Adhesion GPCR Gpr126 (Adgrg6) Expression Profiling in Zebrafish, Mouse, and Human Kidney

Abstract

Adhesion G protein-coupled receptors (aGPCRs) comprise the second-largest class of GPCRs, the most common target for approved pharmacological therapies. aGPCRs play an important role in development and disease and have recently been associated with the kidney. Several aGPCRs are expressed in the kidney and some aGPCRs are either required for kidney development or their expression level is altered in diseased kidneys. Yet, general aGPCR function and their physiological role in the kidney are poorly understood. Here, we characterize in detail Gpr126 (Adgrg6) expression based on RNAscope^® technology in zebrafish, mice, and humans during kidney development in adults. Gpr126 expression is enriched in the epithelial linage during nephrogenesis and persists in the adult kidney in parietal epithelial cells, collecting ducts, and urothelium. Single-cell RNAseq analysis shows that gpr126 expression is detected in zebrafish in a distinct ionocyte sub-population. It is co-detected selectively with slc9a3.2, slc4a4a, and trpv6, known to be involved in apical acid secretion, buffering blood or intracellular pH, and to maintain high cytoplasmic Ca²⁺ concentration, respectively. Furthermore, gpr126-expressing cells were enriched in the expression of potassium transporter kcnj1a.1 and gcm2, which regulate the expression of a calcium sensor receptor. Notably, the expression patterns of Trpv6, Kcnj1a.1, and Gpr126 in mouse kidneys are highly similar. Collectively, our approach permits a detailed insight into the spatio-temporal expression of Gpr126 and provides a basis to elucidate a possible role of Gpr126 in kidney physiology.

1. Introduction

The adhesion G protein-coupled receptor (aGPCR) family comprises 33 members in humans (31 mouse and 24 zebrafish orthologs). The hallmark of aGPCRs is their multitude of adhesion-like domains in a large N-terminus and their unique autocatalytic processing [1]. All, except for one member, contain in their extracellular portion a GAIN domain with a GPCR proteolysis site (GPS) motif. Receptor cleavage generates an N-terminal fragment (NTF) that includes most of the extracellular region and a remaining C-terminal fragment (CTF) containing a 7-transmembrane domain (7TM). Functionally, GPCRs require the interaction of a heterotrimeric G protein on the cytoplasmic side of the cell membrane to conduct their biological response. This response is usually mediated by increased intracellular production of cAMP, which initiates a signaling cascade via the activation of cAMP-dependent protein kinases. Crystallography studies of aGPCRs have described cryptic sequences between GAIN domain β-sheets that could act as tethered agonists for CTF activation. These sequences are called Stachel sequences and might be unmasked by conformational changes in the extracellular domain or by dissociation of the cleaved NTF from the CTF [2,3,4]. In addition, NTFs can be shed off and induce non-cell autonomous signals in neighboring or distant targets [2,5].

aGPCRs have been reported to be important for many pathological processes, including inflammatory conditions [6], tumorigenesis events [7], and lung as well as kidney diseases [8,9]. Notably, aGPCRs represent the second-largest class of the GPCR superfamily whereas GPCRs are currently targeted by ~34% of all approved drugs by the US Food and Drug Administration [10]. Therefore, uncovering the function of understudied GPCRs provides a wealth of untapped therapeutic potential. However, their activation mechanisms, ligands, and downstream signaling remain poorly understood. Thus, research on these receptors is required to unravel their physiological relevance and to expose their clinical potential as pharmaceutical targets.

While a plethora of descriptive data indicate that several aGPCRs are expressed in the kidney from zebrafish to humans [1,8,11], little is known about the role of aGPCRs in kidney development. Recent evidence indicates that aGPCRs can control cell polarity [12], mitotic spindle orientation [13], cell migration [12,14], cell aggregation [15], and the transduction of mechanical stimuli [16,17,18]. These are all processes that also play an important role in kidney ontogenesis and, when disturbed, lead to kidney disease [19,20,21,22,23,24]. The best-characterized aGPCR in kidney development is Celsr1 (Adgrc1) [25]. Mice carrying mutations in Adgrc1 exhibit UB branching defects at embryonic day (E) 13.5. At E17.5, Adgrc1 mutant mice were characterized by kidney growth retardation, dilated cortical tubules, and mitotic spindle misorientation. Adgrf5-knockout (KO) mice exhibited an accumulation of vacuolar-type ATPase pumps at the apical membrane of A-ICs of the collecting ducts, resulting in an excessive acidification of the urine and a metabolic alkalosis. Adgrf5/Adgrl4 double KO mice exhibit loss of endothelial fenestration and fusion of podocyte foot processes resulting in proteinuria, uremia, and death within 3 months [26,27]. Adgrg3 KO mice in the AKI model exhibited significantly less renal injury compared to wildtype (WT) mice [28]. Notably, Adgrg3 is upregulated in the kidneys from patients with biopsy-proven acute tubular necrosis compared with healthy controls.

