Anatomical Evidence for Parasympathetic Innervation of the Renal Vasculature and Pelvis

Significance Statement

The kidneys critically contribute to body homeostasis under the control of the autonomic nervous system. Although the cholinergic effects on renal function have been widely confirmed, there is still lack of evidence for the cholinergic innervation of the kidney. Using a genetically modified mouse model and immunostaining, the authors found evidence suggesting that cholinergic nerves supply the main renal artery, segmental renal artery, and renal pelvis. They also found expression of acetylcholine receptors in the renal artery and its segmental branches. This anatomical evidence for parasympathetic innervation of the kidney may suggest new avenues for investigation into interventional strategies for refractory hypertension, renal innervation mechanisms, and other neuroendocrine diseases associated with renal dysfunction.

Visual Abstract

Abstract

Background

The kidneys critically contribute to body homeostasis under the control of the autonomic nerves, which enter the kidney along the renal vasculature. Although the renal sympathetic and sensory nerves have long been confirmed, no significant anatomic evidence exists for renal parasympathetic innervation.

Methods

We identified cholinergic nerve varicosities associated with the renal vasculature and pelvis using various anatomic research methods, including a genetically modified mouse model and immunostaining. Single-cell RNA sequencing (scRNA-Seq) was used to analyze the expression of AChRs in the renal artery and its segmental branches. To assess the origins of parasympathetic projecting nerves of the kidney, we performed retrograde tracing using recombinant adeno-associated virus (AAV) and pseudorabies virus (PRV), followed by imaging of whole brains, spinal cords, and ganglia.

Results

We found that cholinergic axons supply the main renal artery, segmental renal artery, and renal pelvis. On the renal artery, the newly discovered cholinergic nerve fibers are separated not only from the sympathetic nerves but also from the sensory nerves. We also found cholinergic ganglion cells within the renal nerve plexus. Moreover, the scRNA-Seq analysis suggested that acetylcholine receptors (AChRs) are expressed in the renal artery and its segmental branches. In addition, retrograde tracing suggested vagus afferents conduct the renal sensory pathway to the nucleus of the solitary tract (NTS), and vagus efferents project to the kidney.

Conclusions

Cholinergic nerves supply renal vasculature and renal pelvis, and a vagal brain-kidney axis is involved in renal innervation.

In mammals, most visceral organs are innervated by the autonomic nervous system and receive both sympathetic and parasympathetic nerves.^1,2 For the kidneys, abundant renal nerve strands were illustrated in Willis’s literature as early as 1664.³ These strands contain adrenergic nerve fibers that act as a sympathetic postganglionic pathway.^4,5 Whether the kidney has parasympathetic innervation has been controversial.^5–8

In 1950, Mitchell first proposed the hypothesis that renal nerve bundles contain parasympathetic nerve fibers,⁹ and numerous efforts have been devoted to this issue since then. A typical characteristic of parasympathetic neurotransmission is that the postganglionic nerves are cholinergic.^1,10 Accordingly, the first basis for this hypothesis is to determine the cholinergic nerve supply of the kidney.

In the 1960s, it was widely believed that the kidneys received the cholinergic nerve supply, and the histochemical demonstration of acetylcholinesterase (AChE) in the renal nerves supported that view.^11,12 In the 1970s and 1980s, ultrastructural studies using electron microscopy suggested that the adrenergic nerve terminals in the kidneys can contain AChE,^13,14 meaning that this histochemical evidence lost its persuasiveness. In 2016, van Amsterdam et al. reported the presence of parasympathetic nerves near the renal artery.¹⁵ These nerves were identified using immunohistochemistry staining for nitric oxide synthase (NOS). However, others subsequently pointed out that the NOS was not a specific and selective marker for parasympathetic nerves, and a more reliable marker should be employed.⁷

On the other hand, clinical trials have revealed new understandings. The exploration of catheter-based renal sympathetic denervation in the treatment of hypertension (HTN) has attracted increasing attention, while the results are diverse.^16–19 In early clinical trials of Symplicity Catheter System-based renal denervation (RDN; Symplicity HTN-1 trial and Symplicity HTN-2 trial), treatments resulted in a substantial reduction of BP in patients,^16,17 whereas the Symplicity HTN-3 trial did not show a significant reduction of BP.¹⁸ Murai’s group and Elvan’s group even reported that the cardiac slowdown and BP drop could be caused by catheter-based renal nerve electrical stimulation in the human renal artery.^20,21 The role of autonomic innervation of the kidney in the origin, progression, and outcome of HTN is uncertain.^22,23 There is an appeal to reexamine the exact mode of renal innervation, including that of the renal vasculature, and it is highly desirable to explore the anatomic evidence for parasympathetic innervation of the renal artery.

In this study, we genetically and immunologically labeled the cholinergic plexus along the renal artery and its segmental branches. Moreover, we identified cholinergic ganglion cells within the renal nerve plexus. In the kidney, we found cholinergic nerve varicosities associated with segmental renal arteries and renal pelvis. In addition, single-cell RNA sequencing (scRNA-Seq) suggested that acetylcholine receptors (AChRs) are expressed in the renal artery and its segmental branches. Furthermore, retrograde tracing suggested that vagus afferents conduct the renal sensory pathway to the nucleus of the solitary tract (NTS), and vagus efferents project into the kidney. The results of this study showed that cholinergic nerves supply the main renal artery, segmental renal artery, and renal pelvis, and suggested that a vagal brain-kidney axis is involved in renal innervation.

Materials and Methods

Key study purposes, techniques, and the selection of animal strains can be found in Supplemental Table 1.

Animals

Transgenic mice with specific expression of the Cre recombinase in Chat expressing cells (Chat-ires-Cre mice²⁴; Jackson Laboratory, Bar Harbor, ME; No. 006410) and Cre reporter mice with tdTomato fluorescence (Ai14 mice²⁵; Jackson Laboratory; No. 007914) were used in this study. These transgenic mouse lines were backcrossed and maintained on a C57BL/6 background. Chat-ires-Cre:Ai14 mice were generated by crossing Chat-ires-Cre mice and Ai14 mice. In this mouse strain, the long-range axonal projections of cholinergic neurons are expected to be marked with robust red fluorescence.^25,26 C57BL/6J mice were purchased from Charles River Laboratories (Beijing, PR China).

