Abstract
Endothelial cells (ECs) of blood and lymphatic vessels have distinct identity markers that define their specialized functions. Recently, hybrid vasculatures with both blood and lymphatic vessel-specific features have been discovered in multiple tissues. Here, we identify the penile cavernous sinusoidal vessels (pc-Ss) as a new hybrid vascular bed expressing key lymphatic EC identity genes Prox1, Vegfr3,and Lyve1. Using single-cell transcriptome data of human corpus cavernosum tissue, we found heterogeneity within pc-S endothelia and observed distinct transcriptional alterations related to inflammatory processes in hybrid ECs in erectile dysfunction associated with diabetes. Molecular, ultrastructural, and functional studies further established hybrid identity of pc-Ss in mouse, and revealed their morphological adaptations and ability to perform lymphatic-like function in draining high-molecular-weight tracers. Interestingly, we found that inhibition of the key lymphangiogenic growth factor VEGF-C did not block the development of pc-Ss in mice, distinguishing them from other lymphatic and hybrid vessels analyzed so far. Our findings provide a detailed molecular characterization of hybrid pc-Ss and pave the way for the identification of molecular targets for therapies in conditions of dysregulated penile vasculature, including erectile dysfunction.
Introduction
Blood and lymphatic vessels fulfill important homeostatic functions and play a role in numerous diseases. The endothelial cells (ECs) of blood and lymphatic vessels develop during early embryogenesis and differentiate into distinct cell lineages, characterized by the expression of specific molecular markers. However, recent research shows remarkable plasticity and cell identity transitions within the endothelium during injury, disease and even normal physiological changes in the vasculature. In addition, organ-specific functional specialization is associated with certain vascular beds adopting ‘mixed’ blood–lymphatic vessel identities. For example, blood ECs (BECs) of liver sinusoids and high endothelial venules of secondary lymphoid organs express key lymphatic EC (LEC) markers LYVE1 and VEGFR3 (1). In addition, a ‘hybrid’ blood−lymphatic endothelial phenotype was discovered in ECs of the Schlemm’s canal (SC) of the eye (2, 3, 4), the ascending vasa recta of the renal medulla (5), and the remodeled spiral arteries of the placental decidua (6). Uniquely, these hybrid vasculatures are directly connected to blood vessels and in most cases blood perfused, yet express PROX1, which is considered the master transcriptional regulator of LEC fate (7). The molecular properties of hybrid vessels reflect their functional adaptation to perform ‘lymphatic-like’ functions in response to tissue demands, such as drainage of aqueous humor by the SC in the eye (1).
A remarkable example of a vascular bed that is exposed to varied environmental demands, including a range of pressure, stretch and shear forces, is the penile cavernous sinusoidal vasculature. Penile cavernous sinusoids (pc-Ss) are important for erectile functionality as these vessels entrap blood while engorging, thereby facilitating the organ’s rigidity. The penile cavernous tissues consist of pc-Ss that are embedded within sinusoidal smooth muscle (sSM), supporting connective tissue, and vessels that supply or drain blood, all enclosed by the tunica albuginea (8). Erectile functionality depends on a complex interplay between cavernous nerves and their closely associated sSM, regulating the blood flow from the helicine arteries into the pc-Ss. During the flaccid state, the pc-Ss are tonically constricted by sSMs and only slowly perfused by blood. In contrast, when erected, the sSM and pc-Ss relax due to released neurotransmitters, allowing for increased blood supply by the helicine artery and sinusoidal engorgement (9, 10). Erectile dysfunction (ED), a common condition that decreases quality of life in the male population, is presented by a wide range of psychological, physiological, as well as pathological causes. ED is a common comorbidity in other diseases, including type 1 diabetes (11). While structural abnormalities in pc-Ss have been shown to contribute to ED (12, 13), little is known about their molecular causes.