Recently, we have observed that Gpr126 (Adgrg6) is expressed in the kidney based on a lacZ reporter mouse line [17]. In addition, a database analysis had revealed that a variety of kidney diseases are associated with altered expression of aGPCR including Gpr126 [8]. Gpr126 possesses a signal peptide, a 7TM domain homologous to secretin-like GPCRs, a GAIN domain, GPS motif, an extended N-terminus containing a Complement, Uegf, Bmp1 (CUB), a pentraxin (PTX) and a hormone-binding domain, and 27 putative N-glycosylation sites. Gpr126 is cleaved at the GPS motif in the endoplasmic reticulum. The resulting NTF and CTF locate to the plasma membrane where they are non-covalently associated with each other [29,30]. In silico analysis indicated that the cytoplasmic domain of CTF contains potential sites for palmitoylation; phosphorylation sites for cAMP dependent kinases/protein kinase G, protein kinase C, and casein kinase II; a potential myristoylation signal; and a microbodies C-terminal targeting motif [31]. KO-based phenotypes support a model in which Gpr126 elevates intracellular cAMP through interactions with Gα_s [32,33,34]. Chimeric G-protein assays further indicated that GPR126 can also couple to G_i-proteins [33]. In addition, laminin-211 [35], type IV collagen [36], type VI collagen [37], and the cellular prion protein PrP(C) [38] have been identified as potential Gpr126 ligands that activate G protein signaling. Furthermore, Gpr126 contains in its CTF a short peptide sequence (termed “Stachel” sequence) that functions as a tethered agonist [4,39]. Yet, the consequence of Gpr126-mediated G protein signaling remains poorly understood. Notably, increase in cAMP has also been detected when shaking forces were applied to Gpr126-transfected cells [35]. The interaction of Gpr126 with laminin-211 leads to a synergistic increase in intracellular cAMP levels under dynamic conditions (vibration or shaking). Moreover, Gpr126 expression was found in a variety of tissues known to be subjected to flow or mechanical forces, such as the endocardium of the heart and the hyaline cartilage of the trachea [17].

Gpr126 has been reported to be required for the development of several tissues/organs including the peripheral nervous system, intervertebral disc, ear, placenta, and heart [5,32,34,40,41,42,43]. Yet, despite its clinical importance, little is known about Gpr126-mediated cellular processes and signaling, and thus large efforts are invested in understanding the functional role of Gpr126. Here, we aimed to determine its cellular expression pattern during kidney development in order to provide a basis to elucidate a possible role of GPR126 in kidney function.

2. Materials and Methods

2.1. Human Material

The data obtained in this study were processed according to the principles of scientific practice and all patient samples were analyzed anonymously. Human embryonal kidney tissues were obtained from routine autopsies after confirming the absence of a morphologically recognizable renal phenotype. Normal kidney and bladder tissues without evidence of disease were obtained from distant portions of kidneys or bladders surgically excised because of the presence of a localized neoplasm and diagnosed at the Institute of Pathology, FAU. Kidney and bladder sections showed normal histology without signs of disease. All tissues were provided as formalin-fixed, paraffin-embedded (FFPE) sections of 4 µm thickness. The use of FFPE tissue material was approved by the Ethics Committee of the Friedrich-Alexander-University of Erlangen-Nürnberg for the archive of the Department of Nephropathology (Re.-No.4415 and Re.-No.22-150-D) and Institute of Pathology (Re.-No. 4607) waiving the need of informed consent due to its retrospective nature.

2.2. Animal Housing and Experimentations

Kidneys from wildtype C57BL/6 and lacZ-Gpr126 reporter (Adgrg6tm1a^(EUCOMM)Hmgu) mice [17] were used to determine the endogenous expression of Gpr126. Wildtype AB and Tg(chd17:GFP) zebrafish strains [44] were utilized to identify epithelium of the pronephric and mesonephric tubules and ducts. The transgenic cdh17-GFP line was kindly provided by Prof. Neil Hukriede. Housing, breeding, as well as organ perfusion and extraction conformed with the guidelines from directive 2010/63/EU of the European Parliament of the protection of animals used for scientific purposes and were approved by the local governmental animal protection committee. For experimental details, see below (Re. No. TS-05/11 Pathologie). For LacZ staining, kidneys from Adgrg6tm1a^(EUCOMM)Hmgu mice were obtained as previously described [17]. In brief, mice were anesthetized by inhalation of 3% isoflurane and subjected to transcardial perfusion with 10% dextran 40 supplemented with 0.1% procaine followed by 0.9% NaCl to clear residual blood from tissues (protocol TS-05/11 Pathologie). For RNAscope^® in situ hybridization, wildtype C57BL/6 mice were, after isoflurane sedation, transcardially perfused with 0.9% NaCl (protocol TS-05/11 Pathologie). Organs were extracted and post-fixed by immersion in 4% buffered formalin for 24 h at RT before embedding in paraffin.

