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. Author manuscript; available in PMC: 2025 May 19.
Published in final edited form as: Xenobiotica. 2021 Apr 18;51(6):657–667. doi: 10.1080/00498254.2021.1909783

Influence of vitamin D treatment on functional expression of drug disposition pathways in human kidney proximal tubule cells during simulated uremia

Stacey M Tuey a, Amandla Atilano-Roque a, Georgia Charkoftaki a,b, Joshua M Thurman c, Thomas D Nolin d, Melanie S Joy a,c
PMCID: PMC12088830  NIHMSID: NIHMS2081573  PMID: 33870862

Abstract

1. Effects of cholecalciferol (VitD3) and calcitriol (1,25-VitD3), on the expression and function of major vitamin D metabolizing enzymes (cytochrome P450 [CYP]2R1, CYP24A1) and select drug transport pathways (ABCB1/P-gp, SLCO4C1/OATP4C1) were evaluated in human kidney proximal tubule epithelial cells (hPTECs) under normal and uraemic serum conditions.

2. hPTECs were incubated with 10% normal or uraemic serum for 24 h followed by treatment with 2% ethanol vehicle, or 100 and 240 nM doses of VitD3, or 1,25-VitD3 for 6 days. The effects of treatment on mRNA and protein expression and functional activity of select CYP enzymes and transporters were assessed

3. Under uraemic serum, treatment with 1,25-VitD3 resulted in increased mRNA but decreased protein expression of CYP2R1. Activity of CYP2R1 was not influenced by serum or VitD analogues. CYP24A1 expression was increased with 1,25-VitD3 under normal as well as uraemic serum, although to a lesser extent. ABCB1/P-gp mRNA expression increased under normal and uraemic serum, with exposure to 1,25-VitD3. SLCO4C1/OATP4C1 exhibited increased mRNA but decreased protein expression, under uraemic serum + 1,25-VitD3. Functional assessments of transport showed no changes regardless of exposure to serum or 1,25-VitD3.

4. Key findings indicate that uraemic serum and VitD treatment led to differential effects on the functional expression of CYPs and transporters in hPTECs.

Keywords: Renal metabolism, renal transport, cell culture, CYP enzymes, P-glycoprotein, organic anion transporter, vitamin D, uraemia, proximal tubule cells

Introduction

Vitamin D (VitD) deficiency in the United States is increasingly recognized as an important public health problem, with estimated prevalence rates of 40–100% in older patients.(Holick 2007, Holick et al 2011) Epidemiological evidence supports the association of low serum calcidiol (25-VitD) with not only bone disorders, but cancer, cardio-metabolic disorders, and other chronic medical conditions (Cheng and Coyne 2007, Holick 2007, Jones 2007, Holick et al 2011). VitD insufficiency, defined as serum 25-VitD ranging from 21–29 ng/ml, and deficiency, defined as 25-VitD ≤20 ng/ml, (Holick 2007) are highly prevalent in the chronic kidney disease (CKD) population, with estimates ranging from 57 to 97%. (Thomas et al 1998, Massry et al 2003, González et al 2004, Coresh et al 2007; Mehrotra et al 2008, 2009, Bhan et al 2010, Kandula et al 2011, Mehrotra et al 2013) Studies in CKD patients have shown that low serum 25-VitD concentrations are associated with fatal and non-fatal cardiovascular events. (Mehrotra et al 2013) Because CKD is a strong predictor of premature cardiovascular disease, a positive effect of VitD replacement on cardiovascular health would be beneficial. VitD deficiency has also been associated with progression of CKD, and replacement has been purported to have reno-protective effects. (Schwarz et al 1998, Hirata et al 2002, Jones 2007, Heaney 2008, Tan et al 2008, Zhang et al 2008, Li 2010) Current clinical guidelines recommend treatment with cholecalciferol (VitD3) or ergocalciferol (VitD2) in CKD patients who have 25-VitD concentrations <30 ng/ml. (Holick et al. 2011, Pramyothin and Holick 2012).

Despite the high prevalence of VitD deficiency and subsequent replacement in CKD, limited information exists to inform whether VitD status or treatment has a potential for interactions with co-prescribed medications. This is relevant as CKD patients are prescribed an average of 10–12 different medications (Manley et al 2003) and drug-drug interactions were not previously evaluated with cholecalciferol and ergocalciferol. Additionally, animal studies have suggested changes in the expression and function of drug metabolism and transport pathways after exposure to VitD and its analogs. (Fan et al 2009a, Chow et al 2010, 2011) As VitD and its analogues also undergo extensive cytochrome P450 (CYP) metabolism, effects of these treatments on metabolism of concomitant medications needs to be informed. The majority of VitD3 in humans is obtained through the skin from 7-dehydrocholesterol after ultraviolet light radiation or from the diet as VitD3 and VitD2. VitD is subsequently converted to 25-VitD by the enzyme 25-hydroxylase encoded by the CYP2R1 gene, which is expressed predominately in the liver but has also been shown to be expressed in the kidney and intestine (Tucker et al 1973). The 25-VitD undergoes a second hydroxylation step, primarily in the kidney, resulting in activation to 1,25-dihydroxyvitamin D (1,25-VitD) or catabolism to 24,25-dihydroxyvitamin D (24,25-VitD) (Figure 1).

Figure 1.

Figure 1.

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Scheme of VitD3 metabolism. The VitD3 parent drug is metabolized to 25-VitD3 by CYP2R1 located primarily in liver and kidney. The 25-VitD3 metabolite can be activated through CYP27B1 (primarily in kidney) to 1,25-VitD3. Alternatively, the 25-VitD3 metabolite can be catabolized by CYP24A1 to 24,25-VitD3. While the figure represents VitD3, the scheme is also relevant for VitD2. Abbreviations: VitD3, cholecalciferol; 25-VitD3, calcidiol; 1,25-VitD3, calcitriol; 24,25-VitD3, 24,25-dihydroxycholecalciferol.

As the kidney plays a major role in VitD metabolism, the goal of the current study was to investigate the influence of VitD3 and 1,25-VitD3 on the expression and functional activity of the renal CYP metabolizing enzymes CYP2R1, CYP27B1, and CYP24A1 using human kidney proximal tubule cells (hPTECs). In order to evaluate potential for VitD interactions at the level of renal drug transport pathways, select drug transporters (ABCB1/P-gp, SLCO4C1/OATP4C1) were also evaluated. In order to also inform the concomitant influence of advanced kidney diseases on metabolism and transport pathways, hPTECs were studied during simulated uraemia.

Materials and methods

Materials

VitD3 and 1,25-VitD3 were purchased from Sigma Aldrich (Saint Louis, MO). Epithelial Cell Medium (EpiCM), foetal bovine serum (FBS), epithelial cell growth supplement and penicillin-streptomycin were purchased from ScienCell Research Laboratories (Carlsbad, CA), and 0.25% trypsin-EDTA, 1X (0.25%) and Dulbecco’s Modified Eagle Medium (DMEM) were purchased from Life Technologies (Carlsbad, CA). [3H]Digoxin (specific activity, 29.8 Ci/mmol) was purchased from PerkinElmer (Boston, MA). Six-well Transwell plates (pore diameter 0.4 μm) were purchased from Corning Life Sciences (Lowell, MA). Human normal serum was purchased from Biological Specialty Corporation®. Normal serum was obtained from a 49-year-old, male, with no documented renal impairment. Human uraemic serum was obtained from pooled unidentified end stage renal disease patients (just prior to their scheduled dialysis treatments, e.g. pre-dialysis) at the University of Colorado Hospital under an Institutional Review Board approved protocol.

Cell culture

hPTECs were obtained from ScienCell Research Laboratories (Lot #5127). hPTECs were analysed for cytokeratin-18, –19, and vimentin by immunofluorescence. Cells were passaged twice before use. Cells were cultured in EpiCM supplemented with 2% FBS, 1% epithelial cell growth supplement, 1% penicillin and streptomycin in a humidified incubator (atmosphere of 5% CO2 and 95% relative humidity at 37 °C). Monolayer confluence was verified by visualization using an inverted microscope (10x, Nikon Eclipse, TE 2000-S).