Animals were maintained in a specific pathogen-free environment and supplied with rodent chow and clean water ad libitum under a 12-hour/12-hour light/dark cycle. The room temperature was controlled at 22°C–25°C. Mice aged 4 weeks (immature), 10–13 weeks (adult), and 80 weeks (senior) were used in this study. Unless specifically stated, animals were randomized with respect to animal sex and age (10–13 weeks old). The experimental procedures were in accordance with Chinese law and were approved by the Scientific Committee of Huazhong University of Science and Technology, PR China.

RDN

C57BL/6J mice were anesthetized with 2% isoflurane followed by 1.5% isoflurane with air (800 ml of air per minute) to maintain anesthesia. Oculentum was used to prevent the dehydration of the eyes. After shaving and sanitizing, mice were covered with sterile gauze, and body temperature was maintained near 37°C using a heating pad. A midline incision was made in the abdomen, and the left kidney was exposed. The left renal artery was isolated from the renal vein by blunt dissection and entangled with a thread of cotton soaked with 10% phenol in absolute ethanol.²⁷ After 10 minutes, the thread was removed, and the denervation site was washed three times with normal saline. The abdominal wall was sealed with a sterile suture, and lidocaine hydrochloride gel was placed over the wound. For the sham group, the surgical operation was identical to the RDN group except that normal saline was used instead of phenol ethanol solution. Seven days later, the operated mice were used for further studies.

Subdiaphragmatic Vagotomy

C57BL/6J mice were anesthetized as above, and oculentum was used to prevent dehydration of the eyes. After shaving and sanitizing, the mice were covered with sterile gauze, and body temperature was maintained near 37°C using a heating pad. A midline incision was made in the abdomen, and the gastroesophageal junction was exposed. The ventral and dorsal branches of the vagus nerve were visualized along the esophagus below the diaphragm by a surgical microscope and were transected using fine forceps. For the sham group, ventral and dorsal branches of the vagus nerve were similarly exposed but not transected. Cholera toxin subunit B (CTB) or pseudorabies virus (PRV) tracing followed immediately after subdiaphragmatic vagotomy (sdVx).

CTB Tracing

A total of 1 μl Alexa Fluor 555-conjugated CTB (CTB-555; Thermo Fisher Scientific, Waltham, MA; C34776) was loaded into a pulled glass pipette and held in a micromanipulator. With the aid of a surgical microscope, CTB was slowly injected into the ventral wall of the glandular stomach (0.5 μl per injection at two injection sites). To prevent leakage of the CTB solution, the pipette was held in place for at least 2 minutes following each injection. Once the pipette was removed, the surface of the stomach was immediately cleaned using sterile cotton swabs. Following the procedure, the abdominal wall was sealed with a sterile suture, and lidocaine hydrochloride gel was placed over the wound. Three days later, the brains and stomachs were harvested.

Retrograde Adeno-Associated Virus Tracing

Ai14 mice were anesthetized using isoflurane, with induction at 2% and maintenance at 1.5%–2%. After shaving and sanitizing, the left kidney was exposed as described above. A total of 1 μl AAVretro-hSyn-Cre (5.96 × 10¹² μg/ml; BrainVTA, Wuhan, China) was loaded into a pulled glass pipette and held in a micromanipulator. With the aid of a surgical microscope, virus was slowly injected into the left kidney (0.2 μl per injection at five injection sites, approximately 1.5 mm deep). To prevent leakage of the virus suspension, the pipette was held in place for at least 2 minutes following each injection. Once the pipette was removed, the surface of the kidney was immediately cleaned using sterile cotton swabs. Then, the abdominal wall was sealed with a sterile suture, and lidocaine hydrochloride gel was placed over the wound. After 3 weeks, the brains and nodose/jugular complexes were harvested.

Retrograde Trans-Synaptic PRV Tracing

PRV injection procedure and the housing of infected mice were done in a biosafety level 2 laboratory. We injected 1 μl PRV-carrying green fluorescent protein gene (PRV-CMV-GFP; 2.0 × 10⁹ pfu/ml; BrainVTA; stored at –80°C) into the left kidney of the mice. The PRV injection procedure was identical to the retrograde AAV tracing described above. The brains and spinal cords were harvested 2.5 days after injection.

Nodose/Jugular Complex Dissections

Mice were deeply anesthetized via intraperitoneal injection of 10% urethane and then transcardially perfused with PBS (0.01 M) followed by 4% paraformaldehyde. To locate the nodose ganglion, the cervical vagus nerves were identified and traced cranially with the aid of a surgical microscope, and the nodose/jugular complex was found at the base of the skull. Then, the nodose/jugular complex was resected using micro scissors.

Dissection and Imaging of Spinal Cord

PRV-infected Chat-ires-Cre:Ai14 mice were deeply anesthetized and transcardially perfused with PBS followed by paraformaldehyde as above. The spines were resected and post fixed with 4% paraformaldehyde for 48 hours at 4°C. Then, the spinal cords were stripped from vertebrae, and dorsal root ganglia (DRG) were carefully retained to provide landmarks for segmental recognition. According to the location of DRG, thoracic, lumbar, and sacral segments were cut out. After embedding in 5% agarose, segments were sectioned with a vibrating microtome (VT1200; Leica, Nussloch, Germany) at a thickness of 50 μm. Fluorescence images were then obtained with a confocal laser scanning microscope. Identification of specific spinal cord segments was performed by comparing with a mouse spinal cord atlas,²⁸ and the tdTomato labeling of cholinergic neurons provided a critical reference.