2.3. Antibodies and RNA Probes

The primary antibodies used in this study were anti-cTnI (1:300; Abcam, ab56357), anti-Krt8 (1:6; Developmental Studies Hybridoma Bank, TROMA-I), anti-Claudin-1 (1:100; Abcam, Cambridge, UK, ab15098), anti-E-cadherin (1:200; BD Transduction, BD 610181), anti-Aquaporin-2 (1:100; Sigma-Aldrich, St. Louis, MO, USA, SAB5200110), and anti-V-ATPase B1 (1:100; Invitrogen, PA5-56878). The secondary antibodies were Alexa Fluor^® 488 donkey anti-rabbit (1:200; Life Technologies, Carlsbad, CA, USA, A21206) and Alexa Fluor^® 647 donkey anti-mouse (1:200; Life Technologies, A31571). The RNA probes targeting Gpr126 in zebrafish, mouse, and human tissues were designed and provided by Advanced Cell Diagnostics^® (Dr-adgrg6, 564461; Mm-Adgrg6, 472251; Hs-ADGRG6, 480121).

2.4. LacZ Staining

Organs were washed in cold phosphate-buffered saline (PBS), embedded in Tissue-Tek^® O.C.T. (Sakura, 4583), and frozen in the vapor phase of liquid nitrogen. Samples were stored at −80 °C for a minimum of 24 h and sectioned in 10 µm slices. Fixation was performed with 0.2% glutaraldehyde solution (Sigma-Aldrich, G5882) for 40 min on ice and incubated in a X-gal staining solution (AppliChem, Darmstadt, Germany, 74134) at 37 °C overnight. Sections were counterstained with nuclear fast red (Carl Roth, Karlsruhe, Germany, N069.1) for 5 min at room temperature (RT) and washed in dimethyl sulfoxide (Sigma-Aldrich, D4540) and xylol-free clearing solution (Merck Millipore, Burlington, MA, USA, 1098435000).

2.5. Immunofluorescence Staining

FFPE sections with a thickness of 4 µm were deparaffinized in xylol twice for 10 min each at RT. Then, rehydration of the tissue was performed by washing for 5 min in ethanol solutions with gradually decreased concentrations (100%, 96%, 70%, and 50%) and distilled water. To unmask epitopes possibly altered by the fixation process, sections were incubated in a retrieval solution provided by Advanced Cell Diagnostics^® at 95 °C for 15 min in a programmable pressure cooker. Sections were then subjected to the same treatment as that for RNAscope^® in situ hybridization (see below). Lastly, sections were permeabilized with 0.3% Triton^® X-100 in PBS for 10 min at RT and incubated for 1 h at RT with a blocking buffer containing 3% [w/v] bovine serum albumin (Carl Roth, 8076.4), 5% [w/v] goat serum (Biowest, Nuaillé, France, S1810), 20 mM MgCl₂, and 0.3% [v/v] Tween-20 in PBS. Primary antibodies were incubated at 4 °C overnight, and fluorophore-conjugated secondary antibodies were incubated at RT for 2 h. Sections were finally counterstained with 0.5 μg/mL DAPI (4′,6′-diamidino-2-phenylindole) in 0.1% NP40/PBS at RT for 10 min to visualize nuclei.

2.6. RNAscope^® In Situ Hybridization

Both chromogenic and fluorescent RNAscope^® in situ hybridizations were performed according to the manufacturer’s instructions. Target probes were designed and provided by Advanced Cell Diagnostics^® with the rest of the kit components (RNAscope^® 2.5 HD Detection Reagents RED, 322360; RNAscope^® Multiplex Fluorescent Detection Reagents v2, 323110). FFPE tissue sections were deparaffinized as explained for immunofluorescence staining and subjected to a provided hydrogen peroxidase solution at RT for 10 min to block endogenous peroxidase activity. To restore antigenicity after fixation, an antigen retrieval solution provided by the manufacturer was used at 95 °C for 15 min in a programmable pressure cooker. The tissue was then rinsed in deionized water and immediately treated with a provided protease solution at 40 °C for 30 min. Apart from the zebrafish probe, which had to be diluted first (1:50 in TSA^® buffer), 3–4 drops of the according species-specific target probe were added undiluted to the sections before hybridization was performed at 40 °C for 2 h. From this point, the tissue was incubated in different successive amplifier and signal development solutions depending on the chosen kit. Fluorescent signal development additionally required the incubation with Opal^® reagents (1:1500, Akoya Biosciences, Marlborough, MA, USA, FP1488001KT) in TSA^® buffer at 40 °C for 30 min. To combine in situ hybridization with immunostaining, primary antibodies were incubated overnight at 4 °C after 1 h blocking with a solution containing 0.5% [w/v] bovine serum albumin (Carl Roth, Karlsruhe, Germany, 8076.4). Secondary antibodies were then incubated for 30 min at RT. To counterstain the chromogenic assay, hematoxylin (Carl Roth, Karlsruhe, Germany, T865.2) was used at RT for 30 s and followed by running warm tap water for 5 min. To counterstain the tissue in the fluorescent assay, freshly prepared DAPI solution (0.5 μg/mL in 0.1% NP40/PBS) at RT for 10 min, DAPI solution provided in the kit for 30 s at RT, or the combination of DAPI solution (40 μg/mL in H₂O) with Mowiol mounting medium (40 g Mowiol, Calbiochem, San Diego, CA, USA, 475904 in 200 mL 0.2 M Tris-HCl, pH 8.5; 100 mL glycerin; 8.75 g DABCO, Sigma, D-2522 + 2 mL DAPI solution) was utilized.