Viability assay

hPTECs were seeded onto 96-well plates at a density of 20 000 cells/well. Cells were incubated with 10% normal or uraemic serum for 24 h. Following 24-hour incubation, serum was removed and cells were then treated with 2% ethanol (VEH), VitD3 (100 or 240 nM) or 1,25-VitD3 (100 or 240 nM) for six consecutive days with treatments changed daily. Six days were justified based on a previous published study that investigated the effects of 1,25-VitD3 on gene expression and function in Caco-2 cells (Fan et al 2009a) and preliminary studies in the laboratory which demonstrated adequate cell viability using hPTECs. Due to solubility issues with VitD analogues, a 2% EtOH solution was used as the VEH. Preliminary experiments in the laboratory compared mRNA expression of target genes in hPTECs that were incubated in media only versus those treated with 2% EtOH for 6 days. These initial experiments confirmed that the 2% EtOH VEH did not affect mRNA expression. Uraemic serum was pooled from haemodialysis patients (n = 10) to reduce individual biases. Following treatment, medium was removed, and cells were washed with phenol-free DMEM. 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium (MTS) solution in DMEM at a 1:5 ratio was added to each well and incubated for 45minutes. Cell viability was assessed and quantified using a BioTek Synergy™ 4 Hybrid Microplate Reader (Winooski, VT) and Gen5 1.10 software (BioTek) at a wavelength of 490 nm.

Gene expression

hPTECs were seeded onto 100 × 20 mm/well plates at a density of 5 000 cells/cm2. Cells were incubated with 10% normal or uraemic serum for 24 h. Serum was subsequently removed and cells were treated with VEH, VitD3 (100 or 240 nM) or 1,25-VitD3 (100 or 240 nM) for six consecutive days. Treatments were changed daily. Following treatment, cells were harvested and lysed, and total mRNA was prepared from lysates using an Ambion RNA Extraction Kit (Life Technologies, Grand Island, NY). cDNA was generated using Applied BioSciences Taqman Reverse transcription reagents and a 2720 thermal cycler (Applied Biosystems, Foster City, CA). Materials and reagents for real-time quantitative polymerase chain reaction (RT-qPCR) were purchased from Applied Biosystems® (Foster City, CA). Commercial gene expression assays of CYP2R1 (Hs01379776), CYP24A1 (Hs01096154), CYP27B1 (Hs01096154), ABCB1 (Hs00184500), SLCO4C1 (Hs00698884), and the housekeeping gene GAPDH (Hs02758991) were purchased from Applied Biosystems. The transporter genes ABCB1 and SLCO4C1 were representative drug transporters for the digoxin probe. Real-time PCR was performed using the 7500 Real Time PCR system (Applied Biosystems). Generated data were analysed using the 7500 Software v2.0.6 (Applied Biosystems) using the comparative Ct method (2−ΔΔCt) (Livak and Schmittgen 2001).

Protein expression

hPTECs were seeded onto 100 × 20 mm/well plates at a density of 5 000 cells/cm2. Cells were incubated with 10% normal or uraemic serum for 24 h. Serum was subsequently removed and cells were treated with VEH, VitD3 (240 nM) or 1,25-VitD3 (100 or 240 nM) for six consecutive days. Treatments were changed daily. Following treatment, protein was extracted using a cell RIPA lysis buffer (Santa Cruz Biotechnology, sc24948). Protein concentration was measured using the Pierce™ BCA Protein Assay Kit. Protein (23 mg) was loaded onto a 4–20% Mini-PROTEAN® TGX™ precast gel (Bio-Rad Laboratories, Hercules, CA) and transferred to 0.2 μm nitrocellulose membranes (Bio-Rad Laboratories). Non-specific antibody binding was blocked by incubation of the membranes with 10% w/v non-fat dry milk in 1x Tris-buffered saline (pH 7.4) for 1 h at room temperature. Proteins to probe were selected based on gene expression studies. Membranes were incubated overnight at 4 °C with rabbit anti-P-gp monoclonal antibody (Cell Signalling, Danvers, MA) at 1:1000 dilution, rabbit anti-OATP4C1 polyclonal antibody (Sigma Aldrich, St. Louis, MO) at 1:50 dilution, mouse anti-CYP24A1 monoclonal antibody (Santa Cruz Biotechnology, Dallas, TX) at 1:500 dilution, or rabbit anti-CYP2R1 polyclonal antibody (Abcam, Cambridge United Kingdom) at 1:500 dilution with 1% w/v non-fat dry milk in 1x Tris buffered saline containing 0.1% Tween 20 (TBS-T). CYP27B1 protein was not probed secondary to lack of gene expression. After washing with TBS-T (pH 7.4), membranes were incubated with the secondary antibody, goat anti-rabbit horseradish peroxidase (Cell Signalling, Danvers, MA) or horse anti-mouse horseradish peroxidase (Cell Signalling, Danvers, MA) at 1:2000 for 1 h at room temperature. Bands were visualized with chemiluminescence solution (1:1 ratio, SuperSignal® West Pico Luminol enhancer solution, SuperSignal® West Pico Stable peroxide solution, Thermo Scientific, Rockford, IL) per the manufacturer’s instructions. β-actin (Cell Signalling, Danvers, MA) was used as a loading control. Protein expression was semi-quantified using Image Lab v.5.2.1 build 11, Bio-Rad Laboratories and normalized to β-actin. Quantification (RT) values were compared to control (normal serum + VEH).

CYP2R1 activity assessments

hPTECs were plated into T-25 flasks at a density of 2.5 × 105 cells/flask. Cells were incubated with 10% normal or uraemic serum for 24 h. Subsequently, serum was removed and cells were treated with VEH, VitD3 (240 nM) or 1,25-VitD3 (240 nM) for six consecutive days for potential induction, with treatments changed daily. At Day 6, treatments were removed, cells were washed with PBS and then incubated with 100 nM (38.5 ng/mL) VitD3 (CYP2R1 probe) for 1, 2, or 4 h. The 100 nM dose of VitD3 (CYP2R1 probe) was selected based on the linear range of the ELISA assay. At indicated times, cell media was collected, centrifuged at 1000 xG for 15 minutes at 4 °C and immediately stored at −80 °C. Collected cells were washed three times with PBS to remove residual media, detached with 0.25% trypsin and centrifuged at 1000 xG at room temperature for 5 minutes. Cells were washed an additional 3 times to remove residual trypsin and then resuspended in PBS. Cells were then subjected to 3 freeze/thaw cycles for cell lysis and then centrifuged at 1000 xG for 15 minutes at 4 °C to remove debris. The supernatant was collected and cell lysates were stored at −80 °C. VitD3 and 25-VitD3 concentrations were quantified in cell media and lysates using ELISA kits (MyBioSource, San Diego CA) according to manufacturer’s instructions. Based on the Certificate of Analysis from the manufacturer, the linear range and sensitivity were 0 to 50 ng/ml and 0.1 ng/ml, respectively, for the VitD3 assay, and 0 to 2500 pg/ml and 1.0 pg/ml, respectively, for the 25-VitD3 assay. The manufacturer also reported no significant cross-reactivity with other VitD analogues for either ELISA kit. Area under the concentration vs. time curves from 0 to 4 h (AUC0–4h) for VitD3 and 25-VitD3 in cell median and lysates were calculated using the trapezoidal method. Metabolic ratios were determined by dividing the AUC0–4h of 25-VitD3 by the AUC0–4h of VitD3 in lysates and media, respectively.