Whole-Brain Three-Dimensional Imaging

The procedure of whole-brain imaging was described in our previous study.²⁹ Briefly, PRV-infected C57BL/6J mice were deeply anesthetized and transcardially perfused with PBS followed by 4% paraformaldehyde. The intact brains were dissected from the skull and then post fixed with 4% paraformaldehyde overnight at 4°C. After embedding in 5% agarose, brain samples were placed in a high-throughput light sheet tomography platform equipped with two orthogonal 10× water-immersion objectives (numerical aperture: 0.3; Olympus, Tokyo, Japan). After scanning (excitation wavelength: 488 nm) and sectioning (thickness: 40 μm) layer by layer, a three-dimensional image dataset of the whole brain was acquired.

scRNA-Seq

We collected renal arteries and segmental arteries from 22 male C57BL/6J mice (10 weeks old) to prepare single-cell suspensions. Briefly, mice were deeply anesthetized via intraperitoneal injection of 10% urethane and transcardially perfused with chilled PBS. With the aid of a surgical microscope, the main renal arteries and segmental renal arteries were dissected using fine forceps and micro scissors. The renal parenchyma attached to segmental renal arteries was removed as much as possible. After washing once with chilled PBS, the arteries were cut into pieces of approximately 1 mm³ and incubated with cell dissociation solution at 37°C for 30 minutes. Then, the cell suspension was passed through a 40-μm strainer and washed once with PBS. An automatic cell counter (Countstar BioMed; ALIT Life Sciences, Shanghai, PR China) was used to measure cell viability in the suspensions (84.5% viability, 494 living cells/μl).

Gel beads in emulsion (GEMs) containing single cell, 10× primers, and master mix were generated by using a Chromium Next GEM Single Cell 3′ Kit v3.1 (10× Genomics, Pleasanton, CA; 1000268) and a Chromium Next GEM Chip G Single Cell Kit (10× Genomics; 1000120), following the manufacturer’s instructions. Reverse transcription in individual GEMs was performed on a thermal cycler (S1000; Bio-Rad, Hercules, CA), and the running program was set to 53°C for 45 minutes, 85°C for 5 minutes, and held at 4°C. Next, cDNA was amplified using 12 cycles of PCR and quantified using a bioanalyzer system (4200 TapeStation; Agilent, Beijing, PR China). The library was sequenced using a sequencing system (Novaseq 6000; Illumina, San Diego, CA). Finally, approximately 31,200 reads per cell were achieved (reading strategy was performed by CapitalBio Technology, Beijing, PR China).

Histochemistry

Alexa Fluor 488–conjugated wheat germ agglutinin (WGA) working solution (0.002%; Thermo Fisher Scientific; W11261) and Hoechst 33258 working solution (0.001%; Sigma–Aldrich, St. Louis, MO; B1155) were prepared in the PBS. Primary antibody working solution was prepared in the PBS containing 3% BSA and 0.2% Triton X-100, and secondary antibody working solution was prepared in the PBS containing 3% BSA. Details of each antibody can be found in Supplemental Table 2.

For renal artery immunostaining, mice were deeply anesthetized with 10% urethane and then transcardially perfused with PBS followed by 4% paraformaldehyde. The arteries were resected with the aid of a surgical microscope and post fixed with 4% paraformaldehyde overnight at 4°C. After washing three times in PBS, the arteries were blocked for 2 hours at room temperature in PBS containing 5% BSA and 0.2% Triton X-100. Arteries were then incubated with primary antibodies against choline acetyltransferase (ChAT; goat, 1:50, Millipore, Billerica, MA; AB144P), vesicular acetylcholine transporter (VAChT; rabbit, 1:4000, Sigma–Aldrich; SAB4200559; goat, 1:500, Millipore, ABN100), calcitonin gene-related peptide (CGRP; rabbit, 1:400, Sigma–Aldrich, C8198), neuronal nitric oxide synthase (nNOS; rabbit, 1:500, Abcam, Cambridge, UK, ab76067) or tyrosine hydroxylase (TH; rabbit, 1:500, Sigma–Aldrich; T8700) overnight at 4°C. After washing five times in PBS, arteries were incubated with secondary antibodies (1:500, Thermo Fisher Scientific) for 2 hours at room temperature. Finally, sections were washed at least three times in PBS.

For immunostaining of brain and renal slices, the fixed brains and kidneys were sectioned into 50-μm coronary slices using a vibrating microtome. After blocking as described above, slices were incubated with primary antibodies against ChAT (goat, 1:50, Millipore; AB144P), Synapsin 1 (rabbit, 1:500, Sigma–Aldrich; S193), VAChT (rabbit, 1:4000, Sigma–Aldrich; SAB4200559), or TH (rabbit, 1:500, Sigma–Aldrich; T8700) overnight at 4°C, and then incubated with secondary antibodies (1:500, Thermo Fisher Scientific) for 2 hours at room temperature. Nuclei were labeled by incubating with Hoechst 33258 for 15 minutes at room temperature.

For immunostaining of the renal pelvis, mice were perfused with PBS followed by 4% paraformaldehyde as above described. With the aid of a surgical microscope, the fat in the renal hilum area was cleaned by repeated scraping with fine forceps, and the funnel-shaped renal pelvis was found by following the ureter. Renal pelvises were then dissected using micro scissors and post fixed with 4% paraformaldehyde overnight at 4°C. After blocking as described above, wholemount pelvises were incubated with primary antibodies against Synapsin 1 (rabbit, 1:500, Sigma–Aldrich, S193) overnight at 4°C and then incubated with secondary antibodies (1:500, Thermo Fisher Scientific) for 3 hours at room temperature. In the three-dimensional visualization of the renal pelvis wall, renal pelvises were incubated with Alexa Fluor 488-conjugated WGA (as a plasma membrane probe) and Hoechst 33258 overnight at 4°C to visualize the cytoarchitecture.

For immunostaining of the nodose/jugular complexes, fixed nodose/jugular complexes were blocked for 3 hours as above. After blocking as described above, complexes were incubated with primary antibodies against VGluT2 (rabbit, 1:500, Abcam; ab216463) overnight at 4°C, and then incubated with secondary antibodies (1:300, Thermo Fisher Scientific) for 3 hours at room temperature.