2.7. Microscopy and Image Analysis

Bright-field images were taken using a Zeiss^® Axio Zoom V16 stereoscope or an AMG^® Evos, Keyence^® BZ-900, or Olympus^® BX60 microscope. Confocal images were taken with a Zeiss^® 800 LM laser scanning microscope. For routine rapid examination of the slides, a Zeiss^® AxioTech Vario 100HD and Olympus^® BH2 microscope or a Leica^® S6 stereoscope were utilized. Image processing and analysis was performed using Zeiss^® ZEN (Blue edition), ImageJ with the Fiji extension package, and Adobe^® Photoshop CC 2018. Imported z-stacks were projected into a single plane using maximum intensity projection. Confocal images were acquired as 8-bit at a standard resolution of 1024 × 1024 pixels. Laser intensity and signal amplification parameters were maintained between different conditions of the same experiment. To quantify the RNAscope^® signal, a semi-quantitative scoring approach was applied, as recommended by Advanced Cell Diagnostics^® guidelines. A score from 0 to 4 was given according to the general number of dots and/or dot clusters found in the tissue after RNAscope^® in situ hybridization (0: <1 dot/10 cells; 1: 1–3 dots/cell; 2: 4–9 dots/cell, none or infrequent dot clusters; 3: 10–15 dots/cell and/or <10% dots in clusters; 4: >15 dots/cell and/or >10% dots in clusters).

2.8. Single-Cell RNAseq Analysis

Single-cell RNA-sequencing data of zebrafish was retrieved from Zebrahub [45]. All cell type annotations referenced in this study are derived from the original study. Each annotated cell type of interest was normalized individually with the “normalize_total” function and scaled with the “log1p” function using Scanpy v1.9.1 [46] with default parameters. Neighbors were calculated based on all genes using the UMAP (Uniform Manifold Approximation and Projection) algorithm [47] with default settings and, subsequently, a two-dimensional plot was constructed. The connectivity matrix output by UMAP was passed to the Leiden [48] algorithm to calculate communities with a resolution of 1. Differential expression analysis was conducted with MAST [49]. Analysis of differentially expressed genes was performed with FishEnrichr [50,51].

2.9. Statistical Analysis

Statistical analysis was carried out using GraphPad^® Prism 5. Differences between groups were considered statistically significant when the p-value was ≤0.05. The statistical significance of differences between groups was tested using Mann–Whitney U test for data based on the RNAscope^® score system comparing the means of two groups of data defined by categorical variables. In case of more than two independent groups of data, one-way analysis of variance (ANOVA) was applied followed by Tukey’s honest significance test (Tukey’s HSD).

3. Results

3.1. RNAscope^® Technology Is Suitable to Detect Gpr126 Expression

After testing all commercially available anti-Gpr126 antibodies utilizing Gpr126 KO mice and Gpr126 mutant zebrafish lines, we could not identify a reliable working antibody in mice or zebrafish. In order to study Gpr126 expression at cellular resolution, RNAscope^® technology was used to perform RNA in situ hybridization on FFPE tissues. Previously, Gpr126 has been demonstrated to be expressed in the endocardium of mouse E11.5 hearts by in situ hybridization [5]. Therefore, E11.5 hearts were used as controls, which are composed of an inner single layer of endocardial cells and several layers of cardiomyocytes. RNAscope^® fluorescent signal was observed as punctuated dots lining the lumen of the heart chambers on transversal sections (Figure 1a), suggesting an endocardial location. Subsequently, the RNAscope^® in situ hybridization assay was combined with anti-cardiac troponin I (cTnI) immunofluorescence staining, which served as a specific marker of cardiomyocytes (Figure 1b). This analysis confirmed Gpr126 RNA expression in the cTnI-negative endocardial cells, as previously reported [17]. Hence, it was concluded that RNAscope^® technology is a valid method to detect the murine Gpr126 mRNA.

To evaluate the suitability of the RNAscope^® technology in the kidney, adult human and mouse kidney FFPE sections were subjected to RNAscope^® analysis (Figure 1c,d). In human FFPE renal biopsies, RNAscope^® probes targeting the bacterial 4-hydroxy-tetrahydrodipicolinate reductase (dapB) gene were chosen as a negative control. As a positive control, RNAscope^® probes targeting the human peptidyl-prolyl cis-trans isomerase B (PPIB) gene were utilized. While the bacterial RNA probes are not expected to hybridize in eukaryotic systems, PPIB is known to be conserved and expressed at low levels, serving as a control for low expressed genes [52]. The RNAscope^® fluorescent assay in human tissue showed no punctuated dotty signal when dapB-targeting probes were used (Figure 1c). However, samples exhibited areas of diffuse autofluorescence consistent with the light emission properties of the hemoglobin within erythrocytes, as previously documented [53]. When RNAscope^® probes targeting PPIB were used, a signal was observed as distinctive dotty pattern in addition to the autofluorescence of red blood cells.