Transport assessments

hPTECs were seeded onto 6-well Transwell plates at a density of 4 × 105 cells/cm2. Cells were incubated with 10% normal or uraemic serum for 24 h. Serum was removed and cells were subsequently treated with VEH or 1,25-VitD3 (240 nM) for six consecutive days. Transport experiments were assessed at 1, 2, 4, and 6 h after digoxin treatment to evaluate (i) apical-to-basolateral transport (apically treated and basolaterally measured, A→B) and (ii) basolateral-to-apical transport (basolaterally treated and apically measured, B→A). For A→B transport, the apical compartment contained 33.5 μM (1 μCi) of [3H]digoxin while the basolateral compartment contained 2.6 mL of incubation medium. For B→A transport, the basolateral compartment contained 33.5 μM (1 μCi) of [3H]digoxin while the apical compartment contained 1.5 mL of incubation medium. For all experiments, a 25-μL aliquot was collected from each compartment at the designated time points (1, 2, 4 and 6 h). Liquid scintillation fluid (10 mL) (Bio-safe-II, RPI, Chicago, IL) was added to each 25-μL aliquot (from above), and radioactivity was counted (model LS6000, Beckman Coulter Canada, Inc., Mississauga, ON, Canada). For Transwell studies, % transported in the A→B and B→A direction were calculated. The apparent permeability coefficient (Papp) of [3H]digoxin across hPTEC monolayers was calculated as shown in Equation 1:

Papp=dQdt1A×C0 (1)

Where dQ/dt is the amount accumulated in the receiver compartment over time, A is the surface area of the membrane where transport occurred, and C0 was the initial concentration in the donor compartment. The efflux ratio is the ratio of Papp in the B→A:A→B direction as shown in Equation 2:

EffluxRatio=Papp(B→A)Papp(A→B) (2)

Integrity of cellular barrier and tight junction dynamics in our transwell culture was measured using Transepithelial Electrical Resistance (TEER). Values in the range of 70–75 Ωcm2 were deemed sufficient based on previous data from the laboratory. TEER values < 70 Ω*cm2 were rejected and not used for transport assay assessments.

Statistics

Data are presented as means ± standard error of the mean (SEM) of at least 2 replicates. Differences between VitD treatments and between normal and uraemic serum incubations were evaluated by two-way analysis of variance with Tukey’s post-hoc testing using GraphPad Prism (v.4, San Diego, CA).

Results

Viability assessments

Cell viability was significantly increased in cells treated with normal serum followed by 100 nM and 240 nM 1,25-VitD3 (44% and 26% increase, respectively, p < 0.01) relative to normal serum + VEH (Figure 2). On the contrary, 24 h pre-treatment with uraemic serum followed by 100 nM and 240 nM 1,25-VitD3 had no significant effects on increasing cell viability relative to VEH. VitD3 treatment had no significant effects on cell viability under normal or uraemic serum.

Figure 2.

Figure 2.

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hPTEC Survival. Survival of human proximal tubule epithelial cells (hPTECs) was assessed after 24-hour exposure to normal or uraemic serum followed by treatment for six days with (i) vehicle control (VEH), (ii) 100 nM VitD3, (iii) 240 nM VitD3, (iv) 100 nM 1,25-VitD3, or (v) 240 nM 1,25-VitD3. Cell viability was normalized to 2% v/v ethanol vehicle control + normal serum. Data are means ± SEM of 9 replicates. Data were analysed by analysis of variance (two-way ANOVA) followed by Tukey’s post-hoc test. **p < 0.01 compared to normal serum + VEH, #p < 0.05 compared to normal serum + treatment with the same VitD analogue.

Gene expression

mRNA expression of metabolism and transport genes in hPTECs under VEH and treatment conditions for six days were evaluated (Figure 3). All data were normalized to the normal serum with VEH. Data was not available for CYP27B1 due to low expression (cycle threshold (CT) value > 35). CYP2R1 expression was increased with normal serum plus treatment with either 240 nM VitD3 (3.6-fold; p < 0.01) or 240 nM 1,25-VitD3 (4.7-fold; p < 0.01) compared to normal serum + VEH (Figure 3(A)). Expression of CYP2R1 was significantly increased under uraemic serum followed by treatment with 100 nM VitD3 (3.1-fold; p < 0.01), or 100 nM or 240 nM 1,25-VitD3 (4.8- and 4.0-fold, respectively; p < 0.01) relative to VEH + normal serum. Under uraemic conditions, treatment with 100 nM VitD3 resulted in 45% increase in CYP2R1 expression compared to normal serum (p < 0.01). Uraemic serum followed by treatment with VEH resulted in a 4.5-fold increase in CYP2R1 vs. normal serum + VEH. When compared to normal serum + VEH, CYP24A1 expression increased following exposure to normal serum plus treatment with either 100 nM or 240 nM 1,25-VitD3 (8706-fold, p < 0.01 and 9483fold, p < 0.01, respectively) (Figure 3(B)). CYP24A1 expression also increased following exposure to uraemic serum plus treatment with either 100 nM 1,25-VitD3 (4967-fold, p < 0.01) or 240 nM 1,25-VitD3 (5343-fold, p < 0.01) when compared to normal serum + VEH but to a lesser extent than under normal conditions. ABCB1 expression increased following 100 nM 1,25-VitD3 treatment with normal serum (8.8-fold, p < 0.01) and uraemic serum (10-fold, p < 0.01), as well as by 240 nM 1,25-VitD3 with normal serum (7.6-fold, p < 0.01) (Figure 3(C)). VitD3 treatment had no effect on ABCB1 expression. SLCO4C1 expression was increased under uraemic serum plus treatment with 240 nM 1,25-VitD3 (3.3-fold, p < 0.01) compared to normal serum + VEH (Figure 3(D)). No other significant differences were noted with SLCO4C1.

Figure 3.

Figure 3.

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mRNA expression of metabolism and transport genes. (A) CYP2R1, (B) CYP24A1 (C) ABCB1, and (D) SLCO4C1 gene expression in human proximal tubule epithelial cells (hPTECs). Gene expression was assessed after 24-hour exposure to normal or uraemic serum followed by treatment for six days with (i) vehicle control (VEH), (ii) 100 nM VitD3, (iii) 240 nM VitD3, (iv) 100 nM 1,25-VitD3, or (v) 240 nM 1,25-VitD3. The relative gene expression values were compared to normal serum + VEH treated cells. GAPDH was used as the housekeeping gene. Data are expressed as means ± SEM of 3 replicates and analysed by analysis of variance (two-way ANOVA) followed by Tukey’s post-hoc test. p < 0.01, p < 0.05 compared to normal serum + VEH, ##p < 0.01 compared to normal serum + treatment with the same VitD analogue.

Protein expression

Expression of metabolism and transport proteins in hPTECs under VEH and treatment conditions for six days were evaluated (Figure 4). Given that the high (240 nM) dose of VitD3 appeared to have a greater influence on expression of target genes, this dose of VitD3 was selected for assessment of protein expression. Under uraemic conditions, treatment with 100 nM 1,25-VitD3 and 240 nM 1,25-VitD3 resulted in a 38% and 64% decrease, respectively in CYP2R1 expression compared to the same VitD treatments with normal serum (p < 0.05) (Figure 4(A)). There were no effects from serum or VitD treatment on CYP24A1 protein expression (Figure 4(B)). P-gp expression was not different within and between VitD treatment groups relative to VEH in normal serum (Figure 4(C)). However, treatment with 240 nM 1,25-VitD3 + uraemic serum resulted in a 1.44-fold increase in P-gp expression compared to the same VitD treatment with normal serum (p < 0.05). Under normal serum conditions, treatment with 240 nM VitD3 and 100 nM 1,25-VitD3 had no effect on OATP4C1 expression but 1,25-VitD3 at 240 nM resulted in a 54% decrease in OATP4C1 expression relative to normal serum + VEH (Figure 4(D)). OATP4C1 expression was significantly decreased across all VitD treatments under uraemic serum conditions compared to normal serum + VEH.

Figure 4.