Fluorescence images of stained tissue samples were acquired under a confocal laser scanning microscope (LSM 710; Carl Zeiss, Jena, Germany). In the three-dimensional visualization of the renal pelvis wall, a median filter (radius, two pixels) and a deconvolution algorithm were applied to improve the signal-to-noise ratio.

Data Analyses

For measurements of tdTomato-positive and TH-immunoreactive nerve densities, images were performed skeletonization (created centerlines of nerve fibers in the image by a thinning algorithm^30,31) using ImageJ, and the length of nerve fibers was analyzed by the Analytic skeleton plugin (https://imagej.net/plugins/analyze-skeleton/). The nerve densities were given by dividing the total length of the nerve fibers by the area.

Analysis of the whole-brain three-dimensional image dataset was described in our previous study.³² Briefly, the whole-brain three-dimensional image dataset was nonrigidly transformed to match Allen adult mouse brain atlases (https://atlas.brain-map.org/). Then, the brain regions of interest were segmented through customized software developed in MATLAB R2017a (https://ww2.mathworks.cn/). Finally, the somata of GFP-positive neurons in each brain region were detected and counted using the Spot module of Imaris 7.6 (https://imaris.oxinst.com/). To visualize the distribution patterns of PRV-infected neurons in the whole brain, heatmaps were generated according to the number of detected neurons in the radial range of 250 μm.

For scRNA-Seq data analysis, Cell Ranger 6.1 software (https://www.10xgenomics.com/) was used to convert Illumina basecall files to fastq format. We used the FASTX-Toolkit 0.0.13 (http://hannonlab.cshl.edu/fastx_toolkit/) to generate raw sequence data. Next, the reads were aligned to the mm10 mouse reference genome. Gene counts and unique molecular identifier (UMI) counts were acquired by the Subread software package (https://sourceforge.net/projects/subread/). On the basis on gene counts and UMI counts, expression matrix files for subsequent analyses were generated. For quality control, cells with >25% reads mapping to mitochondria (aligned to the mouse mm10 transcriptome) or with >30,000 UMI counts were filtered out. All gene expression was normalized using the function “NormalizeData()” with a scaling factor of 10,000 (on the basis of the Seurat 4.0 toolkit; https://satijalab.org/seurat/). Then, the top 2000 variable genes were selected using the Seurat function “FindVariableFeautres (),” and the effect of mitochondrial genes was regressed out using the Seurat function “ScaleData().” Next, principal component analysis was performed on the basis of the 2000 highly variable genes. The T-distributed stochastic neighbor embedding (t-SNE) algorithm using the top 30 principal components was applied to visualize cells in the two-dimensional space, and cells were separated into clusters by Seurat function “FindClusters()” with a resolution of 0.6. We used the Seurat function “FindAllmarkers()” (on the basis of the Wilcox likelihood ratio test, with default parameters) to identify marker genes for each cluster. We identified major cell types on the basis of the expression values of the marker genes (log2 fold change >1, P<0.05 versus other clusters) and canonical cell-type genes. Because we focused on the renal artery and its segmental branches, cell types of other kidney tissues (specifically expressed Cldn10, Atp6v1g3, Rhbg, and Calb1)³³ were removed. Then, the retained 6354 cells were reclustered following the same procedure and parameter as described above. Loupe Browser 6.0 (https://support.10xgenomics.com/) was used to visualize t-SNE and to generate violin plots. MATLAB R2017a was used to generate the heatmap.

Statistical Analyses

All data are presented as the mean±SEM. The sample size (n) refers to the number of independent biologic experimental replicates. All histologic findings were replicated in at least three animals. We used Excel (Microsoft, Redmond, WA) and GraphPad Prism v8 (GraphPad Software, San Diego, CA) to generate graphs and statistics. No statistical method was used to predetermine sample size. We first determined whether the data values came from a normal distribution using the Kolmogorov–Smirnov test. Two-sided t tests or Mann–Whitney tests were then used to assess significance levels. Statistical significance was set at P<0.05. More details of statistical tests can be found in the figure legends and Supplemental Material.

Results

Cholinergic Nerve Fibers Traveled with the Renal Artery

A typical characteristic of autonomic neurotransmission is that sympathetic postganglionic nerves are predominantly noradrenergic, whereas those of the parasympathetic nervous system are cholinergic.^1,10 To check whether kidneys receive cholinergic nerve supply, we constructed Chat-ires-Cre:Ai14 transgenic mice, which report strong red fluorescence in Chat-expressing cells and their processes.^25,26 In this mouse strain, we observed dense tdTomato-positive nerve fibers running alongside renal arteries (Figure 1A), and the abundant tdTomato-positive nerve fibers eventually entered the renal hilum.

Figure 1.

Cholinergic nerve fibers around the renal artery. (A) Distribution of Chat-tdTomato positive nerve fibers (red) around the renal artery. The green dashed box in the schematic diagram indicates the imaging region. Scale bar, 200 μm. (B) Representative images of renal arteries from adult Chat-ires-Cre:Ai14 mice, stained for ChAT. Note the tdTomato (red) specifically expressed in ChAT-immunoreactive nerve fibers (green). Scale bars, 10 μm. (C) Representative images of renal arteries from adult Chat-ires-Cre:Ai14 mice, stained for VAChT. Note the tdTomato (red) specifically expressed in VAChT-immunoreactive nerve fibers (green). The white box regions are magnified and shown in the lower-left corner. Scale bars, 10 μm.

To confirm the cholinergic nerves around the renal artery further, we performed immunostaining against ChAT and VAChT, which are both reliable markers of cholinergic nerves.^34,35 On the Chat-ires-Cre:Ai14 mouse renal arteries, we found that there are tdTomato-positive nerve fibers that have ChAT and VAChT immunoreactivity (Figure 1, B and C). On the renal arteries of C57BL/6 mice, ChAT and VAChT immunoreactivity nerve fibers were also observed (Supplemental Figure 1). These results consistently suggested that renal arteries received the cholinergic nerve supply.