To establish the RNAscope^® technology in the mouse kidney, adult kidney samples from a Gpr126-lacZ reporter mouse (Adgrg6^{tm1a(EUCOMM)Hmgu}) [17], which allows determining the Gpr126 RNA expression pattern by X-gal staining, was utilized as a control (Figure 1d). X-gal staining revealed β-galactosidase activity in tubular structures at the renal medullary papilla. This expression pattern was also obtained when utilizing RNAscope^® probes for mouse Gpr126 RNA.

Taken together, these data indicate that the RNAscope^® technology is a reliable method to detect human and mouse Gpr126 RNA on FFPE sections and is compatible with immunohistochemistry. Notably, erythrocytes are identified as diffuse, larger autofluorescent areas that can be distinguished from the RNAscope^® signal, which appears as punctuated dots.

3.2. Gpr126 Expression Changes in Early Development from Ubiquitous to Nephron-Enriched

To evaluate the Gpr126 expression pattern during renal development, embryonic tissues from human and mouse metanephric kidneys were analyzed. Human embryos at 11, 20, and 28 weeks of gestation (wg) and mouse embryonic kidneys from E13.0 and E15.5 wildtype animals were utilized. Notably, metanephroi start to form at around 10 wg in humans or E10.5 in mice with the induction of the ureteric bud [54]. Later in development, the nephrogenesis rate increases exponentially from 18 to 23 wg in humans and from E13.0 to birth in mice [55]. RNAscope^® analysis revealed that Gpr126 expression levels were dependent on the developmental stage in both human and mouse samples (Figure 2).

In humans (Figure 2a,c), dot density analysis revealed that GPR126 expression at 11 wg was moderate in both stromal (RNAscope^® score: 2.4 ± 0.49) and epithelial tubular-forming cells (score 3.0 ± 0.63) (n = 4). At 20 wg, GPR126 expression was increased in both stromal (score 2.6 ± 0.66) and epithelial cells (score 3.8 ± 0.40) (n = 5). Finally, at 28 wg, GPR126 expression was markedly decreased in stromal cells (score 0.4 ± 0.49) and epithelial cells (score 2.6 ± 0.49) (n = 3). Notably, GPR126 expression was not significantly different in epithelial cells comparing 28 wg to 11 wg, whereas stromal GPR126 expression was significantly decreased (p < 0.01). To assess if this spatio-temporal expression pattern is conserved in mice, Gpr126 RNA in situ hybridization was combined with cytokeratin-8 (Krt8) immunostaining (Figure 2b), which serves as a marker of the ureteric bud-derived epithelium [56]. Similar to the human system, Gpr126 expression in mice increased in epithelial cells during kidney development from E13.0 to E15.5 (E13.0: score 1.4 ± 0.49; E15.5: score 3.2 ± 0.40, p < 0.05) but remained low in stromal cells (E13.0: score 1.0 ± 0.63; E15.5: score 0.8 ± 0.40, not significant) (n = 3).

Collectively, these data indicate that both stromal and epithelial cells express Gpr126 during human and mouse metanephros development. Yet, Gpr126 expression is enriched over time in the epithelial cells during active nephrogenesis stages.

3.3. Gpr126 Is Enriched in Parietal Epithelial Cells and the Urinary Collecting System in Mice and Humans

Single-cell RNAseq data by Ransick et al. from adult murine renal tissue indicate that Gpr126 might be expressed in different nephron segments such as the parietal epithelial cells and the collecting duct [57]. To validate the parenchymal expression pattern of Gpr126, we performed RNAscope^® in situ hybridization on FFPE tissue of wildtype C57BL/6 mice. To be able to identify the cell types showing Gpr126 signal, the in situ hybridization was combined with immunofluorescent antibody staining for different cell-specific markers. In contrast to most of the tubular structures in the renal cortex, the Gpr126 signal could be detected in the renal corpuscle colocalizing with the tight junction protein claudin 1 that is specifically expressed in the parietal epithelial cells of the Bowman’s capsule [58] (Figure 3, left panels).

Apart from the parietal epithelial cells, Gpr126 mRNA could be detected in the collecting ducts, which were labelled by co-staining of E-cadherin and the collecting duct-specific water channel protein aquaporin 2 [59]. Distal tubules, also positive for E-cadherin [60,61], and proximal tubules, characterized by a lower epithelial density of nuclei compared to distal tubules and collecting ducts and almost absent E-cadherin staining [60,61], both showed only very low levels of Gpr126 expression. Interestingly, the density of signal dots varied throughout the different segments of the collecting duct. Gpr126 expression first decreased from the cortical collecting duct (CCD) to the outer medullary collecting duct (OMCD) before increasing again in the inner medullary collecting duct (IMCD) and the papillary collecting duct (PCD). Taken together, our findings are consistent with the RNA sequencing data from Ransick et al., with renal parenchymal expression being most abundant in parietal epithelial cells and the collecting duct.

In order to characterize the expression of human GPR126 in adulthood, renal-specific antibodies were explored in combination with RNA in situ hybridization. GPR126 expression was detected in healthy adult human kidneys in both glomerular and tubular regions of the nephron, as assessed by RNAscope^® analysis (Figure 4). In the glomeruli, GPR126 RNA expression colocalized with the parietal epithelial cell marker claudin 1. In the renal tubule, GPR126 expression was observed at low-to-no signal along the proximal and distal tubule segments (Figure 4). However, it was consistently evident in both V-ATPase-positive and V-ATPase-negative tubular cells of the collecting duct, suggesting that GPR126 is expressed in both intercalated and principal cells.