Figure 4.

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Protein expression of metabolism and transport genes (A) CYP2R1, (B) CYP24A1, (C) P-gp, and (D) OATP4C1 in human proximal tubule epithelial cells (hPTECs). Protein expression was assessed after 24-hour exposure to normal or uraemic serum followed by treatment for six days with (1) vehicle control (VEH), (2) 240 nM VitD3, (3) 100 nM 1,25-VitD3, or (4) 240 nM 1,25-VitD3. The relative protein expression values were compared to normal serum + VEH treated cells. β-actin was used as a loading control. Data are expressed as means ± SEM of 3 replicates and analysed by analysis of variance (two-way ANOVA) followed by Tukey’s post-hoc test. **p < 0.01 compared to normal serum + VEH, #p < 0.05 compared to normal serum + treatment with the same VitD analogue.

CYP2R1 activity assessments

CYP2R1 activity was evaluated by measuring concentrations of VitD3 and 25-VitD3 in media and cell lysates from hPTECs after incubation with 10% normal or uraemic serum for 24 h followed by six days of treatment with VEH, 240 nM VitD3 or 240 nM 1,25-VitD3 and then incubated with 100 nM VitD3 as a CYP2R1 probe. Given that the high (240 nM) dose of VitD3 and 1,25-VitD3 showed the greatest influence on CYP2R1 gene and protein expression, this dose was selected for assessment of functional activity. AUC0–4h was calculated for VitD3 and 25-VitD3 and metabolic ratio was calculated by dividing 25-VitD3 AUC0–4h by VitD3 AUC0–4h (Table 1). VitD3 AUC0–4h was reduced in cell lysates treated with uraemic serum + 240 nM 1,25-VitD3 compared to normal serum + VEH (12.5 ± 0.50 ngh/mL vs. 17.6 ± 1.21 ngh/mL, p < 0.01). Serum or treatment with VitD analogues did not affect VitD3 AUC0–4h in media or 25-VitD3 AUC0–4h in cell lysates or media. No statistical differences in metabolic ratio were observed in cell lysates or media.

Table 1.

VitD3 and 25-VitD3 AUC and metabolic ratios in hPTECs.

Matrix Incubation conditions VitD3 AUC0–4 h (ng·h/mL) 25-VitD3 AUC0–4 h (ng·h/mL) Metabolic Ratio (25-VitD3:VitD3)
Cell Normal serum
Lysates  2% VEH 17.6 ± 1.21 19.8 ± 3.73 1.15 ± 0.29
 240 nM VitD3 14.7 ± 0.22 20.7 ± 3.78 1.41 ± 0.28
 240 nM 1,25-VitD3 15.2 ± 0.28 25.8 ± 0.14 1.70 ± 0.02
Uraemic serum
 2% VEH 14.5 ± 0.32 21.5 ± 0.77 1.49 ± 0.09
 240 nM VitD3 14.5 ± 0.78 21.4 ± 1.18 1.49 ± 0.16
 240 nM 1,25-VitD3 12.5 ± 0.50* 23.9 ± 1.14 1.91 ± 0.02
Normal serum
Media  2% VEH 14.1 ± 2.45 25.0 ± 0.64 1.83 ± 0.27
 240 nM VitD3 13.5 ± 0.90 20.3 ± 5.29 1.54 ± 0.49
 240 nM 1,25-VitD3 14.9 ± 0.34 23.0 ± 0.57 1.54 ± 0.07
Uraemic serum
 2% VEH 15.2 ± 1.26 24.2 ± 1.47 1.61 ± 0.23
 240 nM VitD3 15.1 ± 1.18 22.0 ± 0.96 1.47 ± 0.05
 240 nM 1,25-VitD3 13.4 ± 1.10 21.0 ± 3.49 1.60 ± 0.39
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Data was obtained after 24 h incubation with normal or uraemic serum and then treatment with vehicle control (VEH), 240 nM VitD3, or 240 nM 1,25-VitD3 for six days followed by incubation with 100 nM VitD3 (CYP2R1 probe) for 1, 2, or 4 h.

Data are mean ± SEM from duplicate experiments.

Abbreviations- VEH, vehicle control; VitD3, cholecalciferol; 25-VitD3, calcidiol; AUC, area under the concentration time curve. Data were analysed by analysis of variance (two-way ANOVA) followed by Tukey’s post-hoc test.

*

p < 0.05 compared with normal serum + VEH.

VEH: vehicle control; VitD3: cholecalciferol; 25-VitD3: calcidiol; AUC: area under the concentration time curve; hPTEC: human proximal tubule epithelial cells.

Transport assessments

The transport of [3H]digoxin (P-gp and OATP probe, Figure 5) was assessed in hPTECs exposed to normal or uraemic serum for 24 h and then treated with 240 nM 1,25-VitD3 for six days. Given that the 240 nM 1,25-VitD3 dose showed the greatest influence on gene and protein expression of ABCB1/P-gp and SLCO4C1/OATPC1, it was selected to assess the influence on transporter functional activity. B→A transport was similar to A→B transport. Differences in transport were only significant at the 1-h time point. One hour post digoxin dosing, transport in the A→B direction resulted in 21.1 ± 1.32% translocation with normal serum + VEH treatment compared to 23.3 ± 1.29% translocation following uraemic serum + VEH treatment (p < 0.01) (Figure 5(A)). One hour post digoxin dosing, transport in the B→A direction was greater following normal serum + 240 nM 1,25-VitD3 treatment (33.2 ± 2.0%) compared to normal serum + VEH (29.9 ± 2.4%) (p < 0.05) (Figure 5(B)). Overall, Papp B→A was greater than Papp A→B and did not differ across treatments. Following normal serum + VEH treatment, the efflux ratio was 2.7, indicating efflux transport. Uraemic serum and 1,25-VitD3 did not have a significant effect on efflux ratio (Table 2).

Figure 5.

Figure 5.

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Transporter function in human proximal tubule cells (hPTECs). After 24-hour exposure to normal or uraemic serum followed by treatment for six days with vehicle control (VEH) or 240 nM 1,25-VitD3 [3H]Digoxin transport was measured for 1, 2, 4, and 6 h. Experiments were performed in triplicate. (A) A→B represents transport from the apical to the basolateral membrane, (B) B→A represents transport from the basolateral to the apical membrane. Data are expressed as means ± SEM of 3 replicates and analysed by analysis of variance (two-way ANOVA) followed by Tukey’s post-hoc test where *p < 0.05 and **p < 0.01.

Table 2.

Apparent permeability coefficients (Papp)) and efflux ratio of [3H]digoxin across the hPTEC monolayer.

Treatment Papp (x10–6cm/s)
 Efflux Ratio
A → B B → A
Normal Serum + VEH 4.1 ± 0.27 10.9 ± 0.14 2.7
Normal Serum + 1,25-VitD3 3.8 ± 0.07 10.6 ± 0.21 2.8
Uraemic Serum + VEH 3.9 ± 0.14 11.1 ± 0.64 2.8
Uraemic Serum + 1,25-VitD3 3.8 ± 0.12 11.0 ± 0.24 2.9
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Data was obtained after 24 h incubation with normal or uraemic serum and then treatment with vehicle control (VEH) or 240 nM 1,25-VitD3 for six days followed by incubation with [3H]Digoxin for 1, 2, 4, and 6h.

Data are presented as mean ± SEM in triplicates. Data were analysed by analysis of variance (two-way ANOVA) followed by Tukey’s post-hoc test. VEH: vehicle control; VitD3: cholecalciferol; 1,25-VitD3: calcitriol; hPTEC: human proximal tubule epithelial cells; A→B: transport from apical to the basolateral membrane; B→A, transport from the basolateral to the apical membrane.