Renal Periarterial Cholinergic Nerves Are Separated from Both Sympathetic and Sensory Nerves

To evaluate the anatomic relationship between cholinergic, sensory, and sympathetic nerves around the renal artery, we dissected out renal arteries from Chat-ires-Cre:Ai14 mice and stained for CGRP and TH, which are widely accepted specific markers of sensory and sympathetic nerves, respectively.^36–38 At sufficient resolution (confocal imaging, equipped with a 63/1.46× oil-immersion objective), we found cholinergic, sensory, and sympathetic nerve fibers were often intertwined, but cholinergic nerve fibers were separated from both sensory and sympathetic nerve fibers (Figure 2, A–F, Supplemental Figure 2).

Figure 2.

Anatomic relationship between cholinergic, sensory, and sympathetic nerve fibers around the renal artery. (A) Representative images of renal arteries from adult Chat-ires-Cre:Ai14 mice, stained for CGRP. Scale bars, 20 μm. (B) Higher-magnification images of the light blue box regions in (A). Scale bars, 10 μm. (C) Higher-magnification images of the white box regions in (A). Scale bars, 10 μm. Note the separation between tdTomato-positive (red) and CGRP-immunoreactive (green) nerve fibers. (D) Representative images of renal arteries from adult Chat-ires-Cre:Ai14 mice, stained for TH. Scale bars, 100 μm. (E) Higher-magnification images of the light blue box regions in (D). Scale bars, 10 μm. (F) Higher-magnification images of the white box regions in (D). Scale bars, 10 μm. Note the separation between tdTomato-positive (red) and TH-immunoreactive (green) nerve fibers. (G) Comparison of the densities of tdTomato-positive and TH-immunoreactive nerve fibers surrounding the renal artery (n=10 arteries from five female adult mice, two-sided unpaired t test, mean±SEM).

We quantified the densities of tdTomato-positive and TH-immunoreactive nerve fibers along renal arteries. The results showed that tdTomato-positive nerve fibers were significantly less than TH-immunoreactive nerve fibers (Figure 2G). The density (fiber length/vessel surface area) of tdTomato-positive nerve fibers is 30.6±2.5 mm/mm² (n=10 arteries from five female adult mice, mean±SEM), and the density of TH-immunoreactive nerve fibers is 62.5±3.6 mm/mm² (n=10 arteries from five female adult mice, mean±SEM).

There Are Cholinergic Ganglion Cells within the Renal Nerve Plexus

Generally, parasympathetic postganglionic neurons are either located in discrete ganglia closed to the organs they innervated or dispersed in the walls of viscera.^1,10 To find the renal cholinergic ganglion cells, we characterized the ganglia along renal arteries using Chat-ires-Cre:Ai14 mice. In several mice (10 of 22 cases), we found the tdTomato-positive ganglion cells in the renal nerve plexus (Figure 3, A and B). The location of these discrete ganglion cells varied considerably. Immunostaining indicated that there are tdTomato-positive ganglion cells displaying ChAT and VAChT immunoreactivity but no TH immunoreactivity (Figure 3, C–E). These findings suggested that there are small ganglia close to the renal artery that supply cholinergic nerve fibers for the renal nerve plexus.

Figure 3.

Cholinergic ganglion cells within renal nerve plexus. (A) A schematic representation of the location of cholinergic ganglia and nerve tracts around the renal artery. (B) A confocal image shows the tdTomato-positive cells located in a renal nerve bundle. White arrows indicate tdTomato-positive (red) cells. Nuclei were labeled with Hoechst 33258 (blue). The white dashed line delineates the outline of the renal artery. The white box region was magnified and shown in the lower-left corner. Scale bars, 50 μm. (C) Representative images of renal ganglia from adult Chat-ires-Cre:Ai14 mice, stained for ChAT. Scale bars, 20 μm. (D) Representative images of renal ganglia from adult Chat-ires-Cre:Ai14 mice, stained for VAChT. Scale bars, 20 μm. (E) Representative images of renal ganglia from adult Chat-ires-Cre:Ai14 mice, stained for TH (green). Scale bars, 20 μm.

Segmental Renal Artery and Renal Pelvis Are Associated with Cholinergic Axons

Inside the kidney, we investigated the distribution of cholinergic nerve fibers through genetic label and immunostaining against both ChAT and VAChT. This study observed neither intrarenal ganglia nor cholinergic nerve fibers associated with any renal parenchymal cells. However, the Chat-tdTomato-positive varicosities associated with segmental renal arteries and renal pelvis were captured by confocal imaging (Figure 4). In the renal pelvis wall, we found the Chat-tdTomato positive varicosities distributed in the muscle layer and the theca externa (Supplemental Figure 3).

Figure 4.

Cholinergic nerve varicosities are associated with the segmental renal artery and pelvis. (A) Cross-sectional view of a segmental renal artery from Chat-ires-Cre:Ai14 mouse. Stained for synapsin 1 (green). The green box in the schematic diagram indicates the imaging region. Scale bars, 20 μm. (B) Higher-magnification images of the white box regions in (A). Scale bars, 20 μm. Arrows indicate tdTomato and synapsin double-positive varicosities. Colocalization analysis result of varicosities is shown on the right. (n=4 male mice, mean±SEM). (C) Representative images of the renal pelvises from Chat-ires-Cre:Ai14 mice, stained for synapsin 1 (green). Scale bars: whole-mount image, 200 μm; higher-magnification images, 20 μm. (D) Higher-magnification images of the white box regions in (C). Scale bars, 20 μm. Arrows indicate tdTomato and synapsin double-positive varicosities. Colocalization analysis result of varicosities is shown on the right. (n=4 male mice, mean±SEM).

