Pannexin‐1 mediates fluid shear stress‐sensitive purinergic signaling and cyst growth in polycystic kidney disease
2020; Wiley; Volume: 34; Issue: 5 Linguagem: Inglês
10.1096/fj.201902901r
ISSN1530-6860
AutoresEric H. J. Verschuren, Juan Pablo Rigalli, Charlotte Castenmiller, Meike U. Rohrbach, René J.M. Bindels, Dorien J.M. Peters, Francisco J. Arjona, Joost G.J. Hoenderop,
Tópico(s)Renal and related cancers
ResumoThe FASEB JournalVolume 34, Issue 5 p. 6382-6398 RESEARCH ARTICLEOpen Access Pannexin-1 mediates fluid shear stress-sensitive purinergic signaling and cyst growth in polycystic kidney disease Eric H. J. Verschuren, Eric H. J. Verschuren Department of Physiology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, the NetherlandsSearch for more papers by this authorJuan P. Rigalli, Juan P. Rigalli Department of Physiology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, the NetherlandsSearch for more papers by this authorCharlotte Castenmiller, Charlotte Castenmiller Department of Physiology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, the NetherlandsSearch for more papers by this authorMeike U. Rohrbach, Meike U. Rohrbach Department of Physiology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, the NetherlandsSearch for more papers by this authorRené J. M. Bindels, René J. M. Bindels Department of Physiology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, the NetherlandsSearch for more papers by this authorDorien J. M. Peters, Dorien J. M. Peters Department of Human Genetics, Leiden University Medical Center, Leiden, the NetherlandsSearch for more papers by this authorFrancisco J. Arjona, Francisco J. Arjona Department of Physiology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, the NetherlandsSearch for more papers by this authorJoost G. J. Hoenderop, Corresponding Author Joost G. J. Hoenderop Joost.Hoenderop@radboudumc.nl Department of Physiology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, the Netherlands Correspondence Joost G. J. Hoenderop, Department of Physiology (286), Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, P.O. Box 9101, 6500 HB Nijmegen, the Netherlands. Email: Joost.Hoenderop@radboudumc.nlSearch for more papers by this author Eric H. J. Verschuren, Eric H. J. Verschuren Department of Physiology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, the NetherlandsSearch for more papers by this authorJuan P. Rigalli, Juan P. Rigalli Department of Physiology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, the NetherlandsSearch for more papers by this authorCharlotte Castenmiller, Charlotte Castenmiller Department of Physiology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, the NetherlandsSearch for more papers by this authorMeike U. Rohrbach, Meike U. Rohrbach Department of Physiology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, the NetherlandsSearch for more papers by this authorRené J. M. Bindels, René J. M. Bindels Department of Physiology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, the NetherlandsSearch for more papers by this authorDorien J. M. Peters, Dorien J. M. Peters Department of Human Genetics, Leiden University Medical Center, Leiden, the NetherlandsSearch for more papers by this authorFrancisco J. Arjona, Francisco J. Arjona Department of Physiology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, the NetherlandsSearch for more papers by this authorJoost G. J. Hoenderop, Corresponding Author Joost G. J. Hoenderop Joost.Hoenderop@radboudumc.nl Department of Physiology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, Nijmegen, the Netherlands Correspondence Joost G. J. Hoenderop, Department of Physiology (286), Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, P.O. Box 9101, 6500 HB Nijmegen, the Netherlands. Email: Joost.Hoenderop@radboudumc.nlSearch for more papers by this author First published: 11 March 2020 https://doi.org/10.1096/fj.201902901RCitations: 2 Francisco J. Arjona and Joost G. J. Hoenderop are contributed equally to this work. AboutSectionsPDF ToolsRequest permissionExport citationAdd to favoritesTrack citation ShareShare Give accessShare full text accessShare full-text accessPlease review our Terms and Conditions of