  was subtracted from the total 45 Ca 2  transport to determine the transcellular 45 Ca 2  transport. Transcellular 45 Ca 2  flux was then calculated using the formula J Ca 2  = {(dpm)/( * [Ca 2  ] tA )}10 6 ( 1 ), where J Ca 2  is the unidirectional Ca 2  flux, dpm denotes the total number of disintegrations per minute, * [Ca 2  ] is the specific activity of 45 Ca 2 , t the time in minutes, A the surface area of the transwell filter insert, and 10 6 is a correction factor to convert micromoles to picomoles. Ca 2  flux from the basolateral to the apical compartment was similarly determined by adding the radiolabeled 45 Ca 2  to the bottom (basolateral) compartment and measuring its appearance in the top (apical) compartment.
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Statistical analysis. Each experiment was repeated a minimum of three times and all data were expressed as means ± SE. Statistical differences were analyzed by Student's t -test using StatView, and results were considered to be statistically significant at P & A; l, I2 Q  N; _
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RESULTS, C- B5 F- h+ |& o
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Treatment with VitD results in upregulation of PMCAs in MDCK cells. We previously showed that PMCA1b and PMCA4b are the two main PMCA isoforms expressed in MDCK cells, whereas PMCA2 is present in minor amounts and PMCA3 is virtually undetectable ( 19 ). To determine if VitD treatment leads to a change in the expression of the PMCAs in MDCK cells, we incubated 70-80% confluent cultures with 10 nM VitD for up to 4 days and analyzed the PMCA levels on Western blots of total cell lysates. When probed with an antibody recognizing all PMCAs (5F10 ), a band of the expected size of 140 kDa was readily observed in both control and VitD-treated cells. When standardized to -actin, the amount of total PMCA increased in the VitD-treated cells in a time-dependent manner ( Fig. 1, top ), representing a 1.2-, 1.4-, and 1.7-fold upregulation after 2, 3, and 4 days, respectively. With the use of a specific antibody against PMCA4 (JA9), this upregulation was more readily apparent, reaching 1.2-, 1.6-, and 1.9-fold control levels after 2, 3, and 4 days, respectively ( Fig. 1, middle ). As previously noted, the PMCA1-specific antibody NR-1 detected only a faint band of appropriate size for the full-length pump in MDCK cell lysates and instead reacted with a prominent band at 110 kDa, which presumably reflects a cleavage product of the PMCA1 ( 19 ). Similar to the trend observed for PMCA4, VitD treatment resulted in an enhancement of the PMCA1 signal in a time-dependent manner ( Fig. 1, bottom ). Because our previous study indicated that PMCA4b is the major pump isoform in MDCK cells, we focused our further work on this isoform as well as on the sum of all PMCAs.

Fig. 1. Vitamin D 3 (VitD) upregulates plasma membrane Ca 2  -ATPases (PMCAs) in total lysates of Madin-Darby canine kidney (MDCK) cells. MDCK cells were either kept in normal growth medium [control (C)] or incubated in the presence of 10 nM VitD for up to 4 days. Cells were harvested at different times after addition of VitD as indicated on the top, and total lysates were run on SDS-polyacrylamide gels, blotted onto nitrocellulose membrane, and probed with an antibody recognizing all PMCAs ( top ) or with antibodies specific for PMCA4 ( middle ) or PMCA1 ( bottom ). Right : size of the major PMCA band is indicated in kDa. Note that the band for PMCA1 is smaller than expected for the full-length protein, likely due to proteolytic degradation. The blots were stripped and reprobed for actin ( bottom half of each panel) as a control for equal protein loading. A moderate increase in total PMCA and in PMCA4 and PMCA1 isoforms was seen in the total lysates of MDCK cells treated for several days with VitD.* Z  P6 `( ^  R1 w% g* |