Contrary to the human penis, the mouse penis is anatomically segmented into the distal glans and the proximal body, connecting at a 90°-angle bend of the urethra (14). During the nonerected resting state, the mouse penis is internalized and projects outward during erection. Several different, functionally specialized cavernous tissues have been identified in the mouse penis, of which those found in the mouse penile body are considered to be analogous to the human corpus spongiosum and corpus cavernosum (15). Additionally, cavernous rigidity is supported in mice by the baculum bone (os penis) in the glans penis (16, 17).
Here we report that the blood-perfused pc-Ss display molecular features of hybrid vessels and display distinct transcriptomic alterations in ED. Like other hybrid vessels, they express the LEC master regulator Prox1 but, uniquely, develop independent of the key lymphangiogenic growth factor VEGF-C. Molecular characterization of pc-Ss can provide insight into their specialized functionality and identify potential therapeutic targets of the male reproductive system.
Materials and methods
Mice
R26-mTmG (18) and R26-tdTom (19) reporter lines were obtained from the Jackson Laboratory. Prox1-GFP (20), Cldn5-GFP (21), Cdh5-CreERT2 (22), and Vegfr3-CreERT2 (23) lines were described previously. All mice were maintained on a C57BL/6J genetic background. For AAV experiments, neonatal pups received a single intraperitoneal injection (10 µL) with 5 × 1010 viral particles of a recombinant AAV9 encoding the ligand-binding domains 1–4 of VEGFR3 fused to an mFc domain (mVEGFR31–4-mFc), or the ligand non-binding domains 4–7 of VEGFR3 fused to an mFc domain (mVEGFR34–7-mFc) (24). For induction of Cre recombinase in adult mice, tamoxifen, dissolved in peanut oil (10 mg/mL), was administered by oral gavage. In juvenile mice, 4-hydroxytamoxifen (4-OHT), dissolved in peanut oil (10 mg/mL), was administered by intraperitoneal injection as indicated in figures and/or legends. Experimental procedures on mice were approved by the Uppsala Animal Experiment Ethics Board (permit numbers 130/15 and 5.8.18-06383/2020) and performed in compliance with Swedish legislation.
Tracer experiments
To visualize blood perfusion, intravenous tail vein injection with biotinylated Lycopersicon esculentum (tomato) lectin (Vector Laboratories; B-1175-1) or L. esculentum (tomato) lectin, DyLight® 649 (Vector Laboratories; DL-1178-1) was followed by terminal anesthesia with 100 mg/kg Ketador (Richter Pharma AG) and 10 mg/kg Rompun (Bayer; 02-25-45), and cardiac perfusion fixation with 1% paraformaldehyde (PFA, Sigma, P6148) in PBS. Tissues were post-fixed for 1.5–2 h in 3% PFA in PBS at room temperature and further processed for immunofluorescence analysis.
To assess fluid and macromolecular uptake by penile cavernous sinusoids, 1 µL of lysine-fixable tetramethylrhodamine conjugated dextran, 2,000,000 MW (Invitrogen™, D7139; 10 mg/mL in sterile filtered PBS), was injected subcutaneously in the penile glans using a Hamilton syringe (34G). Animals were sacrificed 20 min after tracer injection by cervical dislocation. Penis and lymph nodes were dissected and fixed for 1.5–2 h in 3% PFA in PBS (penis) or overnight in 1% PFA in PBS at 4°C (lymph nodes) and further processed for immunofluorescence analysis.