Use and check box below to share full-text version of article.I have read and accept the Wiley Online Library Terms and Conditions of UseShareable LinkUse the link below to share a full-text version of this article with your friends and colleagues. Learn more.Copy URL Share a linkShare onFacebookTwitterLinkedInRedditWechat Abstract Tubular ATP release is regulated by mechanosensation of fluid shear stress (FSS). Polycystin-1/polycystin-2 (PC1/PC2) functions as a mechanosensory complex in the kidney. Extracellular ATP is implicated in polycystic kidney disease (PKD), where PC1/PC2 is dysfunctional. This study aims to provide new insights into the ATP signaling under physiological conditions and PKD. Microfluidics, pharmacologic inhibition, and loss-of-function approaches were combined to assess the ATP release in mouse distal convoluted tubule 15 (mDCT15) cells. Kidney-specific Pkd1 knockout mice (iKsp-Pkd1−/−) and zebrafish pkd2 morphants (pkd2-MO) were as models for PKD. FSS-exposed mDCT15 cells displayed increased ATP release. Pannexin-1 inhibition and knockout decreased FSS-modulated ATP release. In iKsp-Pkd1−/− mice, elevated renal pannexin-1 mRNA expression and urinary ATP were observed. In Pkd1−/− mDCT15 cells, elevated ATP release was observed upon the FSS mechanosensation. In these cells, increased pannexin-1 mRNA expression was observed. Importantly, pannexin-1 inhibition in pkd2-MO decreased the renal cyst growth. Our results demonstrate that pannexin-1 channels mediate ATP release into the tubular lumen due to pro-urinary flow. We present pannexin-1 as novel therapeutic target to prevent the renal cyst growth in PKD. Abbreviations Abcc6 ATP binding cassette subfamily c member 6 BB-FCF brilliant blue-FCF Cx30 connexin30 Cx30.3 connexin30.3 Cx37 connexin37 Entpd2 ectonucleoside triphosphate diphosphohydrolase 2 Entpd3 ectonucleoside triphosphate diphosphohydrolase 3 FBS fetal bovine serum FSS fluid shear stress Gapdh glyceraldehyde 3-phosphate dehydrogenase MO translation blocking morpholino Nt5e ecto-5′-nucleotidase Panx1 pannexin-1 PC1 polycystin-1 PC2 polycystin-2 PKD polycystic kidney disease Pkd1 polycystic kidney disease 1 Pkd2 polycystic kidney disease 2 Ptgs2 prostaglandin-endoperoxide synthase 2 1 INTRODUCTION Mutations in the genes polycystic kidney disease 1 (PKD1) and polycystic kidney disease 2 (PKD2) lead to polycystic kidney disease (PKD). PKD is characterized by increased cell proliferation, fluid accumulation, and altered extracellular matrix synthesis in the kidney. These characteristics lead to the renal cyst formation and growth, predominantly in the distal tubules and collecting duct (CD), and ultimately to end-stage kidney disease.1-4 PKD is one of the most common inherited renal diseases and accounts for ~10% of all patients on renal replacement therapy worldwide.5, 6 The Pkd1 gene encodes for the protein polycystin-1 (PC1), which locates to the primary cilium and plasma membrane of renal tubular epithelial cells. PC1 forms a complex with polycystin-2 (PC2, encoded by Pkd2), which is a nonselective cation channel.7-11 PC1 is suggested to act as a mechanosensor of fluid shear stress (FSS) generated by pro-urine flow, regulating the physiological responses in renal tubular epithelial cells.12, 13 In PKD, extracellular ATP plays an important role in disease progression.14, 15 Under physiological conditions, extracellular ATP activates ionotropic P2X and metabotropic P2Y receptors at the luminal cell surface to regulate and maintain the kidney function.16 In recent years, autocrine and paracrine effects of the purinergic signaling have been suggested to provide a detrimental acceleration of cyst growth in PKD.17-20 It has been shown that the renal cyst fluid in PKD rats and PKD patients contains high ATP levels.21, 22 Also, cystic PKD cell cultures display increased ATP release at basal conditions.23 Furthermore, previous studies have related purinergic receptors P2Y2 and P2X7 to cyst growth in an in vitro cystic model, and in vivo, in zebrafish and mouse PKD models.23-26 Thus far, translation of these findings into therapeutic applications for PKD related to purinergic signaling are limited.24 Conversely, approaches that did reach clinical