Discussion

The current study sought to fill gaps in the understanding of interactions between VitD and CYP metabolizing enzymes and drug transporter pathways in the setting of kidney disease or uraemia. The aims of the current investigation were to evaluate the influence of VitD3 and 1,25-VitD3 on the expression of VitD metabolizing CYP enzymes (CYP2R1, CYP27B1, and CYP24A1) and select drug transporters (ABCB1/P-gp, SLCO4C1/OATP4C1) in hPTECs exposed to normal serum and uraemic serum conditions. The key study results indicated differential expression of CYPs and transport genes and proteins, based on whether the cells were exposed to normal serum or uraemic serum, as well as which specific VitD analogue was administered.

Evidence from literature has shown increased cell viability with exposure to VitD analogs.(Ozerkan et al 2015) In the current study, under normal serum conditions, dosing of 1,25-VitD3 led to increased cell proliferation relative to VEH control (Figure 2). On the contrary, culture of hPTECs under uraemic serum resulted in no significant increase in cell proliferation as compared to exposures to normal serum treated with either low (100nM) or high (240 nM) doses of 1,25-VitD3. This suggests that uraemic serum can inhibit proliferative and potentially protective effects of 1,25-VitD3 on cells. Several studies have reported inhibition of cell proliferation in a variety of cell types upon exposure to uraemic toxins. (Mozar et al 2011, Pan et al 2017, Kawakami et al 2010). The exact mechanism(s) of the effects of uraemic toxins on cell proliferation remains unclear. However, proposed mechanisms have included direct inhibition of growth factors, increased production of reactive oxygen species, and endoplasmic reticulum stress (Mozar et al 2011, Pan et al 2017, Kawakami et al 2010). VitD was also previously reported to protect against cellular oxidative stress and decrease expression of inflammatory genes (Tohari et al 2016).

The current study sought to evaluate the influence of VitD moieties on drug transport and metabolism specifically in hPTECs, as CKD directly effects the kidneys. Previous studies have evaluated VitD metabolism in uraemic rats, (Michaud et al 2010, Helvig et al 2010) but published data on human kidney tubule cells was absent. Uraemic toxins accumulate in patients with moderate to severe CKD as kidney function declines and this has been documented to have effects on select pathways of drug metabolism and transport (Sun et al 2004, Naud et al 2011, Naud et al 2008, Nolin et al 2008). Various uraemic toxins, including 3-carboxy-4-methyl-5-propyl-2-furan-propanoic acid, indoxyl sulphate, hippuric acid, indole acetic acid, guanidinosuccinic acid, and indoxyl-β-D-glucuronide have been identified in the blood of patients with advanced kidney disease (Sun et al 2004, Naud et al 2011, Naud et al 2008, Nolin et al 2008). The current study exposed hPTECs to normal and uraemic serum in order decipher direct effects of the CKD environment on metabolism and transport pathways. The 24-hour exposure to normal or uraemic serum was selected based on previously published methods using rat hepatocytes and human microsomes (Michaud et al 2005, Michaud et al 2008, Volpe et al 2014). Serum was not co-incubated with VitD treatments in order to reduce protein binding and transcription factor interactions. The study results indicated differential expression of CYP metabolism and transport genes, which was dependent on the specific VitD analogue and serum to which the hPTECs were exposed.

CYP2R1 is the first enzyme encountered by VitD as it progresses along the metabolism pathway to its active metabolite. It is responsible for the 25-hydroxylation of VitD. While the liver is the primary site of VitD 25-hydroxylation in the human body, renal CYP2R1 has been purported to be capable of conversion of VitD to both 25-VitD3 and 1,25-VitD3 (Blomberg Jensen et al 2010, Urbschat et al 2013, Tucker et al 1973). The current study found that under normal serum CYP2R1 expression was increased with high dose (240 nM) VitD compounds relative to VEH (Figure 3(A)). CYP2R1 also had increased expression with uraemic serum under both low (100 nM) and high (240 nM) VitD analogue doses, with the exception of 240 nM VitD3 compared to normal serum + VEH. In fact, uraemic serum led to increases in CYP2R1 even in the absence of VitD analogues. CYP2R1 was also increased following treatment with 100 nM VitD3 under uraemic serum compared to the same treatment under normal serum. On the contrary, CYP2R1 protein was decreased with exposure to uraemic serum and 1,25-VitD3. Protein expression of CYP2R1 was lower under uraemic serum with low (100 nM) and high (240 nM) doses of 1,25-VitD3 compared the same VitD treatments under normal serum (Figure 4(A)) The discrepancy observed between gene and protein expression following treatment with uraemic serum and doses of 1,25-VitD3 in the current study could be due to post-translational modifications or differences in stability of mRNA vs. protein (Gedeon and Bokes 2012, Greenbaum et al 2003).

The functional activity of CYP2R1 can be assessed directly by measuring exposures of VitD and its immediate metabolic product 25-VitD and then calculating a metabolic ratio of the metabolite to the parent drug. The results of the current study suggest that hPTECs incubated with uraemic vs. normal serum, under pre-incubations (for potential induction) with low and high doses of VitD3 and 1,25-VitD3 demonstrated no overall differences in CYP2R1 activity (Table 1). AUC0–4h of 25-VitD3 and VitD3 did not differ across incubation conditions in cell media. However, the AUC0–4h of VitD3 was reduced in the cell lysates of hPTECs under uraemic serum + 1,25-VitD3 incubation conditions, suggesting a potential induction of CYP2R1 or metabolism through another enzymatic pathway that was not assessed in the current study. While there was not a statistically significant increase in 25-VitD3 AUC0–4h in cell lysates to correspond with the decrease in VitD3 AUC0–4h, there was a trend towards an increase in exposure to 25-VitD3. The influence of subsequent activation (CYP27B1) and catabolism (CYP24A1) to 1,25-VitD3 and 24,25-VitD3, respectively, on VitD3 and 25-VitD3 exposures were not assessed, as sensitive analytical testing for these metabolites in cell media and lysates were not available for the current study. Future studies with highly sensitive assays will be needed to determine the impact of subsequent metabolism on measured exposures to earlier substrates in the VitD metabolism cascade.

While CYP2R1 is an activating enzyme for VitD, CYP24A1 is a key enzyme responsible for catabolism of the active 1,25-VitD3 to the inactive 1,24,25-VitD3. It is present in all cells that express the vitamin D receptor (VDR) including intestinal enterocytes,(Baker et al 1988) osteoblasts (Narbaitz et al 1981), and renal tubules (Stumpf et al 1979, Berger et al 1988). Its primary role is to reduce active VitD (1,25-VitD3) concentrations to prevent high serum calcium levels (Jones et al 1998). In the current study, CYP24A1 expression was induced following treatment with both low (100 nM) and high (240 nM) doses of 1,25-VitD3 relative to VEH under normal serum conditions (Figure 3(B)). These finding were consistent with the literature reporting that 1,25-VitD3 concentrations are feedback regulated and increased levels of 1,25-VitD3 induce CYP24A1 (Bossé et al 2007, Horvath et al 2012, Deeb et al 2007). A study in HepG2 cells showed CYP24A1 expression increased 5,300-fold in response to treatment with a 4 nM dose of 1,25-VitD3 (Horvath et al 2012). This suggests the influence of 1,25-VitD3 on CYP24A1 expression may be dose and/or cell type dependent. In the current study, CYP24A1 expression was also increased, albeit to a less extent following treatment with low (100 nM) and high (240 nM) doses of 1,25-VitD3 under uraemic serum conditions relative to normal serum + VEH. A previous study in porcine proximal tubule kidney cells dosed with 1,25-VitD3 and parathyroid hormone (PTH), a confirmed uraemic toxin, resulted in suppressed CYP24A1 expression by reducing mRNA stability and enhancing degradation (Zierold et al 2001). CYP24A1 protein expression showed no changes under serum conditions or under exposures to VitD treatments in the current study (Figure 4(B)). Furthermore, a study in benign and malignant breast tissue and cell lines also reported discrepancies in CYP24A1 mRNA and protein expression and suggested alternative splicing of CYP24A1 could be the cause differences in mRNA and protein levels (Fischer et al 2009).