VitD upregulates PMCA4 expression at the mRNA level and in the membrane fraction of MDCK cells. Using semiquantitative RT-PCR with PMCA4-specific primers, we found an increased level of PMCA4b mRNA in MDCK cells treated for 3 days with 10 nM VitD ( Fig. 2 A ). When standardized against a GAPDH control, PMCA4b mRNA increased about twofold, suggesting that long-term treatment with VitD increases transcription and/or stability of the pump mRNA. We next asked if the VitD-dependent upregulation of PMCAs in total MDCK cell lysates was due to increased pump expression in the membrane fraction. As shown in Fig. 2 B, the amount of total PMCA increased significantly in the purified total membrane fraction after treatment with 10 nM VitD for 3 days. A similar enhancement of expression in the membrane fraction was observed when PMCA4 was analyzed separately after standardization against Na -K -ATPase immunostaining ( Fig. 2 B ). By contrast, the increase of PMCA1 in the membrane fraction was less pronounced (data not shown), suggesting that most of the VitD-dependent PMCA upregulation in total membranes was due to PMCA4. Compared with the controls, however, we noted an increase in PMCA1 in the pellets (containing nuclear debris and unlysed cells) of VitD-treated cells, perhaps indicating a redistribution of this pump isoform to a different cellular compartment other than the membranes ( Fig. 2 C ).

Fig. 2. VitD treatment results in increased PMCA expression at the RNA level and in total membranes from MDCK cells. MDCK cells were grown in the absence (C) or presence of 10 nM VitD (VitD3) for 3 days and then harvested for the preparation of RNA for RT-PCR or for total membranes as described in MATERIALS AND METHODS. A : RT-PCR was performed with PMCA4-specific primers and GAPDH-specific primers and the resulting products were separated on agarose gels, yielding fragments of 350 nt for PMCA4b ( top ) and 220 nt for GAPDH ( bottom ). Standardizing against GAPDH revealed a 2-fold increase of PMCA4b after VitD treatment. A 100-bp ladder was run in the marker lane (M); the position of the 500-bp fragment is indicated on the left. B : equal amounts of total membrane protein were separated by SDS-polyacrylamide gel electrophoresis, transferred to nitrocellose, and the membranes were probed with antibodies recognizing all PMCAs ( top ) or PMCA4 ( middle ). The blots were then reprobed with antibodies against the -subunit of Na -K -ATPase ( bottom ) to control for equal protein loading. C : equal amounts of protein in the pellet remaining after membrane isolation were run as in B and probed with an antibody against PMCA1.
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Dose- and time-dependent upregulation of PMCAs in the total plasma membrane fraction of VitD-treated MDCK cells. To determine the dose dependence of the VitD-induced PMCA upregulation, we treated MDCK cells with 10, 50, and 100 nM VitD for 3 days and then probed for PMCA expression in the purified total plasma membrane fractions. As shown above, 10 nM VitD induced a marked 3-fold) upregulation of PMCA expression in the plasma membrane, 10-fold) in the presence of 50 and 100 nM VitD ( Fig. 3 A, top ). As a control, we confirmed that the VitD-dependent intracellular Ca 2  binding protein calbindin-D 28K showed the expected upregulation by VitD in the MDCK cells ( Fig. 3 A, bottom ). Although maximal PMCA upregulation was observed at 50 nM VitD, we used 10 nM VitD in all subsequent studies to remain closer to physiologically relevant conditions. As was observed in total lysates, 10 nM VitD strongly upregulated the PMCAs expressed in the purified plasma membrane fraction in a time-dependent fashion ( Fig. 3 B ).

Fig. 3. Dose- and time-dependent upregulation of PMCAs in purified total plasma membranes from MDCK cells treated with VitD. A : MDCK cells were grown for 3 days in the absence (C) or presence of 10, 50, or 100 nM VitD as indicated on top of each lane. Cells were then harvested and equal amounts of the purified total plasma membrane fractions were separated by SDS-polyacrylamide gel electrophoresis, followed by Western blot analysis using an antibody recognizing all PMCAs ( top ) or calbindin-D 28K (CB D28K; bottom ). B : MDCK cells were grown in the absence (C) or presence of 10 nM VitD for up to 4 days, and aliquots were harvested after 2, 3, or 4 days as indicated on top of each lane. Equal amounts of the purified total plasma membrane fractions were electrophoretically separated and probed for the presence of all PMCAs as described above.