The study of Gpr126 expression was extended beyond the renal parenchyma. For this purpose, human FFPE sections from tumor-free areas of different urinary tract tissues and mouse sections from the renal pelvis were subjected to chromogenic and/or fluorescent RNAscope^® analysis (Figure 5). Human GPR126 expression was detected in the inner surface of the renal pelvis, ureter, and bladder (Figure 5a), suggesting a urothelial expression. Fluorescent RNAscope^® analysis on human and mouse renal pelvises revealed that Gpr126 was predominantly detected at the urothelium in both species, while low-to-no signal was found in the underlying lamina propria (Figure 5b).

Collectively, these data suggest that Gpr126 is expressed throughout the urothelium of the urinary collecting system in humans and mice.

3.4. gpr126 Is Expressed in Ionocytes as well as Pro- and Mesonephros in the Zebrafish

Despite the lack of metanephroi, pronephric and mesonephric nephrons in zebrafish display segment-specific morphological and physiological features and spatio-temporal gene expression that are homologous to the human kidney [62]. gpr126 expression analysis with RNAscope^® combined with anti-GFP immunostaining in transversal FFPE sections from Tg(cdh17:GFP) larvae indicates that gpr126 is expressed in the pronephros as well as ionocytes (Figure 6a), which exhibit analogous functions to collecting duct epithelial cells in the embryonic skin and adult gills of zebrafish [63,64]. The analysis of recently published single-cell RNAseq data describing zebrafish development [45] confirmed that gpr126 is highly expressed in ionocyte progenitor cells at 14 h and in ionocytes at 5 and 10 days post fertilization (hpf or dpf) (Figure 6b,c and Table S1). As expected, high gpr126 expression was, for example, also detected at 10 dpf in the otic vesicle [32], cartilage [40], peripheral nervous system [34], and renal glomerulus (based on our mouse data above) (Table S1) [1].

Collecting duct cells and ionocytes plays an important role in ion homeostasis. Notably, the ion-transport-related genes nkcc1 and kcnq1 were reported to be downregulated in the otic vesicles, a main location of ionocytes, in gpr126 zebrafish mutants [32]. Previously, four types of ionocytes have been described in zebrafish (H⁺-ATPase-rich: apical H⁺-ATPase (HA, atp6v1aa), NHE3b (slc9a3.2), Rhcg1 (rhcgb), rhbg, AE1b (slc4a1b), atp1a1a.5, ca15a, ca2 (cytosolic CA2-like a), ceacam1, gcm2; Na⁺-K⁺-ATPase-rich: ECaC (trpv6), PMCA2 (atp2b2), NCX1b (slc8a1b), Na⁺-K⁺-ATPase (atp1a1a.1); Na⁺-Cl⁻ co-transporter-expressing: slc12a10.2, atp1a1a.2, NBCe1b (slc4a4b); K⁺-secreting: atp1a1a.4, Kir1.1 (kcnj1a.1)) [65,66,67,68]. ndrg1a has been described as a general marker of ionocytes and the pronephric duct [69]. Therefore, we analyzed the single-cell RNAseq data in regard to these genes to determine in which ionocyte sub-types gpr126 expression can be detected (Figure 6d).

At 10 dpf, four distinct clusters of ionocytes could be derived from the single-cell RNAseq data (1, 2, 3, 4) (Figure 6b,c). Cluster 1 contains ionocytes in which exclusively slc4a1b, ca15a, rhcgb, and rhbg expression could be detected. In addition, they are characterized by enriched expression of H⁺-ATPase (atp6v1aa) and ca2. Thus, cluster 1 ionocytes represent H⁺-ATPase-rich ionocytes. Notably, several genes known to be expressed by H⁺-ATPase-rich ionocytes are co-expressed in clusters 1 and 2 (ceacam1, atp1a1a.5, and gcm2), while slc9a3.2 expression could exclusively be detected in cluster 2 (Figure S1a). Expression of genes previously identified as marker genes for Na⁺-K⁺-ATPase-rich ionocytes are selectively detectable in ionocytes at 5 dpf, represented by clusters 5 and 6 (Figure S1b). Yet in cluster 5, there are also 10 dpf cells included (Figure 6b). Genes previously identified as marker genes for Na⁺-Cl⁻ co-transporter-expressing ionocytes are exclusively detected in cluster 3 or in both cluster 3 and 4 (Figure S1c). In contrast to these 3 ionocyte-types, K⁺-secreting ionocytes could not, based on atp1a1a.4 and kcnj1a.1 expression, be identified as a distinct cell population/cluster (Figure S1d).

In order to predict a possible function of gpr126, we determined in which clusters gpr126 expression can be detected. Notably, gpr126 expression was selectively detected at 10 dpf in cluster 2 together with slc9a3.2, slc4a4a, and trpv6 expression. In addition, cells of cluster 2 were enriched in kcnj1a.1 expression and co-expressed a subset of the H⁺-ATPase-rich ionocyte marker genes (ceacam1, atp1a1a.5, and gcm2). Taken together, these data indicate that Gpr126 is expressed in a subset of H⁺-ATPase-rich and possibly Na⁺-K⁺-ATPase-rich ionocytes.