research stages for PKD involve vasopressin antagonists, mTOR inhibitors, and somatostatin analogues. However, these approaches are not a definitive curative treatment option.1, 25-29 An improved understanding of the molecular mechanisms behind ATP signaling in PKD may promote the development of novel therapeutic strategies for this disease. ATP can be released from renal epithelial cells upon the mechanical stimulation triggered by variable pro-urine flow.30, 31 Yet, the ATP extrusion mechanisms mediating flow-sensitive ATP release as well as their regulation are not fully elucidated.32, 33 The extracellular ATP in the tubule lumen signals the apical membrane of renal cells to trigger an intracellular response which results in regulation of electrolyte and water transport.34, 35 In this study, we aimed to disclose the mechanisms regulating the apical ATP release in renal cells upon flow sensing. To this end, the urine of healthy human volunteers and cellular and organismal kidney models were interrogated. The relevance of these mechanisms for disease progression in the context of PKD was investigated, where purinergic signaling was anticipated to play a pathologic role. 2 MATERIALS AND METHODS 2.1 Ethics approval Urine samples of healthy human volunteers were collected after acute water loading experimentation in accordance with the principles expressed in the Declaration of Helsinki. All participants gave written informed consent, and the study protocol was approved by the Institutional Review Board of the Radboud University Medical Center (approval no. NL47178.091.13). The local animal experimental committee of the Leiden University Medical Center and the Commission Biotechnology in Animals of the Dutch Ministry of Agriculture approved the animal procedures performed. 2.2 Cell culture To assess the role of FSS on ATP release, μ-slide I0.4 and VI0.4 channels (Ibidi GmbH, Planegg, Germany) were used for ATP and gene expression experiments, respectively. Static ATP release was assessed using 12-well plates. Mouse distal convoluted tubule 15 (mDCT15) cells (gift from Dr Robert Hoover, Emory University, Atlanta, GA, USA), a model of the distal convoluted epithelial cell, were used in all experiments.36 mDCT15 cells (either wild type, Ift140−/− or Pkd1−/−) were seeded in the channels (μ-slide I0.4:1.2 × 105 cells/cm2 and μ-slide VI0.4: 5 × 104 cells/cm2) with DMEM/F-12 1:1 nutrient mixed media supplemented with 5% v/v fetal bovine serum (FBS, GE Healthcare, Little Chalfont, United Kingdom) and 10 μg/mL ciprofloxacin, and maintained at 37°C and 5% (v/v) CO2. The following day, cells were serum-starved for 24 hours to induce cell differentiation. Shortly before FSS experimentation, biocompatible silicone tubing (Ø0.8 mm, Ibidi GmbH, Planegg, Germany) was connected to a peristaltic pump (ISM931C, Ismatec, Wertheim, Germany) and the microfluidic channels. During microfluidic experiments serum-free DMEM/F-12 1:1 nutrient mixed media was used. 2.3 ATP measurements To assess the ATP release by mDCT15 cells due to FSS, cells were exposed during 1 minute to different physiologically relevant FSS levels (ie, 0.3, 0.6, and 1.2 dyn/cm2). Cells were also exposed to repetitive cycles of 0.3 dyn/cm2 (1 min) followed by 1.2 dyn/cm2 (1 min) with static intervals between cycles of 1 to 25 minutes, thus simulating dynamic changes in the pro-urine flow. To investigate the involvement of mTORC1 and pannexin-1 in FSS-modulated ATP release, cells were pre-incubated for 1hour with rapamycin (100 nM, Santa Cruz Biotechnology, Santa Cruz, CA, USA) and brilliant blue FCF (BB-FCF, 100 μM, Sigma-Aldrich, St. Louis, MO, USA), respectively.37, 38 In this case, during microfluidic experimentation, media was supplemented with the corresponding inhibitor. Control cells were exposed tomedia supplemented with the corresponding vehicle (DMSO or MilliQ-water, for rapamycin or BB-FCF, respectively). Flow-through was collected after 1 minute of FSS exposure and used for the ATP measurement (Figure 1A). Cell morphology after exposure to different FSS conditions was monitored by brightfield microscopy (Figure 1B). Figure 1Open in figure viewer Fluid shear stress increases ATP release in vitro and in