VitD increases PMCA expression in the basolateral plasma membrane. In addition to the time- and dose-dependent upregulation of PMCA expression in cell lysates and purified plasma membranes of MDCK cells, we observed a redistribution of the PMCAs among apical and basolateral membrane domains following VitD treatment. MDCK cells were either untreated (controls) or treated for 3 days with 10 nM VitD. Total membranes and purified plasma membrane subfractions enriched in apical or basolateral domains were then prepared as described in MATERIALS AND METHODS and analyzed for total PMCA content as well as for the expression of PMCA4. As previously reported ( 19 ) and as shown in Fig. 4 A, top, the PMCAs are enriched in the basolateral membrane of resting cells, although a significant amount is also present in the apical domain. Upon treatment with VitD, the amount of total PMCA in the apical membrane domain decreased while that in the basolateral membranes increased about two- to threefold ( Fig. 4 A, top, compare lanes 2 and 5, and lanes 3 and 6; and Fig. 4 B, left ). An essentially identical pattern of distribution was seen when the antibody against PMCA4 was used, i.e., VitD treatment increased PMCA4 expression in the basolateral plasma membrane domain while decreasing its intensity in the apical membrane fraction ( Fig. 4 A, middle, and Fig. 4 B, right ). In evaluating the data in Fig. 4 A, note that only 1 µg of protein was loaded per lane for the basolateral membranes, compared with 2 or 2.5 µg per lane for apical membranes, and 6 µg for total membranes. The relative "purity" of basolateral membrane fractions was determined by the presence of the basolateral marker protein Na -K -ATPase ( Fig. 4 A, bottom ). Similarly, the absence of detectable Na -K -ATPase in the apical membrane fractions indicates the relative purity of these membranes.
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Fig. 4. VitD treatment leads to increased PMCA expression in the basolateral plasma membrane of MDCK cells. A : monolayers of MDCK cells were grown for 3 days in the absence (C) or presence of 10 nM VitD and were then harvested for membrane fractionation as described in MATERIALS AND METHODS. Six micrograms of total membranes (tm), 2-2.5 µg of apical plasma membranes (api), and 1 µg of basolateral plasma membranes (bl) were separated by SDS-polyacrylamide electrophoresis followed by Western blot analysis to detect all PMCAs ( top ) or PMCA4 ( middle ). The blots were then reprobed for the basolateral marker protein Na -K -ATPase ( bottom ). Note that different amounts of total protein were loaded in each lane due to limitations in the amounts of purified membrane fractions obtained. B : quantification of the change in the level of expression of all PMCAs (blue bars) and of PMCA4 (red bars) in total, apical, and basolateral membranes upon treatment with VitD. Data from gels as shown in A were quantified as described in MATERIALS AND METHODS and standardized to the amount of protein loading. The PMCA expression level in membranes from control cells (C) was set to 1.0 (light-colored bars). The expression level in membranes from VitD-treated cells (dark-colored bars) was then compared with the controls and expressed as fold change ± SE in optical density (OD). The number of independent repeats of each experiment is indicated on top of the bars; * P , R6 z, q' H/ L7 ]

Immunofluorescence confocal microscopy was performed on polarized MDCK cells grown in confluent monolayers on glass coverslips in the presence or absence of 10 nM VitD for 3 days. In agreement with the biochemical data showing enrichment of the PMCAs in the basolateral plasma membrane, immunocytochemical localization using the anti-pan-PMCA antibody 5F10 showed the honeycomb pattern typical of lateral membrane staining displayed by the tight junction marker occludin ( Fig. 4 C ). Although not readily quantifiable, using identical settings for fluorescence microscopy this staining pattern appeared to be enhanced in VitD-treated cells ( Fig. 4 C, compare top right to bottom right ).4 O) \- Z* U( f" J