To determine whether other aGPCRs exhibit a similar expression pattern as gpr126 in ionocytes, we analyzed all genes starting with “adgr” (Table S2). This analysis revealed that only a few aGPCRs are enriched in a specific cluster (Table S2 and Figure S2a). adgrl3 was the only aGPCR found that exhibits a similar pattern as gpr126 (Figure S2a). In addition, we performed a differential expression analysis, where we compared cells in which gpr126 was detected versus cells that lack gpr126 UMIs (Table S3). Among the differentially expressed genes, we detected three genes that have previously been associated to Gpr126 (Figure S2b). It has been described that GPR126 regulates angiogenesis through modulation of VEGFR2 receptor signaling [70]. Further, GPR126 expression in periodontal ligament cells increased expression of bone sialoprotein, osteopontin, and Runx2 among others through inhibitor of DNA binding 4 (ID4) [71]. Moreover, TMED and GPR126 are both associated with height in humans [72]. While vegfab was, like gpr126, enriched at 10 dpf in cluster 2, id4 and tmed10 were more broadly detected (Figure S2a). The analysis of all differentially expressed genes with FishEnrichr [50,51] indicated that these genes are, for example, associated with the biological processes “sodium ion homeostasis”, “calcium ion homeostasis”, “response to osmotic stress”, and “response to ethanol” (Figure S2c and Table S4).

The analysis of gpr126 expression in trunk sections of 3-months-old adult wildtype zebrafish revealed gpr126 expression in both stromal and tubular mesonephric cells (Figure 6e), resembling the Gpr126 expression pattern seen in mammalian embryonic kidneys (Figure 2).

4. Discussion

Our work indicates that Gpr126 is involved in kidney development. Gpr126 is detected in the pronephric and mesonephric kidneys of zebrafish, as well as in the metanephric kidneys of mice and humans. While Gpr126 is widely expressed during development, its expression enriches in adulthood in parietal epithelial cells and epithelial cells of the collecting duct and the urothelium.

Most human aGPCR members and several orthologs have been detected in the kidney from zebrafish to human [8]. However, most of these data are based on RNA or “omics” approaches and only one member, Adgrc1, has been characterized in detail during renal development. Thus, it is crucial to explore strategies and methodologies aimed at providing a more detailed spatio-temporal analysis at cellular resolution. In this work, the RNAscope^® technology has been established to detect Gpr126 at cellular resolution in the kidneys of zebrafish, mice, and humans during development and adulthood. The presented data show that RNAscope^® technology is suitable to detect Gpr126 RNA in kidney FFPE sections and offers the possibility of combining in situ hybridization with immunohistochemistry on the same slide. Hence, RNAscope^® probes targeting Gpr126 in combination with several kidney-specific antibodies served to conclude that Gpr126 expression switches during metanephros development from ubiquitous to a tubule-enriched pattern in mice and humans, and remains low but consistently expressed in adult stages in the parietal epithelial cells and collecting ducts. In contrast, tubular epithelial collecting duct cells in the papillary region and the urothelial cells at the renal pelvis, ureter, and bladder show high Gpr126 expression. Importantly, it has to be considered that the RNAscope^® technology detects only RNA levels and does not prove that the RNA will be translated into proteins or provides information such as subcellular localization of the protein, which would allow us to speculate on the function of the protein. Thus, it is important to continue to establish working anti-Gpr126 antibodies.

In the mouse and human embryonic kidneys, the data presented here indicate that Gpr126 is initially ubiquitously expressed in both mesenchymal and epithelial cell types, but ends up being enriched in the adult papillary collecting ducts and parietal epithelial cells. This suggests that Gpr126 might be involved in differentiation of common early progenitor cells. There are other genes that show this behavior during kidney development, such as PDGFRB and PECAM1, glomerular markers of mesangial and endothelial cells, respectively. During human early kidney development, PDGFRB and PECAM1 expressions are found in the renal mesenchyme but become enriched inside the glomeruli by 10 wg [73]. This might also explain why Gpr126 is expressed in such different cell types, such as parietal epithelial cells and epithelial cells of the collecting duct and the urothelium, which to our knowledge exhibit no common function.