Immunoblotting
To validate AAV transduction, whole blood was collected from the thoracic cavity prior to cardiac perfusion, followed by coagulation (30 min, room temperature) and serum (supernatant) collection. Serum was purified by centrifugation (two times for 10 min, 2000 g, 4°C) and consecutive collection on ice. Bromophenol blue and 5% β-mercaptoethanol were added to the obtained serum, boiled at 95°C for 5 min and used to detect the soluble VEGFR31–4-Ig or VEGFR34–7-Ig serum proteins by western blotting. Recombinant mouse VEGFR3 chimera protein (R&D Systems, 743-R3-100) was used as quantitative standard. Proteins were separated by sodium dodecyl sulfate−polyacrylamide gel electrophoresis (SDS-PAGE) and blotted onto polyvinylidene difluoride membranes. Mouse VEGFR3 domains 1−4 and 4−7 were detected by probing with the polyclonal goat anti-mouse VEGFR3 antibody (R&D Systems, AF743, 1:1000) against the extracellular domain of VEGFR3. Signal detection was done using donkey anti-goat secondary antibodies conjugated to horseradish peroxidase (0.375 µg/mL, Jackson ImmunoResearch) in combination with enhanced chemiluminescence solution (ThermoFisher Scientific, WP20005). Visualization was performed on a ChemiDoc MP imaging system (Biorad).
Immunofluorescence
Fixed penile tissue was decalcified in 0.5 M EDTA (pH 8) at room temperature for five days, with solution exchanged daily. The tissue was embedded in gelatin−albumin matrix and 60-90 µm coronal sections were prepared using a vibratome. The sections were permeabilized in 0.3% Triton X-100 in PBS (PBS-Tx), followed by blocking in 3% bovine serum albumin (BSA; Sigma, A3294-100G) in PBS‑Tx for 2 h rocking at room temperature. Subsequently, the sections were incubated with primary antibodies diluted in 1% BSA in PBS‑Tx for 2–3 days at 4°C and washed three times in PBS-Tx prior to a 2-h incubation with fluorescent‑conjugated secondary antibodies, which were diluted in 1% BSA in PBS‑Tx. The stained sections were washed for 10 min with PBS‑Tx, stained for 10 min with 4′,6-diamidino-2-phenylindole (DAPI; Sigma, D9542, 1:1000 in PBS), and washed extensively in PBS before mounting in RapiClear® 1.52 (SUNJin Lab; RC1522001).
Whole-mount penises and ears were fixed in 4% PFA in PBS at room temperature for 2 h, permeabilized for 10 min with PBS-Tx, blocked with 3% BSA in PBS-Tx and then incubated for 3 h at room temperature or overnight at 4°C with primary antibodies diluted in 1% BSA in PBS-Tx. After washing, secondary antibody and DAPI staining was performed as described earlier, followed by mounting in Fluoroshield (Sigma-Aldrich, F-6182). Alternatively, DAPI staining was omitted and samples were mounted in Fluoroshield with DAPI (Sigma-Aldrich, F-6057).
The following primary antibodies were used for immunofluorescence staining: goat anti-mouse VE-cadherin (R&D Systems, AF1002; 1:200), hamster anti-mouse PDPN (Developmental Studies Hybridoma Bank, 8.1.1-a; 1:200), rabbit anti-mouse LYVE1 (Reliatech, 103-PA50AG; 1:200), rat anti-mouse EMCN (Santa Cruz Biotechnology, sc-65495; 1:200), rat anti-mouse PLVAP (BD Biosciences, 550563; 1:200), rabbit anti-mouse AQP1 (BiCell Scientific, 550563; 1:100). Secondary antibodies conjugated to Alexa Fluor 488, Alexa Fluor 594, Alexa Fluor 647, Cy3, or Cy5 were obtained from Jackson ImmunoResearch and used at a concentration of 3.75 µg/mL.
Image acquisition
Displayed confocal images are either single or multiple tile scan images and represented as maximum intensity projections of z-stacks unless indicated differently. All confocal images were acquired using the Leica TCS SP8 or Stellaris 5 confocal microscope with the Leica LAS X software (Leica Microsystems). Following objectives were used with the Leica TCS SP8: HC FLUOTAR L 25×/0.95 W VISIR and HC PL APO CS2 63×/1.30 GLYC objectives. The Leica Stellaris 5 was used with: HC FLUOTAR L 25×/0.95 W VISIR and HC PL APO CS2 63×/1.30 GLYC objectives.