VitD enhances transcellular Ca 2  transport in MDCK cells and shifts net flux toward the basolateral side. The transcellular Ca 2  flux across a tight monolayer of MDCK cells was measured in control cells and in cells treated with 10 nM VitD for 3 days. 45 Ca 2  transport from the apical-to-basolateral chamber, or in the reverse direction from the basolateral to the apical side, was determined after 30 min of incubation as described in MATERIALS AND METHODS. The transcellular 45 Ca 2  flux from the apical to the basolateral direction averaged 7.4 pmol·cm -2 ·min -1 in control cells. This value was significantly increased (1.9-fold) to 14.2 pmol·cm -2 ·min -1 in VitD-treated cells ( n = 20, P
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Fig. 5. VitD treatment increases apical-to-basolateral Ca 2  flux in MDCK cells. Transcellular 45 Ca 2  efflux assays were performed on tight layers of polarized MDCK cells grown for 3 days in the absence (C) or presence of 10 nM VitD. A : 45 Ca 2  flux from the apical-to-basolateral compartment is significantly increased in VitD-treated cells ( P * a, K! Z3 Y6 C9 w3 F

DISCUSSION
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Active Ca 2  reabsorption in the distal kidney occurs via a transcellular pathway and represents a final and tightly controlled mechanism for recovery of filtered Ca 2  before it is excreted in the urine. Accordingly, Ca 2  reabsorption in the distal kidney is under the control of calciotropic hormones, notably parathyroid hormone, calcitonin, and VitD ( 3, 7, 16 ). The importance of VitD for proper Ca 2  metabolism is underlined by a variety of genetic and acquired diseases where VitD deficiency or problems with the VitD receptor and VitD-dependent signaling compromise normal Ca 2  homeostasis. For example, different forms of congenital rickets have been associated with mutations in the genes for the 25-hydroxyvitamin D 3 -1 -hydroxylase (1 -OHase) and the VitD receptor, respectively ( 17, 20 ).: W! y" h5 K; T7 E

In the distal kidney, VitD may target three principal mechanisms: 1 ) Ca 2  inflow at the apical membrane of the tubular epithelial cells, 2 ) intracellular Ca 2  buffering and shuttling between the apical and basolateral membrane, and 3 ) Ca 2  extrusion at the basolateral membrane. Recent evidence showed that the apical Ca 2  channel ECaC1 is regulated by VitD at the genomic and posttranscriptional level. In VitD-depleted rats and homozygous mice deficient in the 1 -OHase, VitD administration increased the mRNA and protein level of ECaC1 and helped restore normal plasma Ca 2  levels ( 12, 13 ). Similarly, the intracellular Ca 2  -buffering protein calbindin-D 28K has long been known to be upregulated by VitD in the kidney ( 5, 31 ). In a recent study on 1 -OHase -/- mice, Hoenderop et al. ( 12 ) found that VitD treatment also resulted in a significant upregulation of the mRNAs for the basolateral Ca 2  efflux proteins NCX1 and PMCA1b. Our results showing a significant increase in total PMCA protein in VitD-treated MDCK cells are in agreement with these animal studies and confirm recent data by Glendenning et al. ( 8 ) who reported VitD-dependent upregulation of PMCA1b mRNA and protein in MDBK cells.$ W, z3 `9 C5 X7 ^6 M+ H0 z. D