vivo. A, Schematic overview of the in vitro microfluidic set-up used to measure cellular ATP release. B, Brightfield microscopy of mDCT15 cellular morphology before and after exposure to either 0.3 and 1.2 dyn/cm2 FSS. C, Elevated FSS-modulated ATP release in mDCT15 cells exposed to 1 minute of 0.3, 0.6, and 1.2 dyn/cm2 FSS (n = 4), significant difference (P < .05) denoted with symbol a. D-F, Attenuated FSS-modulated ATP release in mDCT15 cells after repeated 1 min sampling during exposure, for 5 min in total, to 0.3, 0.6, and 1.2 dyn/cm2 FSS, respectively (n = 4). G, Recovery of FSS-modulated ATP release after static intervals of 1 to 25 min between 1 min FSS exposure periods (0.3 to 1.2 dyn/cm2) (n = 4), significant differences between 0.3 and 1.2 dyn/cm2 are denoted with symbols a, b and c. H, In vivo resemblance of FSS-modulated ATP release in healthy human subjects after acute water loading (n = 7), significant differences indicating elevated urinary ATP (from t = 0 to t = 60 and t = 90, respectively) are denoted with symbol a, whereas significant differences indicating a decreased urinary ATP (from t = 90 to t = 150) are denoted with symbol b. Values are presented as the mean ± SEM ATP levels were determined in the (flow-through) medium, total cell lysates, and urine using the ATPlite Luminescence Assay System (Perkin Elmer, Waltham, MA, USA) according to manufacturer's instructions. A standard curve, ranging from 0.6 × 106 to 2.5 × 106 and 0.3 × 102 to 10.0 × 103 pmol/L ATP for intracellular and extracellular ATP, respectively, was used. All samples measured were within the confines of the standard curve used. ATP content in the medium and total cell lysates was corrected to the total volume and normalized to the total cell number in the corresponding channel or well. For in vitro experimentation, fold change of extracellular ATP for each condition to control is depicted. In vivo, ATP content in urine was normalized to urinary creatinine for human subjects and to urinary volume collected in 24 hours for mice.39 2.4 Acute water loading Urine samples of healthy human volunteers were collected, for ATP measurement, after acute water loading experimentation. These samples were obtained from a previously performed study and supplemented with protease inhibitors at a final concentration of 50 μM PMSF, 20 μM aprotinin, 10 μM pepstatin A, and 20 μM leupeptin, and stored at −80°C.40 All volunteers were healthy males between the age of 25 and 35years with a body weight lower than 100kg. Subjects were asked to refrain from coffee, tea, and alcohol intake and exercise for 24 hours before and during the study. In 30 minutes, each subject ingested 20ml/kg body weight of water. Midstream urine was collected before (t = 0) and after water ingestion in 30 minutes intervals for up to 150 minutes. Urine samples were immediately stored at −80°C. 2.5 Mouse experimentation Inducible kidney-specific Pkd1 knockout mice (iKsp-Pkd1lox/lox) were used to assess the role of PC1, in vivo, in urinary ATP excretion. Tamoxifen was administered, via oral gavage, to iKsp-Pkd1lox/lox mice on postnatal days 18, 19, and 20 (PN18) to induce a kidney specific knockout of Pkd1 (iKsp-Pkd1del).41 iKsp-Pkd1lox/lox mice which received no tamoxifen were considered as age and genotype-matched controls. At PN18 + 29 days, mice were placed in the metabolic cages for 24 hours to collect urine in a continuously cooled container (4°C) and stored at −20°C for ATP measurements. Finally, mice were killed by cervical dislocation. Precystic kidneys were extracted, weighed, and collected in liquid nitrogen and stored at −80°C for mRNA isolation. These samples were obtained from a previously performed study.41 2.6 Gene expression analysis To evaluate the effect of FSS on gene expression, cells in μ-slide VI0.4 channels were exposed during 3 hours to either no flow (=static) or 0.6 dyn/cm2 FSS (Figure 4A). After experimentation, cells were lysed and RNA was isolated using the RNeasy mini kit according to manufacturer's instructions (Qiagen, Hilden, Germany). cDNA synthesis using Molony Murine Leukemia Virus Reverse Transcriptase (Invitrogen, Carlsbad, CA, USA) and RTqPCR, using SYBR green master mix (Bio-Rad Laboratories, Hercules, CA, USA) were performed as previously described.42 The expression of the following genes was assessed: Abcc6, Cx37, Entpd2, Entpd3, Nt5e, P2rx4, P2rx5, P2rx6, P2rx7, P2ry1, P2ry2, Panx1, and Ptgs2. Gapdh was used for each gene of interest as a reference gene and non-template controls were used during RTqPCR determinations as negative controls. Since all primers used had an efficiency of approximately 100% at the concentration (400nM) used, the relative gene expression was analyzed using the Livak method (2−ΔΔCt) (Supporting Table S1). The RTqPCR procedures described here complied with the MIQE guidelines. 