In mammals, cholinergic components are ubiquitously expressed among widely diverse tissues, not just in the nervous system.^39,40 To investigate whether these intrarenal tdTomato-positive varicosities are neurogenic, we immunostained renal slices and renal pelvises of Chat-ires-Cre:Ai14 mice for synapsin 1 (a specific synaptic vesicles marker⁴¹). The results showed that 93.2%±2.8% tdTomato-positive varicosities around the segmental renal artery are synapsin immunoreactive (n=4 male mice; Figure 4B), and 93.8%±1% tdTomato-positive varicosities in the pelvic wall are synapsin immunoreactive (n=4 male mice; Figure 4D). Furthermore, we found that the cholinergic, sensory, and sympathetic nerves were separate fibers in the renal pelvis wall (Supplemental Figure 4). Overall, the above findings suggest that there are synaptic vesicle-carrying cholinergic nerves supplying the segmental renal artery and renal pelvis.

AChRs Are Expressed in the Renal Artery and Its Segmental Branches

In mammals, 16 nicotinic ACh receptor subtypes (nAChRs, α1–α7, α9, α10, β1–β4, γ, δ, and ɛ) and five muscarinic ACh receptor subtypes (mAChRs, M1–M5) have been indentified.⁴² To examine the expression of various AChRs in the renal artery and its segmental branches, we dissected it and performed scRNA-seq analysis (Figure 5, A and B). After an unsupervised clustering analysis of 6354 cells, eight distinct clusters were projected onto t-SNE plots (Figure 5C). On the basis of the expression values of known marker genes, we identified seven major cell types: smooth-muscle cells (SMCs; Tagln, Acta2, and Myh11;),⁴³ endothelial cells (ECs; Fbln2 and Bmx),⁴³ fibroblasts (Pdgfra and Tcf21),^44,45 myeloid cells (Cd68 and Csf1r),^46,47 Schwann cells (Plp1 and Cnp),^48,49 T cells (Cd3d and Cd3g), and B cells (Ms4a1 and Cd79a; Figure 5, D and E). In these cells, expression of multiple receptor subtypes was detected, with higher expression of mAChR3, nAChRa7, and nAChRβ1 (Figure 5F, Supplemental Figure 5, Supplemental Table 3). The mAChR3 expression in ECs is most pronounced across cell subpopulations (Figure 5G, Supplemental Table 3).

Figure 5.

Expression of AChRs in the renal artery and its segmental branches. (A) Several renal arteries and their segmental branches for scRNA-seq. Scale bar, 1 mm. (B) Workflow of scRNA-seq. Renal arteries and their segmental branches were digested to the single cell suspension. After barcoded cDNA library construction, sequencing was performed on a 10× Genomics platform, followed by the unsupervised clustering analysis. (C) T-SNE representation of 6354 cells. (D) Violin plots showing the expression distribution of selected marker genes across cell subpopulations. (E) Treemap showing the composition of arterial cells analyzed by unsupervised clustering analysis. (F) T-SNE plot showing the biased expression of mAChR3 in ECs. (G) Heatmap showing relative expression of 16 AChRs in seven major cell types of the renal artery and its segmental branches.

Glutamatergic Neurons Arising in the Nodose/Jugular Complexes Conduct the Renal Sensory Pathway to the NTS

Previous tracing studies revealed that the nodose ganglion contains cell bodies of neurons that project to the kidney.^6,50,51 To clarify the vagal afferent pathway from the kidney to brain, we injected the retrograde AAV-carrying Cre genes (AAVretro-hSyn-Cre) into the left kidney of Ai14 mice. RDN models were established as a control, and the efficacy of denervation was confirmed by the diminishment of TH-immunoreactive nerve fibers in the renal cortex (Figure 6, A and B). With the AAV retrograde tracing strategy, neurons that projected directly to the kidney were expected to be labeled with strong tdTomato fluorescence, including their long-range axonal projections (Figure 6C).

Figure 6.

Assessment of the brain-kidney vagal afferent pathway. (A) Representative images of renal sections at 7 days after the sham procedure (left) and RDN (right), stained for TH (white). Scale bars, 500 μm. The green box regions are magnified and shown below, respectively. Scale bars, 50 μm. (B) Densities of TH-immunoreactive nerve fibers in the renal cortex of sham-operated or RDN mice. (n=5 male adult C57BL/6J mice, two-sided unpaired t test with Welch’s correction, mean±SEM). (C) Cartoon illustrates the scheme for retrograde AAV tracing. (D) TdTomato-positive neurons (red) in ipsilateral nodose/jugular complex from sham-operated and RDN Ai14 mice, infected with AAVretro-hSyn-Cre. Scale bars, 100 μm. (E) Statistical results for (D) (n=5 male heterozygous Ai14 mice for each group, two-sided unpaired t test, mean±SEM). (F) TdTomato-positive neurons (red) in the ipsilateral nodose/jugular complex from Ai14 mice infected with AAVretro-hSyn-Cre, stained for VGluT2 (green). Scale bars, 50 μm. (G) Statistical results for (F) (n=5 male mice, mean±SEM). (H) TdTomato-positive axons (red) distributed in NTS. White lines delineate outlines of brain regions. Scale bar, 200 μm. A higher-magnification image of the white box region is shown on the right. Scale bar, 20 μm.

Three weeks after AAVretro-hSyn-Cre injection, tdTomato-positive neurons were found in nodose/jugular complexes (Figure 6D). Compared with sham-operated control mice, the number of tdTomato-positive neurons was significantly reduced in renal denervated mice (Figure 6E). Immunostaining for vesicular glutamate transporter 2 (VGluT2; a glutamatergic neuron specific marker⁵²) indicated that most of these tdTomato-positive neurons in nodose/jugular complexes are glutamatergic (93.7%±2.6% tdTomato-positive neurons were VGluT2-immunoreactive, n=5 male mice; Figure 6, F and G). Moreover, the tdTomato-positive nerve varicosities were found in the NTS (Figure 6H). Overall, these results suggest that glutamatergic afferent neurons in nodose/jugular complexes conduct the renal sensory pathway to the NTS.