In addition to CYP metabolism pathways, drug transporters play a critical role in the disposition of many drugs (Morrissey et al 2013). As kidney transporters can be directly impacted in CKD, the functional implications could be significant. P-gp is an efflux transporter and is primarily found on the apical surface of epithelial cells in a variety of tissues, including hPTECs, and regulates intracellular exposures (Melaine et al 2002). P-gp is promiscuous and responsible for the transport of numerous drugs, including dexamethasone, cimetidine and fexofenadine (Cvetkovic et al 1999, Karyekar et al 2004, Ueda et al 1992). It is commonly identified in drug-drug interactions and is interrogated in drug development.(Zhang et al 2009, Fenner et al 2009) In the current study, under normal serum, the high (240 nM) dose of 1,25-VitD3 resulted in an increase in ABCB1 relative to VEH (Figure 3(C)). ABCB1 expression was also increased in hPTECs treated with a low (100 nM) dose of 1,25-VitD3 under normal or uraemic serum. P-gp protein changes were concordant in direction with mRNA expression, with an increase under high (240 nM) doses of 1,25-VitD3 and uraemic serum vs. normal serum (Figure 4(C)). Previous studies in several cell types (Caco-2, LS180, LS174T) have shown upregulation of ABCB1/P-gp expression under treatment with 1,25-VitD3 through vitamin D receptor (VDR) interactions (Aiba et al 2005, Fan et al 2009b, Tachibana et al 2009). Binding of 1,25-VitD3 to VDR triggers the formation of a heterodimer of VDR and retinoic X receptor (RXR). This heterodimer is then able to recognize and bind to the vitamin D response element (VDRE) in the promoter region of target DNA sequences. ABCB1 has been reported to contain multiple VDREs, supporting the gene’s regulation through VDR (Saeki et al 2008).

OATP4C1 is an uptake transporter localized to the basolateral membrane and is predominately expressed in the proximal tubules of the kidney (Sato et al 2017). OATP4C1 is involved in the transport of numerous compounds including digoxin, ouabain, methotrexate, and glycocholic acid (Sato et al 2017, Mikkaichi et al 2004b). In the current study, SLCO4C1/OATP4C1 mRNA expression was increased in hPTECs treated with high (240nM) dose 1,25-VitD3 and uraemic serum as compared to normal serum + VEH (Figure 3(D)). However, uraemic serum appeared to inhibit OATP4C1 expression in hPTECs treated with VEH or VitD analogues (Figure 4(D)). While the mechanism is unclear, it is possible that degradation of OATP4C1 may be circumventing the increase in gene expression to protect the cell from increasing intracellular concentrations of compounds under uraemia.

The functional relevance of uraemic serum and VitD compounds on transport were assessed in the current study using [3H] digoxin as a probe compound. Digoxin is a well-known substrate of P-gp and in the kidney proximal tubule cells, OATP4C1 is believed to be the major uptake transporter of digoxin on the basolateral membrane (Mikkaichi et al 2004a, Mikkaichi et al 2004b). In the current study, transport experiments were assessed at 1, 2, 4, and 6 h after digoxin treatment to evaluate apical-to-basolateral transport (apically treated and basolaterally measured, A→B, Figure 5(A)) and basolateral-to-apical transport (basolaterally treated and apically measured, B→A, Figure 5(B)). While basolateral to apical transport of [3H] digoxin was higher than apical to basolateral transport in hPTECs exposed to 1,25-VitD3, these overall differences failed to reach statistical significance. Significant differences in digoxin translocation were noted at the 1 h assessment time, but the importance of these differences is uncertain. Calculation of the efflux ratio (Papp, B→A to Papp, A→B) under the experimental conditions failed to support any differences. Overall, functional assessments of transport in hPTEC, showed no overall changes regardless of normal or uraemic serum conditions or exposure to 1,25-VitD3. Therefore, any gene or protein expression changes in transporters due to serum conditions were not observed to result in changes to overall transporter activity. The transport of the P-gp and MRP4 substrates adefovir dipivoxil and its metabolite adefovir was previously reported in Caco-2 cells exposed to 1,25-VitD3 treatment (Maeng et al 2012). Increased P-gp and MRP4 transport activity through assessment of basolateral to apical transport of adefovir dipivoxil was demonstrated after 3 day treatment with 1,25-VitD3 (100 nM). This may suggest treatment with VitD analogues have differential effects in the type of cells evaluated. However, in the adefovir study, Caco-2 cells were not incubated with normal or uraemic serum as in the current study which may also contribute to the observed differences secondary to effects on transcription factors. A study in hepatocellular carcinoma cells, showed differential transcription of 22–32% of genes evaluated following exposure to human serum versus foetal bovine serum (Steenbergen et al 2018). The presence and function of sodium dependent transporters located on the basolateral or apical membrane of hPTECs may have obscured the effects of incubation conditions on digoxin transport in the current study (Taub et al 2011). The discordance between gene or protein expression changes to transporters due to serum conditions and subsequent lack of changes in functional transporter activity in the current study requires additional investigation.

Conclusions

The current study fill gaps in the understanding of how VitD may interact with CYP metabolizing enzymes and transporter pathways (P-gp and OATP4C1) under the influence of kidney disease or uraemia. The key study findings indicated differential expression of CYPs and transport genes and proteins, dependent upon whether the cells were exposed to normal serum or uraemic serum, and which specific VitD treatment (and dose) was administered. Additional studies are needed to understand lack of concordance in direction of gene and protein expression results observed in this study, which may include assessment of post-transcriptional or -translational mechanisms (Baldi and Long 2001, Szallasi 1999, Greenbaum et al 2003). However, observations from this study could be extended to other transport and metabolism substrates in hPTECs and may aid in drug development and clinical applications in the CKD population.

Funding

This work was supported by National Institute of General Medical Sciences.

Footnotes

Disclosure statement

JMT receives royalties from Alexion Pharmaceuticals, Inc. and is a consultant for Q32 Bio, Inc., a company developing complement inhibitors. He also holds stock and will receive royalty income from Q32 Bio, Inc.