2.7 Fura-2-based Ca2+ imaging Intracellular Ca2+ was measured using the ratiometric dye Fura-2. Press-to-Seal silicone isolators with adhesive 24 wells (Molecular Probes, Eugene, OR, USA) were attached to a 24 × 50 mm cover glass. The wells were coated with Poly-L-Lysine (Sigma-Aldrich, St. Louis, MO, USA) and washed with PBS. The procedures were performed in a Ca-HT buffer (4.2mM KCl, 132mM NaCl, 1mM MgCl2, 1.4mM CaCl2, 5.5mM glucose, and 10mM HEPES in MilliQ). Subsequent, 15 μL of cell suspension was added to each well and incubated for 1.5 hours. About 5 μL of medium was removed and 10 μL Fura-2-AM mixture (0.3% v/v Fura-2-AM, 0.015% v/v Pluronic acid (10% v/v in Ca-HT buffer) was added and incubated for 1hr. For the conditions in which P2Y1 was inhibited, MRS2500 (Tocris Bioscience, Bristol, United Kingdom) was added to the Fura-2 AM mixture at a concentration of 1 μM. One measurement per well was performed. The well was placed into a perfusion chamber mounted onto the stage of an inverted microscope (Zeiss Axiovert 200M, Carl Zeiss, Jena, Germany). Intracellular Ca2+ was monitored by exciting Fura-2 with monochromatic light of wavelength 340 and 380nm (Polychrome IV, TILL Photonics, Gräfelfing, Germany). Fluorescence emission light was directed by a 415DCLP dichroic mirror (Omega Optical, Brattleboro, VT, USA) through a 510WB40 emission filter (Omega Optical, Brattleboro, VT, USA) onto a CoolSNAP HQ monochrome CCD camera (Roper Scientific, Vianen, The Netherlands). The integration time of the CCD camera was set at 200ms with a sampling interval of 3 seconds. After 30 seconds of measuring, either ATP (100 μM) in Ca-HT, ATP (100 μM) + MRS2500 (1 μM) in Ca-HT or just Ca-HT was added to the mini-well. At each timepoint the 340/380nm ratio was calculated per region. To determine the maximal response, the delta peak was calculated by subtracting the baseline ratio from the maximum ratio for every cell. Only the responding cells were used for data analysis. 2.8 Immunocytochemistry Primary cilia in mDCT15 cells were visualized as previously described.42 The following antibodies were used: primary antibody rabbit polyclonal anti-ARL13B (1:200, Proteintech, Rosemont, IL, USA) and secondary antibody Alexa Fluor 594-conjugated anti-rabbit IgG (1:250, Molecular Probes, Eugene, OR, USA). Nuclei were counterstained and cells mounted using DAPI-Fluoromount-G (ITK Diagnostics, Uithoorn, The Netherlands). Acquisition of images was performed using confocal laser scanning microscopy (FV1000, Olympus, Tokyo, Japan) equipped with a 60X oil-immersion objective. ARL13B-positive primary cilia (594 nm laser), identified as dots or small dashes on the xy-plane, and DAPI-positive nuclei (405 nm laser) were visualized. A z-stack with 0.25 μM distance between each focal plane was obtained. Images were processed using Fiji (https://fiji.sc/) software (National Institutes of Health, Bethesda, MD, USA). 2.9 sgRNA design Guide RNA's (sgRNA) targeting exon 9 of the Mus musculus gene Pkd1 and exon 1 of the Mus musculus gene Panx1 were designed using http://crispor.tefor.net/ (Supporting Table S1).43 The oligonucleotide pair for each sgRNA was phosphorylated and annealed using T4 Polynucleotide Kinase (New England Biolabs, Ipswich, MA, USA) in a thermocycler (37°C for 30 minutes, 95°C for 5 minutes and ramped down to 25°C with 5°C/min). The plasmids PX458 (Addgene #48138)44 and PX333 (Addgene #64073)45 were linearized using BbsI, or BsaI, restriction enzymes (New England Biolabs, Ipswich, MA, USA) at 37°C for 1hr. The annealed oligonucleotides were ligated into the linearized PX458 or