Cranial Parasympathetic Efferents Project to the Kidney

Parasympathetic efferent nerves either originate from the certain cranial nerve nuclei of the brainstem or the intermediate gray matter of the sacral segments of spinal cord.^1,10 To assess whether the cranial parasympathetic efferents project to the kidney, we injected retrograde PRV-CMV-GFP into the left kidney of the Chat-ires-Cre:Ai14 and C57BL/6 mice, and evaluated the distribution of PRV-infected neurons in brain and spinal cord (Figure 7A). As revealed by whole-brain imaging, we found that GFP-positive neurons were mainly confined within the paraventricular hypothalamic nucleus and the medulla of brainstem (Figure 7, B and C, Supplemental Figure 6, Supplemental Table 4). Previous studies have shown that these regions critically regulate renal sympathetic nerve activity.⁵³ Moreover, we found a few PRV-infected neurons in the DMX (Figure 7C, Supplemental Figure 6, Supplemental Table 4).

Figure 7.

Assessment of the brain-kidney parasympathetic connection. (A) Cartoon illustrates the scheme for retrograde PRV tracing with sdVx or RDN. CTB tracing was used to confirm the successful vagotomy. (B) Workflow of the whole brain three-dimensional imaging. Scale bars, 2 mm. (C) Whole-brain three-dimensional view of the distribution of infected neurons after PRV-CMV-GFP injection. Modeled digital spots were derived from the GFP-positive neurons, and the spots located in the DMX were rendered in red. The surfaces of the DMX, paraventricular hypothalamic nucleus, and paragigantocellular reticular nucleus (PGRN) were rendered in light blue. Scale bar, 2 mm. A, anterior; P, posterior; D, dorsal; V, ventral; L, lateral; M, medial. (D) Numbers of GFP-positive neurons in DMX of the sdVx and sham-operated mice (n=5 male mice for each group, two-sided unpaired t test with Welch’s correction, mean±SEM). (E) Frequencies of GFP-positive neurons in left DMX of the renal denervated and sham-operated mice (n=5 male mice, two-sided Mann–Whitney test, mean±SEM). (F) PRV-infected cholinergic neurons in DMX. Scale bars, 50 μm. Arrows indicate Chat-tdTomato (red) and PRV-GFP (green) double-positive neurons. Dashed lines delineate the outlines of the DMX. Statistical result was shown on the right (n=5 male mice, mean±SEM). (G) The absence of PRV-labeled cells within the sacral 4. Scale bars, 200 μm. (H) The cell count results of Chat-tdTomato and PRV-GFP double-positive neurons (red lines) and all PRV-GFP-positive neurons (black lines) in representative coronal slices from multiple spinal cord levels (n=2 male mice).

Considered the disturbance from nonspecific viral spread, we established sdVx and RDN models as the controls. The efficacy of sdVx was confirmed by CTB tracing and enlarged stomachs (Supplemental Figure 7, A, B, and D). Compared with the sham-operated group, the number of PRV-infected neurons in the bilateral DMX was significantly reduced in sdVx mice (Figure 7D and Supplemental Figure 7C), and the frequency of PRV-infected neurons in the ipsilateral DMX (the number of GFP-positive neurons in the left DMX divided by the number in the bilateral DMX) was significantly reduced in renal denervated mice (Figure 7E, Supplemental Figure 8). In the DMX, colocalization analysis showed that most infected neurons are Chat-tdTomato positive (91%±3.6% PRV-infected neurons have tdTomato, n=5 male mice; Figure 7F). Immunostaining for ChAT indicated that the genetic labeling of cholinergic neurons in Chat-ires-Cre:Ai14 mice DMX is faithful (92.8%±0.7% tdTomato-positive neurons were ChAT-immunoreactive, n=3 female mice; Supplemental Figure 9). Together, these results suggest that a small population of cholinergic neurons in the DMX are components upstream of the renal nerves.

To assess the possibility of synaptic connection between the sacral parasympathetic preganglionic nuclei and the kidneys, we evaluated the distribution of infected neurons in the intermediolateral column of the spinal cords after PRV injection. As expected, we observed that PRV-infected neurons were mainly confined within the ipsilateral intermediolateral column of T9–L2 (Supplemental Figure 10). These regions were previously described to contain renal sympathetic preganglionic neurons.⁵³ In sacral spinal cords, there were no PRV-infected neurons (Figure 7, G and H, and Supplemental Figure 10). These results implied no renal nerves originating from the sacral cord.

Discussion

Cholinergic effects on renal function have been a subject of interest for decades. Numerous studies indicate that ACh vasodilates the renal artery,⁵⁴ increases renal blood flow,⁵⁵ promotes sodium excretion,⁵⁶ and regulates the contractile frequency of the renal pelvis.⁵⁷ Several routes have been described regarding the source of ACh in the kidney.^58,59 This study showed that the cholinergic axons supply the main renal artery, segmental renal artery, and renal pelvis wall. The newly discovered cholinergic nerves represent a more conventional path for ACh to arrive at the renal vasculature and pelvis.

Previous studies have found that there are small ganglia in the renal plexus, and the location, number, and size of these small ganglia vary considerably. In 1950, Mitchell reported small ganglia along the course of the renal nerves or near their points of entry into the kidney.⁶⁰ In 1968, McKenna et al. showed the single AChE-containing ganglion cell in the nerve bundle close to the renal hilus.¹¹ In 2016, Mompeo et al. reported small ganglia at the bifurcation of the hilar renal artery.⁶¹ Consistently, we found there are small ganglia in the renal plexus of the mouse (Figure 3). Furthermore, we found there are discrete cholinergic ganglion cells contained in these small ganglia, suggesting that parasympathetic postganglionic fibers relay in the renal plexus.