VitD complexed to its nuclear receptor is thought to affect the Ca 2  regulatory proteins primarily at the genomic level, i.e., by altering mRNA transcription upon occupancy of specific promoter regulatory site(s) ( 22 ). On the other hand, VitD also affects several of its target proteins by posttranscriptional effects such as mRNA stabilization or increased translation and/or protein trafficking to the membrane. Glendenning et al. ( 8 ) noted that the increase in PMCA1b mRNA in VitD-stimulated Madin-Darby bovine kidney cells was due, at least in part, to preferential mRNA stabilization. VitD treatment of VitD-depleted rats resulted in a more modest increase in ECaC1 mRNA than ECaC1 protein, indicating a posttranscriptional mechanism to increase protein expression at the membrane ( 13 ). Using semiquantitative RT-PCR, we found that VitD treatment increased the level of PMCA4b mRNA in MDCK cells, suggesting that the observed upregulation of the PMCAs in these cells is at least partially due to increased transcription and/or increased mRNA stability. Our data also indicate that VitD may facilitate the concentration of PMCAs (mainly PMCA4b) in the basolateral membrane domain. This redistribution of the PMCA favors apical-to-basolateral Ca 2  flux, as would be expected for a VitD-stimulated increase in Ca 2  reabsorption in the distal kidney (see Fig. 5 C ). Finally, it should be noted that VitD may also have indirect effects on PMCA expression, e.g., via a sustained increase in intracellular [Ca 2  ]. The VitD-dependent increase in Ca 2  influx via ECaC1 might lead to an elevation of resting [Ca 2  ], and a rise in [Ca 2  ] has been shown to stimulate PMCA transcription in different cell types ( 9, 21 ).

What is the relative role of the different PMCA isoforms in VitD-stimulated Ca 2  reabsorption? Distal convoluted tubules and MDCK cells express PMCA1b and -4b, as well as smaller amounts of PMCA2b ( 19, 23 ). Under resting conditions, some Ca 2  extrusion must occur at the apical plasma membrane to counteract Ca 2  leakage. Accordingly, small amounts of PMCA are detected in the apical membrane. It is conceivable that VitD selectively decreases the amount of PMCA in the apical membrane via nongenomic effects that disrupt the membrane-stabilizing interactions of a specific PMCA isoform. Interestingly, we noted an increased amount of PMCA1b in the insoluble pellet after VitD treatment in MDCK cells, concomitant with a decrease in PMCA in the apical membrane. In addition to upregulating the total amount of PMCA, VitD may thus induce a redistribution of the pumps by promoting the selective removal of apical PMCA isoforms and increasing the trafficking or stabilization of basolateral PMCAs. These effects could conceivably be transmitted via altered phosphorylation of specific PMCA isoforms or through changes in the lipid environment of the pumps or alternatively through effects on the synthesis and distribution of PMCA-interacting proteins.: z8 W: U: I% b

In conclusion, we showed that VitD upregulates both PMCA1b and -4b in polarized MDCK cells and induces the preferential expression of the pump in the basolateral membrane, thereby increasing apical-to-basolateral Ca 2  efflux. VitD therefore acts at multiple levels to enhance transcellular Ca 2  reabsorption in the distal kidney: at the apical membrane by increasing the expression of the Ca 2  influx channel ECaC1 ( 12, 13 ), intracellularly by increasing the expression of the Ca 2  -buffering protein calbindin-D 28K ( 31 ), and at the basolateral membrane by upregulating the expression of PMCAs. The coordinate regulation of the complex machinery for transcellular Ca 2  reabsorption is a prerequisite for the exquisite control of net Ca 2  flux to maintain overall Ca 2  homeostasis and react to the body's changing demands for Ca 2 . Further studies will be required to investigate if PMCA1b and PMCA4b play unique roles in Ca 2  reabsorption and how they are individually regulated by other calciotropic hormones or diverse pharmacological and pathological stimuli.

GRANTS

This work was supported by National Institute of General Medical Sciences Grant GM-58710 and the Mayo Foundation for Medical Education and Research.! `: ?8 f; ]8 V1 m/ G
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ACKNOWLEDGMENTS
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We thank A. Filoteo and J. T. Penniston for the PMCA-specific antibodies, R. Kumar for the gift of VitD, A. S. Lienhard for technical assistance, and all members of the Strehler lab for helpful discussions.
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