References

  1. Aiba T, et al. , 2005. The effects of culture conditions on CYP3A4 and MDR1 mRNA induction by 1α, 25-dihydroxyvitamin D3 in human intestinal cell lines, Caco-2 and LS180. Drug metabolism and pharmacokinetics, 20 (4), 268–274. [DOI] [PubMed] [Google Scholar]
  2. Baker AR, et al. , 1988. Cloning and expression of full-length cDNA encoding human vitamin D receptor. Proceedings of the national academy of sciences of the United States of America, 85 (10), 3294–3298. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Baldi P, and Long AD, 2001. A Bayesian framework for the analysis of microarray expression data: regularized t -test and statistical inferences of gene changes. Bioinformatics, 17 (6), 509–519. [DOI] [PubMed] [Google Scholar]
  4. Berger U, et al. , 1988. Immunocytochemical detection of 1,25-dihydroxyvitamin D receptors in normal human tissues. The journal of clinical endocrinology and metabolism, 67 (3), 607–613. [DOI] [PubMed] [Google Scholar]
  5. Bhan I, et al. , 2010. Clinical measures identify vitamin D deficiency in dialysis. Clinical journal of the American society of nephrology, 5 (3), 460–467. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Blomberg Jensen M, et al. , 2010. Expression of the vitamin D receptor, 25-hydroxylases, 1alpha-hydroxylase and 24-hydroxylase in the human kidney and renal clear cell cancer. The journal of steroid biochemistry and molecular biology, 121 (1–2), 376–382. [DOI] [PubMed] [Google Scholar]
  7. Bossé Y, Maghni K, and Hudson TJ, 2007. 1α, 25-Dihydroxy-vitamin D3 stimulation of bronchial smooth muscle cells induces autocrine, contractility, and remodeling processes. Physiological genomics, 29 (2), 161–168. [DOI] [PubMed] [Google Scholar]
  8. Cheng S, and Coyne D, 2007. Vitamin D and outcomes in chronic kidney disease. Current opinion in nephrology and hypertension, 16 (2), 77–82. [DOI] [PubMed] [Google Scholar]
  9. Chow EC, et al. , 2011. Comparative effects of doxercalciferol (1alpha-hydroxyvitamin D(2)) versus calcitriol (1alpha,25-dihydroxyvitamin D(3)) on the expression of transporters and enzymes in the rat in vivo. Journal of pharmaceutical sciences, 100 (4), 1594–1604. [DOI] [PubMed] [Google Scholar]
  10. Chow EC, et al. , 2010. Effects of 1alpha,25-dihydroxyvitamin D3 on transporters and enzymes of the rat intestine and kidney in vivo. Biopharmaceutics & drug disposition, 31 (1), 91–108. [DOI] [PubMed] [Google Scholar]
  11. Coresh J, et al. , 2007. Prevalence of chronic kidney disease in the United States. Jama, 298 (17), 2038–2047. [DOI] [PubMed] [Google Scholar]
  12. Cvetkovic M, et al. , 1999. OATP and P-glycoprotein transporters mediate the cellular uptake and excretion of fexofenadine. Drug metabolism and disposition: the biological fate of chemicals, 27 (8), 866–871. [PubMed] [Google Scholar]
  13. Deeb KK, Trump DL, and Johnson CS, 2007. Vitamin D signalling pathways in cancer: potential for anticancer therapeutics. Nature reviews. cancer, 7 (9), 684–700. [DOI] [PubMed] [Google Scholar]
  14. Fan J, et al. , 2009a. Up-regulation of transporters and enzymes by the vitamin D receptor ligands, 1alpha,25-dihydroxyvitamin D3 and vitamin D analogs, in the Caco-2 cell monolayer. Journal of pharmacology and experimental therapeutics, 330 (2), 389–402. [DOI] [PubMed] [Google Scholar]
  15. Fan J, et al. , 2009b. Up-regulation of transporters and enzymes by the vitamin D receptor ligands, 1α, 25-dihydroxyvitamin D3 and vitamin D analogs, in the Caco-2 cell monolayer. Journal of pharmacology and experimental therapeutics, 330 (2), 389–402. [DOI] [PubMed] [Google Scholar]
  16. Fenner KS, et al. , 2009. Drug-drug interactions mediated through P-glycoprotein: clinical relevance and in vitro-in vivo correlation using digoxin as a probe drug. Clinical pharmacology & therapeutics, 85 (2), 173–181. [DOI] [PubMed] [Google Scholar]
  17. Fischer D, et al. , 2009. Vitamin D-24-hydroxylase in benign and malignant breast tissue and cell lines. Anticancer research, 29 (9), 3641–3645. [PubMed] [Google Scholar]
  18. Gedeon T, and Bokes P, 2012. Delayed protein synthesis reduces the correlation between mRNA and protein fluctuations. Biophysical journal, 103 (3), 377–385. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. González EA, et al. , 2004. Vitamin D insufficiency and deficiency in chronic kidney disease. American journal of nephrology, 24 (5), 503–510. [DOI] [PubMed] [Google Scholar]
  20. Greenbaum D, et al. , 2003. Comparing protein abundance and mRNA expression levels on a genomic scale. Genome biology, 4 (9), 117. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Heaney RP, 2008. Vitamin D in health and disease. Clinical journal of the American society of nephrology, 3 (5), 1535–1541. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Helvig CF, et al. , 2010. Dysregulation of renal vitamin D metabolism in the uremic rat. Kidney international, 78 (5), 463–472. [DOI] [PubMed] [Google Scholar]
  23. Hirata M, et al. , 2002. 22-Oxacalcitriol prevents progressive glomerulosclerosis without adversely affecting calcium and phosphorus metabolism in subtotally nephrectomized rats. Nephrology, dialysis, transplantation, 17 (12), 2132–2137. [DOI] [PubMed] [Google Scholar]
  24. Holick MF, 2007. Vitamin D deficiency. New England journal of medicine, 357 (3), 266–281. [DOI] [PubMed] [Google Scholar]
  25. Holick MF, et al. ,; Endocrine Society. 2011. Evaluation, treatment, and prevention of vitamin D deficiency: an Endocrine Society clinical practice guideline. The journal of clinical endocrinology and metabolism, 96 (7), 1911–1930. [DOI] [PubMed] [Google Scholar]
  26. Horvath E, et al. , 2012. Marked increase of CYP24A1 mRNA level in hepatocellular carcinoma cell lines following vitamin D administration. Anticancer research, 32, 4791–4796. [PubMed] [Google Scholar]
  27. Jones G, 2007. Expanding role for vitamin D in chronic kidney disease: importance of blood 25-OH-D levels and extra-renal 1alpha-hydroxylase in the classical and nonclassical actions of 1alpha,25-dihydroxyvitamin D(3). Seminars in dialysis, 20 (4), 316–324. [DOI] [PubMed] [Google Scholar]
  28. Jones G, Strugnell SA, and Deluca HF, 1998. Current understanding of the molecular actions of vitamin D. Physiological reviews, 78 (4), 1193–1231. [DOI] [PubMed] [Google Scholar]
  29. Kandula P, et al. , 2011. Vitamin D supplementation in chronic kidney disease: a systematic review and meta-analysis of observational studies and randomized controlled trials. Clinical journal of the american society of nephrology, 6 (1), 50–62. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Karyekar CS, et al. , 2004. Renal interaction between itraconazole and cimetidine. The Journal of Clinical Pharmacology, 44 (8), 919–927. [DOI] [PubMed] [Google Scholar]
  31. Kawakami T, et al. , 2010. Indoxyl sulfate inhibits proliferation of human proximal tubular cells via endoplasmic reticulum stress. American journal of physiology. Renal physiology, 299 (3), F568–F576. [DOI] [PubMed] [Google Scholar]
  32. Li YC, 2010. Renoprotective effects of vitamin D analogs. Kidney international, 78 (2), 134–139. [DOI] [PubMed] [Google Scholar]
  33. Livak KJ, and Schmittgen TD, 2001. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) Method. Methods, 25 (4), 402–408. [DOI] [PubMed] [Google Scholar]
  34. Maeng HJ, et al. , 2012. Effects of 1α,25-dihydroxyvitamin D3 on transport and metabolism of adefovir dipivoxil and its metabolites in Caco-2 cells . European journal of pharmaceutical sciences, 46 (3), 149–166. [DOI] [PubMed] [Google Scholar]
  35. Manley HJ, Drayer DK, and Muther RS, 2003. Medication-related problem type and appearance rate in ambulatory hemodialysis patients. BMC nephrology, 4, 10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Massry SG, et al. , 2003. K/DOQI clinical practice guidelines for bone metabolism and disease in chronic kidney disease. American journal of kidney diseases, 42, S1–S201. [PubMed] [Google Scholar]