In the zebrafish, gpr126 was detected in the pronephros and mesonephros. Similar to mammalian metanephros, gpr126 expression was predominantly observed in tubular cells. However, in contrast to the mammalian kidney, some interstitial cells expressed very high levels of gpr126. Finally, gpr126 was also found in the skin of zebrafish larvae, which is known to be a tissue rich in ionocytes. Ionocytes are found in the embryonic skin and adult gills of zebrafish. They express a variety of apical and basolateral cell surface transporters that control transepithelial movement of protons, sodium, potassium, ammonium, calcium, chloride, and bicarbonate [63,64]. Thus, ionocytes present analogous functions to collecting duct epithelial cells and their study might serve to investigate a possible role of Gpr126 in ion homeostasis. In zebrafish, we found that gpr126 is expressed in a sub-cluster of H⁺-ATPase-rich ionocytes. The epithelial transport functions of these cells are related to H⁺ secretion, Na⁺, and HCO₃⁻ uptake, as well as ammonium excretion and are thus analogous to proximal tubular cells and type A-intercalated cells of the kidney in terms of transporter expression and function [65]. Ammonium secretion is mediated by Rhcg1 (rhcgb) and Rhbg, which are both not detected in gpr126-expressing cells [65,74]. Apical acid secretion is largely mediated by H⁺-ATPase atp6v1aa and the sodium/hydrogen exchanger (NHE3b/slc9a3.2) [65,75]; expressions of both were detected in gpr126-expressing cells, whereby our data show that slc9a3.2 expression is selectively detected in gpr126-expressing cells in cluster 2. Thus, gpr126 is highly associated with apical acid secretion and might play a role in this process. Notably, gpr126 was also detected in type A-intercalated cells of the mouse kidney. Furthermore, the selective detection of the renal electrogenic Na⁺/HCO₃⁻ cotransporter NBCe1-A/slc4a4a in gpr126-expressing cells indicates that gpr126 might play also a role in HCO₃⁻ uptake, contributing to the buffering of blood pH or intracellular pH [76]. Notably, ECaC (trpv6) expression was also selectively detected in gpr126-expressing cells. ECaC (trpv6) enables ionocytes to maintain high cytoplasmic Ca²⁺ concentration. trpv6 encodes a constitutively open Ca²⁺ channel that functions as the first and rate-limiting step in the transcellular Ca²⁺ transport pathway [77]. In mouse kidney, Trpv6 has been described as being expressed at the apical domain of the distal convoluted tubules (DCT2), connecting tubules (CNT), and cortical and medullary collecting ducts (CD), whereby it co-localized with AQP2 in the CCD, OMCD, and IMCD [78] and thus shows a highly similar expression pattern to Gpr126 in mice. These data suggest that gpr126 might also play a role in transcellular Ca²⁺ transport. This is supported by the co-detection of the cell-fate-related transcription factor gcm2, which regulates expression of the calcium sensor receptor gene casr [79]. The function of the novel marker of mature H⁺-ATPase-rich ionocytes, ceacam1, is currently unknown. Finally, gpr126 was co-detected with the Kir1.1 (kcnj1a.1) potassium transporter, also known as renal outer medullary K⁺ (ROMK). Its activity has been detected in the apical membrane of the thick ascending limb (TAL), DCT2, CNT, and CCD. Notably, gpr126 was not detected in mice in TAL. However, while kcnj1a.1 is important in the TAL for K⁺ recycling, which is essential for maintaining NaCl reabsorption via type II Na⁺-K⁺-2Cl-cotransporters, kcnj1a.1 plays an important role in mediating K⁺ excretion in the DCT2, CNT, and CCD [80,81]. Collectively, the zebrafish ionocyte data fit with the observed Gpr126 kidney data in mice.

The analysis of co-detection of genes together with gpr126 suggests that Gpr126 plays a role in the different functions described above. On the other hand, Gpr126 might be required for the maturation of ionocyte progenitor cells into a distinct sub-population of ionocytes. Future experiments modulating Gpr126 expression followed by functional analysis will reveal the role of Gpr126 in heart development and physiology.

Notably, the collecting ducts consist of principal and intercalated cells (similar to H⁺-ATPase-rich ionocytes), but the colocalization analyses revealed that Gpr126 is not restricted to either cell type. Gpr126 is highly detected throughout the collecting duct epithelial cells of the medullary papilla and in the urothelial cells of the renal pelvis, ureter, and bladder. Notably, inner medullary and papillary collecting ducts are continuously subject to osmotic and hydrostatic stress. They collect the hyperosmotic filtrate from multiple nephrons and withstand a fluid shear stress that oscillates between 0.2 and 20 dyn/cm², which is equivalent to the physiological range of blood shear stress in resistance vessels [82]. The urothelial cells of the renal pelvis, ureter, bladder, and proximal urethra receive the final urine and are also reported to respond actively to mechanochemical changes [83] in many cases through GPCRs [84,85]. Gpr126 has been detected in a variety of tissues and cell types, including vascular endothelial cells, subject to mechanical stress [17] and has also been described as a mechanosensor that responds to signals in the extracellular matrix by binding with type VI collagen [36] and laminin-211 [35]. This supports the idea that Gpr126 could act as a mechanosensitive gene in the epithelium of the urinary collecting system: collecting ducts, renal pelvis, ureters, bladder, and urethra, and possibly also parietal epithelial cells.

5. Conclusions

The results shown in this work demonstrate that Gpr126 is expressed in the kidney during development and adulthood, while its expression pattern is evolutionarily conserved from zebrafish to human. We provide a detailed expression analysis of Gpr126 and identified several associations to possible functions of Gpr126 in the kidney as a basis to elucidate a possible role of Gpr126 in kidney development and/or physiology. Notably, database analyses associated increased Gpr126 expression also with kidney disease [8]. Thus, in the future, it will be important to characterize how Gpr126 expression is altered in kidney disease and to determine if Gpr126 deletion affects kidney development and/or physiology.