PX333 plasmid using T4 Ligase (New England Biolabs, Ipswich, MA, USA) overnight at 16°C. Next, the ligation mix was transformed by 42°C heat shock into TOP10F competent cells. The next day, colonies were picked and the PX458 or PX333 plasmid, containing the sgRNA, isolated. The PX333 was modified by inserting T2A-eGFP (derived from PX458) after NLS-Cas9-NLS resulting in the plasmid PX333-GFP, suitable for GFP-positive FACS sorting. The PX333 plasmid allows for the dual expression of sgRNA's. 2.10 CRISPR/Cas9-mediated genome editing mDCT15 cells were transfected with the PX458 (Addgene #48138,44 or PX333-GFP plasmid (modified from Addgene #64073),45 containing the sgRNA's, using Lipofectamin 2000 reagent (Invitrogen, Carlsbad, CA, USA). GFP-positive cells were single cell sorted into 96-well plates by fluorescence-activated cell sorting (FACS, Aria, BD Biosciences, San Jose, CA, USA) at 48 hours post-transfection. Next, single cell clones were grown (37°C, 5% (v/v) CO2) for 7 to 14days in DMEM/F-12 1:1 nutrient mixed media (Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 5% (v/v) FBS (GE Healthcare, Little Chalfont, United Kingdom) and ciprofloxacin (10 μg/mL). Once cells reached confluency, genomic DNA was isolated using the prepGEM DNA extraction kit (ZyGEM, Hamilton, New Zealand). The targeted region of either Ift140 exon 8 or Pkd1 exon 9 was amplified using AmpliTaq GOLD (Thermo Fisher Scientific, Waltham, MA, USA) in a thermocycler (95°C for 5 min and 30 cycles of 95°C for 15 seconds, 58°C for 30 s, and 72°C for 30 s followed by a final elongation step of 72°C for 7 min). PCR products were run on a 2% (w/v) agarose gel to assess that one single amplicon was amplified for the PX458 transfected cells, as for the PX333-GFP transfected cells, multiple amplicons corresponding to wild type, hetero- or homozygous can be expected. Next, PCR products were ligated into the pGEM-T Easy vector (Promega, Madison, WI, USA) using TA cloning and transformed into TOP10F competent cells. Plasmids containing an insert were sequenced, using sanger sequencing and SnapGene software (version 4.2.4, GSL Biotech LLC, Chicago, IL, USA), to assess each clone for homozygous frame shift mutations. The Pkd1−/− mDCT15 clones were validated by western blot (Figure 5A) using 4%-15% Criterion TGX Precast Gels (Bio-Rad Laboratories, Hercules, CA, USA), PVDF membrane (Immobilon-P, Millipore Corporation, Bedford, MA, USA), and the antibodies mouse anti-PC1 (1:1000, Santa Cruz Biotechnology, Dallas, TX, USA) and peroxidase-conjugated sheep anti-mouse (1:10,000, Sigma-Aldrich, St. Louis, MO, USA). Blots were visualized using enhanced chemiluminescence (Thermo Fisher Scientific, Waltham, MA, USA). Both Pkd1−/− clones used in this study were generated independently by either use of the PX458 or PX333-GFP plasmid. The Ift140−/− mDCT15 cell line was previously generated and validated by sanger sequencing and immunocytochemistry.46 2.11 Zebrafish experimentation The effect of BB-FCF on cyst growth was studied in an established zebrafish model of PKD, where PKD is induced by the use of a translation blocking morpholino (MO) targeting the zebrafish ortholog of human PKD2 (pkd2).47 Wild-type Tupfel long-fin zebrafish were used for experimentation. In brief, PKD was induced in zebrafish larvae by microinjecting one- to two-cell stage embryos with the following pkd2-MO:5′-AGGACGAACGCGACTGGAGCTCATC-3′. This MO has been previously validated as pkd2-morphants phenocopy pkd2 mutants.48 In parallel, control embryos were injected with a standard mismatch control MO (5′-CCTCTTACCTCAGTTACAATTTATA-3′), directed against a human β-globin intron mutation, at the amount of 2 ng/embryo (Gene Tools, Philomath, OR, USA). At 4 hours postfertilization (hpf), BB-FCF was added to the medium (standard E3 medium: 5mM NaCl, 0.17mM KCl, 0.33mM CaCl2, 0.33mM MgSO4) at a final concentration of 100 μM. Non-treated fish were also included in our study. In this case, only the vehicle (MilliQ water) was added (same volume as used when adding BB-FCF) to the medium. In total, four experimental groups were realized: two groups microinjected with the pkd2-MO, exposed and non-exposed to BB-FCF; and two groups microinjected with the mismatch