Department of Biochemistry and Molecular Biology, Mayo Clinic, Rochester, Minnesota 55905
Submitted 20 February 2003 ; accepted in final form 15 October 2003
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
calcium transport; kidney distal tubule; Madin-Darby canine kidney; transcellular ion flux
Because of its importance for overall Ca2+ homeostasis, active transcellular Ca2+ reabsorption in the distal kidney is under tight hormonal control. The "classic" Ca2+-mobilizing hormones 1,25-(OH)2 vitamin D3 (VitD), calcitonin, and parathyroid hormone are all known to act on Ca2+ 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). Ca2+ influx at the apical membrane is likely the rate-limiting step for transcellular Ca2+ 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 Ca2+ reabsorption, however, Ca2+ buffering/shuttling mechanisms as well as basolateral extrusion must be coordinately upregulated. Calbindin-D28K, the major Ca2+-shuttling protein in the kidney, is a well-known target of VitD upregulation, and mRNAs for both the Na+/Ca2+ exchanger (NCX1) and the plasma membrane Ca2+ pump (PMCA) isoform 1b have recently been found to be upregulated in 25-hydroxyvitamin D3-1-hydroxylase knockout mice following treatment with VitD (12). However, no detailed studies have yet been reported on the effects of VitD on the basolateral Ca2+ extrusion mechanisms at the protein and functional levels.
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 Ca2+ 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 Ca2+ 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 Ca2+ 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.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
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% CO2. MDCKs grown to 70-80% confluency were treated with 10 nM (1x), 50 nM (5x), or 100 nM (10x) 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 Na3VO4] 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 KH2PO4, 1 CaCl2, 1 MgSO4, 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 dH2O, 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).
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 NaHCO3, 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.
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-D28K (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.
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 Ca2+ and Mg2+ (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.
Transcellular Ca2+ flux across monolayers of MDCK cells. Functional Ca2+ flux studies were performed on polarized MDCK cells grown on permeable inserts (Costar, Cambridge, MA) as described (19). The transepithelial transport of 45Ca2+ 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 Na2HPO4, 0.44 KH2PO4, 0.8 MgSO4, 20 HEPES, 4 glutamine, 0.5 CaCl2, 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 45Ca2+, 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 45Ca2+ from the apical toward the basolateral compartment. To estimate the paracellular 45Ca2+ 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 45Ca2+ was subtracted from the total 45Ca2+ transport to determine the transcellular 45Ca2+ transport. Transcellular 45Ca2+ flux was then calculated using the formula JCa2+ = {(dpm)/(*[Ca2+]tA)}106 (1), where JCa2+ is the unidirectional Ca2+ flux, dpm denotes the total number of disintegrations per minute, *[Ca2+] is the specific activity of 45Ca2+, t the time in minutes, A the surface area of the transwell filter insert, and 106 is a correction factor to convert micromoles to picomoles. Ca2+ flux from the basolateral to the apical compartment was similarly determined by adding the radiolabeled 45Ca2+ to the bottom (basolateral) compartment and measuring its appearance in the top (apical) compartment.
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 0.05.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
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. 2A). 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. 2B, 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. 2B). 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. 2C).
|
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, and this upregulation reached even higher levels (>10-fold) in the presence of 50 and 100 nM VitD (Fig. 3A, top). As a control, we confirmed that the VitD-dependent intracellular Ca2+ binding protein calbindin-D28K showed the expected upregulation by VitD in the MDCK cells (Fig. 3A, 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. 3B).
|
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. 4A, 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. 4A, top, compare lanes 2 and 5, and lanes 3 and 6; and Fig. 4B, 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. 4A, middle, and Fig. 4B, right). In evaluating the data in Fig. 4A, 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. 4A, bottom). Similarly, the absence of detectable Na+-K+-ATPase in the apical membrane fractions indicates the relative purity of these membranes.
|
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. 4C). Although not readily quantifiable, using identical settings for fluorescence microscopy this staining pattern appeared to be enhanced in VitD-treated cells (Fig. 4C, compare top right to bottom right).
VitD enhances transcellular Ca2+ transport in MDCK cells and shifts net flux toward the basolateral side. The transcellular Ca2+ flux across a tight monolayer of MDCK cells was measured in control cells and in cells treated with 10 nM VitD for 3 days. 45Ca2+ 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 45Ca2+ 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 < 0.0001; Fig. 5A). Transcellular 45Ca2+ flux in the reverse direction (from the basolateral to the apical side) was determined at 8.2 pmol·cm-2·min-1 and did not change significantly in VitD-treated MDCK cells (10.5 pmol·cm-2· min-1, P = 0.07, n = 17; Fig. 5B). Overall, transcellular Ca2+ flux across monolayers of resting MDCK cells at equilibrium was similar in both directions. In marked contrast, the net direction of Ca2+ efflux was from the apical-to-basolateral compartment in the VitD-treated cells, indicating a shift in the directionality of the overall Ca2+ flux under these conditions (Fig. 5C). Importantly, in the distal kidney, this would correspond to calcium reabsorption from the urine into the blood.
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
In the distal kidney, VitD may target three principal mechanisms: 1) Ca2+ inflow at the apical membrane of the tubular epithelial cells, 2) intracellular Ca2+ buffering and shuttling between the apical and basolateral membrane, and 3) Ca2+ extrusion at the basolateral membrane. Recent evidence showed that the apical Ca2+ 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 Ca2+ levels (12, 13). Similarly, the intracellular Ca2+-buffering protein calbindin-D28K 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 Ca2+ 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.
VitD complexed to its nuclear receptor is thought to affect the Ca2+ 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 Ca2+ flux, as would be expected for a VitD-stimulated increase in Ca2+ reabsorption in the distal kidney (see Fig. 5C). Finally, it should be noted that VitD may also have indirect effects on PMCA expression, e.g., via a sustained increase in intracellular [Ca2+]. The VitD-dependent increase in Ca2+ influx via ECaC1 might lead to an elevation of resting [Ca2+], and a rise in [Ca2+] 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 Ca2+ reabsorption? Distal convoluted tubules and MDCK cells express PMCA1b and -4b, as well as smaller amounts of PMCA2b (19, 23). Under resting conditions, some Ca2+ extrusion must occur at the apical plasma membrane to counteract Ca2+ 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.
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 Ca2+ efflux. VitD therefore acts at multiple levels to enhance transcellular Ca2+ reabsorption in the distal kidney: at the apical membrane by increasing the expression of the Ca2+ influx channel ECaC1 (12, 13), intracellularly by increasing the expression of the Ca2+-buffering protein calbindin-D28K (31), and at the basolateral membrane by upregulating the expression of PMCAs. The coordinate regulation of the complex machinery for transcellular Ca2+ reabsorption is a prerequisite for the exquisite control of net Ca2+ flux to maintain overall Ca2+ homeostasis and react to the body's changing demands for Ca2+. Further studies will be required to investigate if PMCA1b and PMCA4b play unique roles in Ca2+ reabsorption and how they are individually regulated by other calciotropic hormones or diverse pharmacological and pathological stimuli.
![]() |
GRANTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
![]() |
ACKNOWLEDGMENTS |
---|
![]() |
FOOTNOTES |
---|
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.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|