The existence of renal parasympathetic innervation was controversial. Previous tracing studies gave conflicting results. Using horseradish peroxidase and Fluoro-gold tracing, John et al. and Maeda et al. found the absence of labeled cells within the DMX, and hence concluded that the kidney does not have a parasympathetic innervation.^62,63 However, Vincent et al. detected a few isolated labeled cells in the DMX by horseradish peroxidase–WGA tracing, although these labels were ignored as sparse.⁵⁰ Recently, van Amsterdam et al. provided the distribution pattern of parasympathetic nerves on human renal artery by immunostaining for NOS.¹⁵ Consistently, nNOS–immunoreactive axons also supply the mouse renal artery and renal pelvis (Supplemental Figure 11). However, NOS is not a specific and selective marker for the parasympathetic nerves. Previous studies have shown that NOS is found in renal sensory nerves.⁶⁴ Considering that a typical characteristic of parasympathetic neurotransmission is that the postganglionic nerves are cholinergic,^1,10 we identify renal cholinergic nerves by visualizing the biosynthetic enzyme for ACh (ChAT) and the vesicular transporter for ACh (VAChT), which are widely accepted specific markers for cholinergic nerves to date.^65–70 Furthermore, renal sensory, sympathetic, and cholinergic nerve fibers were distinguished in this study. The new findings provide significant anatomic evidence for parasympathetic innervation of the kidney. The functional effect of induced cholinergic renal nerves remains uncertain. It would be essential to validate further the action of cholinergic nerves in the kidney because the functional effects will be the ultimate standard for parasympathetic innervation.

As a neurotropic virus that can retrogradely spread across synapses and replicate itself, recombinant PRV has become a powerful tool to map the functional connections between the central nervous system and peripheral organs.^71,72 Through PRV tracing, previous studies revealed the sympathetic and sensory connections between the central nervous system and the kidneys.^73–75 On the basis of our anatomic findings, there are cholinergic nerve varicosities in the kidney (Figure 4). We reassessed whether parasympathetic efferents project to the kidney by PRV tracing. Previous studies have shown that a high titer PRV is crucial for complete representation of neural pathways, especially for the neural pathways with fewer density terminal fields.^76,77 Thus, we hypothesize that exiguous cholinergic terminals in the kidney have limited ability to take up the virus, and the combination of high-titer virus injection and extensive imaging of intact brain regions may be necessary for complete characterization of the brain-kidney neural pathway. Except for the sympathetic nuclei that had been clarified in previous studies, a small number of PRV-infected neurons in the DMX (a cranial parasympathetic nucleus) were also found in this study through higher titer (10⁹ versus 10⁶–10⁸ pfu/ml^73–75) virus infection and sequential imaging of the brain slices (Figure 7, B and C, Supplemental Figure 6, Supplemental Table 4). The projection of vagal nerves to the kidneys seems very likely because both RDN and sdVx can weaken PRV infection in the DMX (Figure 7, D and E, Supplemental Figures 7 and 8).

The anatomic findings in this study indicated that parasympathetic nerves supply the main renal artery, segmental renal artery, and renal pelvis. In addition, we found AChR expression in the renal artery and its segmental branches and a vagal brain-kidney axis involved in the renal innervation. The new findings may contribute to the understanding of neural regulation of renal function. Moreover, further understanding of the renal innervation paradigm may promote the improvement of RDN strategies in the clinic.^21,78,79

Disclosures

All authors have nothing to disclose.

Funding

This work was supported by grants from the National Natural Science Foundation of China (Grant number: 31630029), the National Program on Key Basic Research Project of China (Grant number: 2015CB755603), the Science Fund for Creative Research Group of China (Grant number: 61721092), and the Director Fund of Wuhan National Laboratory for Optoelectronics, Huazhong University of Science and Technology.

Author Contributions

S. Zeng, X. Cheng, and X. Liu were responsible for conceptualization; S. Zeng was responsible for funding acquisition, project administration, and validation; X. Cheng wrote the original draft of the manuscript; X. Cheng and S. Zeng were responsible for the formal analysis; X. Cheng, S. Zeng, and Y. Zhang were responsible for the methodology; X. Cheng, Y. Zhang, R. Chen, H. Lv, and S. Qian were responsible for visualization; R. Chen, X. Cheng, S. Qian, X. Liu, and S. Zeng were responsible for data curation; S. Zeng and X. Liu were responsible for supervision and reviewed and edited the manuscript; and Y. Zhang was responsible for software.

Data Sharing Statement

All data are available in the main text or the Supplementary Materials. All whole-brain imaging datasets and scRNA-Seq data are stored at the Wuhan National Laboratory for Optoelectronics, Huazhong University of Science and Technology. Customized software and any datasets generated during the current study are available from the corresponding author upon reasonable request.

Supplemental Material

This article contains the following supplemental material online at http://jasn.asnjournals.org/lookup/suppl/doi:10.1681/ASN.2021111518/-/DCSupplemental.

Supplemental Figure 1. ChAT and VAChT immunoreactivity nerve fibers traveled with the renal artery of C57BL/6 mouse.

Supplemental Figure 2. Anatomical relationship between cholinergic and catecholaminergic nerve fibers around the renal artery at different ages.

Supplemental Figure 3. The Chat-tdTomato positive fibers located in the renal pelvis wall.

Supplemental Figure 4. Anatomical relationship between cholinergic, sympathetic, and sensory nerve fibers in the renal pelvis wall.

Supplemental Figure 5. Expression of individual AChRs in renal artery and its segmental branches.

Supplemental Figure 6. Assessment of the synaptic connection between the brain and the left kidney.

Supplemental Figure 7. CTB and PRV tracing with sdVx.

Supplemental Figure 8. Distribution of infected neurons in the DMX after PRV injection.

Supplemental Figure 9. Immunostaining indicates that tdTomato expression is faithful to cholinergic neurons in the DMX.

Supplemental Figure 10. Assessment of the synaptic connection between spinal cord and the left kidney.

Supplemental Figure 11. Nitrergic nerve fibers associated with the renal artery and pelvic wall of C57BL/6 mouse.

Supplemental Table 1. Key study purposes, techniques, and mouse strains.

Supplemental Table 2. Primary and secondary antibodies used in this study.

Supplemental Table 3. Expression of AChRs in renal artery and its segmental branches.

Supplemental Table 4. Number and frequency of PRV-infected neurons in brain regions (Excel file).

Supplemental Source Data (Excel file).