control MO, exposed and non-exposed to BB-FCF. At 5days postfertilization (dpf) larvae were anesthetized with tricaine/Tris pH 7.0 solution. Using brightfield microscopy (20x magnification) mortality, pronephros cystic phenotype, curved tail and pericardial edema were assessed, and high-resolution images of larvae were obtained. Images were processed using Fiji (https://fiji.sc/) software (National Institutes of Health, Bethesda, MD, USA) (Supporting Figure S1A,B). All animal procedures detailed here were performed in accordance with national and international legislations and were approved by the ethical review committee of the Radboud University Nijmegen. 2.12 Statistical analyses Differences between groups were analyzed using an unpaired Student's t test when two experimental groups were compared and a factor of variance was considered. When more than two experimental groups were considered, a one-way (one factor of variance) or a two-way ANOVA (two factors of variance) was applied, followed by the Tukey's test for multiple comparisons. Data in the figures are expressed as mean ± SEM. In the case of the experiments performed in vitro, the mean is the average value of independent experiments (n = 3-4). Each independent experiment consisted of at least two (in the case of ATP measurements) or three (in the case of gene expression measurements) replicates for each of the conditions tested. For in vivo experiments, the sample size is depicted in the figure legend. Differences in survival of zebrafish larvae were analyzed with the logrank (Mantel-Cox) test. P < .05 was considered statistically significant. All statistical analyses were performed with Prism 8 (GraphPad, La Jolla, CA, USA). 3 RESULTS 3.1 Flow modulates ATP extrusion in vitro and in vivo To assess whether FSS influences the cellular ATP release from renal cells, mDCT15 cells were exposed to physiological FSS (Figure 1A). Cell morphology and monolayer integrity were not affected by the culture conditions (Figure 1B). A significant elevated ATP release was observed in cells exposed to 1.2 dyn/cm2 FSS for 1 minute when compared to 0.3 dyn/cm2 FSS (100 ± 75 vs 481 ± 127%, P < .05) (Figure 1C). ATP release attenuated when cells were exposed up to 5 minutes to these rates (Figure 1D-F). Cells exposed to cycles of alternating 1 minute exposures to 0.3 and 1.2 dyn/cm2 followed by 1 minute rest displayed episodic peaks of ATP release when switching from 0.3 to 1.2 dyn/cm2 FSS (100 ± 31 vs 220 ± 23% and 5 ± 1 vs 19 ± 4%, respectively, P < .05). Increasing the static interval to 25 minutes resulted in higher ATP release after applying FSS compared to a static interval of 1 minute (7 ± 5 vs 69 ± 11%, P < .05) (Figure 1G). To assess whether ATP extrusion is FSS-modulated in vivo, an acute water loading test in healthy human subjects was performed. Acute water loading significantly increased the urinary output, indicative of elevated urinary flow, resulting in higher urinary ATP excretion (27 ± 12 vs 2,426 ± 530 and 27 ± 12 vs 2,951 ± 696 pmol ATP/mmol creatinine for t = 0 vs t = 60 and t = 0 vs t = 90 minutes, respectively, P < .05) (Figure 1H). Overtime, urinary output decreased, resulting in lower urinary ATP excretion (2,951 ± 696 vs 901 ± 266 pmol ATP/mmol creatinine for t = 90 vs t = 150 minutes, respectively, P < .05). 3.2 ATP extrusion not dependent on primary cilia Studies have shown that FSS-modulated ATP release is primary cilia-dependent.49 Therefore, Ift140−/− mDCT15 cells were used, where ciliogenesis is inhibited, to study the ATP extrusion upon FSS mechanosensation.46 After staining for the primary cilia-specific protein ARL13B, no primary cilia were detected in Ift140−/− cells. Wild-type cells displayed primary cilia (Figure 2A). Cells exposed to 0.3 and 1.2 dyn/cm2 FSS exhibited similar FSS-modulated ATP release between wild-type and Ift140−/− cells (100 ± 34 vs 894 ± 107 and 100 ± 44 vs 770 ± 274%, respectively, P < .05) (Figure 2B). Figure 2Open in figure viewer FSS-modulated ATP release is not dependent on the presence of primary cilia. A, Immunocytochemical observation of primary cilia, detected through ARL13B staining, in wild-type mDCT15 ce
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