  37. Mehrotra R, et al. , 2008. Hypovitaminosis D in chronic kidney disease. Clinical journal of the american society of nephrology, 3 (4), 1144–1151. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Mehrotra R, et al. , 2009. Chronic kidney disease, hypovitaminosis D, and mortality in the United States. Kidney international, 76 (9), 977–983. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Mehrotra R, et al. ,; on behalf of the Kidney Early Evaluation Program (KEEP) Investigators. 2013. No independent association of serum phosphorus with risk for death or progression to end-stage renal disease in a large screen for chronic kidney disease. Kidney International, 84 (5), 989–997. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Melaine N, et al. , 2002. Multidrug resistance genes and p-glycoprotein in the testis of the rat, mouse, Guinea pig, and human. Biology of Reproduction, 67 (6), 1699–1707. [DOI] [PubMed] [Google Scholar]
  41. Michaud J, et al. , 2005. Effects of serum from patients with chronic renal failure on rat hepatic cytochrome P450. British journal of pharmacology, 144 (8), 1067–1077. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Michaud J, et al. , 2010. Reduced hepatic synthesis of calcidiol in uremia. Journal of the American society of nephrology, 21 (9), 1488–1497. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Michaud J, et al. , 2008. Effect of hemodialysis on hepatic cytochrome P450 functional expression. Journal of pharmacological sciences, 108 (2), 157–163. [DOI] [PubMed] [Google Scholar]
  44. Mikkaichi T, et al. , 2004a. Isolation and characterization of a digoxin transporter and its rat homologue expressed in the kidney. Proceedings of the national Academy of sciences, 101, 3569–3574. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Mikkaichi T, et al. , 2004b. The organic anion transporter (OATP) family. Drug metabolism and pharmacokinetics, 19 (3), 171–179. [DOI] [PubMed] [Google Scholar]
  46. Morrissey KM, et al. , 2013. Renal transporters in drug development. Annual review of pharmacology and toxicology, 53, 503–529. [DOI] [PubMed] [Google Scholar]
  47. Mozar A, et al. , 2011. Uremic toxin indoxyl sulfate inhibits human vascular smooth muscle cell proliferation. Therapeutic apheresis and dialysis, 15 (2), 135–139. [DOI] [PubMed] [Google Scholar]
  48. Narbaitz R, Stumpf WE, and Sar M, 1981. The role of autoradiographic and immunocytochemical techniques in the clarification of sites of metabolism and action of vitamin D. The journal of histochemistry and cytochemistry, 29 (1), 91–100. [DOI] [PubMed] [Google Scholar]
  49. Naud J, et al. , 2011. Effects of chronic renal failure on kidney drug transporters and cytochrome P450 in rats. Drug metabolism and disposition, 39 (8), 1363–1369. [DOI] [PubMed] [Google Scholar]
  50. Naud J, et al. , 2008. Effects of chronic renal failure on liver drug transporters. Drug metabolism and disposition, 36 (1), 124–128. [DOI] [PubMed] [Google Scholar]
  51. Nolin T, et al. , 2008. Emerging evidence of the impact of kidney disease on drug metabolism and transport. Clinical pharmacology and therapeutics, 83 (6), 898–903. [DOI] [PubMed] [Google Scholar]
  52. Ozerkan D, Ozsoy N, and Yilmaz E, 2015. Vitamin D and melatonin protect the cell’s viability and ameliorate the CCl4 induced cytotoxicity in HepG2 and Hep3B hepatoma cell lines. Cytotechnology, 67, 995–1002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Pan L, et al. , 2017. Antiproliferation effect of the uremic toxin para-cresol on endothelial progenitor cells is related to its antioxidant activity. Molecular medicine reports, 15 (4), 2308–2312. [DOI] [PMC free article] [PubMed] [Google Scholar] [Retracted]
  54. Pramyothin P, and Holick MF, 2012. Vitamin D supplementation: guidelines and evidence for subclinical deficiency. Current opinion in gastroenterology, 28 (2), 139–150. [DOI] [PubMed] [Google Scholar]
  55. Saeki M, et al. , 2008. Identification of the functional vitamin D response elements in the human MDR1 gene. Biochemical pharmacology, 76 (4), 531–542. [DOI] [PubMed] [Google Scholar]
  56. Sato T, et al. , 2017. Potential drug interactions mediated by renal organic anion transporter OATP4C1. Journal of pharmacology and experimental therapeutics, 362 (2), 271–277. [DOI] [PubMed] [Google Scholar]
  57. Schwarz U, et al. , 1998. Effect of 1,25 (OH)2 vitamin D3 on glomerulosclerosis in subtotally nephrectomized rats. Kidney international, 53 (6), 1696–1705. [DOI] [PubMed] [Google Scholar]
  58. Steenbergen R, et al. , 2018. Establishing normal metabolism and differentiation in hepatocellular carcinoma cells by culturing in adult human serum. Scientific reports, 8 (1), 11685. [DOI] [PMC free article] [PubMed] [Google Scholar]
  59. Stumpf WE, et al. , 1979. Target cells for 1,25-dihydroxyvitamin D3 in intestinal tract, stomach, kidney, skin, pituitary, and parathyroid. Science, 206 (4423), 1188–1190. [DOI] [PubMed] [Google Scholar]
  60. Sun H, et al. , 2004. Effects of uremic toxins on hepatic uptake and metabolism of erythromycin. Drug metabolism and disposition, 32 (11), 1239–1246. [DOI] [PubMed] [Google Scholar]
  61. Szallasi Z, 1999. Genetic network analysis in light of massively parallel biological data acquisition. Pacific Symposium on Biocomputing, 4, 5–16. [DOI] [PubMed] [Google Scholar]
  62. Tachibana S, et al. , 2009. Involvement of vitamin D receptor in the intestinal induction of human ABCB1. Drug metabolism and disposition, 37 (8), 1604–1610. [DOI] [PubMed] [Google Scholar]
  63. Tan X, Wen X, and Liu Y, 2008. Paricalcitol inhibits renal inflammation by promoting vitamin D receptor-mediated sequestration of NF-kappaB signaling. Journal of the American society of nephrology, 19 (9), 1741–1752. [DOI] [PMC free article] [PubMed] [Google Scholar]
  64. Taub ME, et al. , 2011. Digoxin is not a substrate for organic anion-transporting polypeptide transporters OATP1A2, OATP1B1, OATP1B3, and OATP2B1 but is a substrate for a sodium-dependent transporter expressed in HEK293 cells. Drug metabolism and disposition, 39 (11), 2093–2102. [DOI] [PubMed] [Google Scholar]
  65. Thomas MK, et al. , 1998. Hypovitaminosis D in medical inpatients. New England journal of medicine, 338 (12), 777–783. [DOI] [PubMed] [Google Scholar]
  66. Tohari AM, Zhou X, and Shu X, 2016. Protection against oxidative stress by vitamin D in cone cells. Cell Biochemistry and Function, 34 (2), 82–94. [DOI] [PubMed] [Google Scholar]
  67. Tucker G 3rd, Gagnon RE, and Haussler MR, 1973. Vitamin D 3 −25-hydroxylase: tissue occurrence and apparent lack of regulation. Archives of biochemistry and biophysics, 155 (1), 47–57. [DOI] [PubMed] [Google Scholar]
  68. Ueda K, et al. , 1992. Human P-glycoprotein transports cortisol, aldosterone, and dexamethasone, but not progesterone. Journal of biological chemistry, 267 (34), 24248–24252. [PubMed] [Google Scholar]
  69. Urbschat A, et al. , 2013. Vitamin D hydroxylases CYP2R1, CYP27B1 and CYP24A1 in renal cell carcinoma. European journal of clinical investigation, 43 (12), 1282–1290. [DOI] [PubMed] [Google Scholar]
  70. Volpe DA, et al. , 2014. Effect of uremic serum and uremic toxins on drug metabolism in human microsomes. Regulatory toxicology and pharmacology, 68 (2), 297–303. [DOI] [PubMed] [Google Scholar]
  71. Zhang L, et al. , 2009. Predicting drug-drug interactions: an FDA perspective. The AAPS journal, 11 (2), 300–306. [DOI] [PMC free article] [PubMed] [Google Scholar]
  72. Zhang Z, et al. , 2008. Renoprotective role of the vitamin D receptor in diabetic nephropathy. Kidney international, 73 (2), 163–171. [DOI] [PubMed] [Google Scholar]
  73. Zierold C, Mings JA, and Deluca HF, 2001. Parathyroid hormone regulates 25-hydroxyvitamin D(3)-24-hydroxylase mRNA by altering its stability. Proceedings of the national academy of sciences of the United States of America, 98 (24), 13572–13576. [DOI] [PMC free article] [PubMed] [Google Scholar]

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