Two-photon images were acquired using the Leica SP8 DIVE system with the HPX APO 20×/1.0 water dipping objective and the infrared femtosecond Ti:Sapphire (multiphoton) laser (Spectra-Physics) set to 920 nm for excitation of GFP and 1240 nm for excitation of Lectin-DyLight® 649.
Scanning electron microscopy
Anesthetized animals were perfused with PBS for 2 min, followed by perfusion with EM fixative (2.5% glutaraldehyde (Sigma, G5882) and 0.1 M sodium cacodylate (Sigma, C4945) in 4% PFA (Sigma, P6148), pH adjusted to 7.4) for 2 min. The tissue was post-fixed in EM fixative for 2 h, followed by decalcification with 1:1 1 M sodium cacodylate buffer (pH 7.4) and 0.5 M EDTA (pH 8) at room temperature for five days with daily exchange of the decalcification solution. Fixed penile tissue was embedded in a gelatin-albumin matrix for vibratome sectioning. The coronal penile sections (200 µm) were kept in sodium cacodylate buffer until processing. For SEM imaging, vibratome sections were washed in 0.1M sodium cacodylate buffer (pH 7.4), post-fixed in 1% w/v unbuffered OsO4 for 1 h, washed three times in water, dehydrated in EtOH (20–100%, 20% steps) and then EtOH–acetone (70:30, 50:50, 30:70, 100%; 15 min steps), critical point dried in an Agar E3000 critical point dryer (Quorum Technologies Ltd., East Essex, UK), mounted on stubs and coated with Au with an Emitech K550X sputter device (Emitech Ltd., Kent, UK) before examining using a Philips XL 30 ESEM (Philips, Eindhoven, Netherlands) operated at 15 kV accelerating voltage. Images were recorded digitally.
Image analysis
Images were analyzed using Fiji ImageJ (version 2.0.0-rc-69/1.53c, http://imagej.nih.gov/ij/, NIH). Three-dimensional renderings were computed from two-photon images using the Imaris x64 software (v.9.5.1.) (Bitplane).
where r is the radius of the corresponding fenestration, Atotal is the total area of the quantified image, and Afenestration is the area of fenestrations presented in the quantified area, respectively. All images were acquired from one animal. Data are representative of seven representative detail images taken from one section and represented as mean ± s.e.m.
Flow cytometry
Mouse penises were divided into penile glans and body and cut into smaller pieces, followed by digestion in collagenase IV (Life Technologies; 5 mg/mL), DNase I (Roche; 0.1 mg/mL) and FBS (Life Technologies; 0.5%) in PBS at 37°C for 10–15 min, with constant shaking at 950 rpm. Collagenase activity was quenched by addition of EDTA to a final concentration of 2 mM and digestion products were filtered through 50 µm nylon filters (Sysmex) and washed with FACS buffer (PBS, 0.5% FBS, 2 mM EDTA). Afterwards, cells were immediately processed for immunostaining. The Fc receptor binding was blocked with rat anti-mouse CD16/CD32 (eBioscience, 14-0161-82) followed by an incubation with antibodies (all obtained from eBioscience) against Ter119 (48-5921-82), CD45 (48-0451-82), CD11b (17-0112-82), PECAM1 (25-0311-82), and PDPN (12-5381-82). The antibody staining was followed by an FACS buffer washing step, followed by staining for dead cells using SYTOX Blue (Life Technologies, L34961). Cells were analyzed on a BD LSR Fortessa cell analyzer (BD Biosciences) equipped with fixed laser lines (used: 405, 488, 561, and 643 nm). Single viable cells were gated from FSC-A/SSC-A, FSC-H/FSC-W and SSC-H/SSC-W plots and subsequent exclusion of dead cells. FMO controls were used to set up presented gating schemes, allowing to distinguish and quantify distinct cell populations. Acquired flow data were processed using using FlowJo software, version 10.5.0 (TreeStar).
scRNAseq data processing
Eight count matrices from human penile tissue samples, containing all cell types, were obtained from the GEO database (GSE206528), with individual sample accession numbers GSM6255907, GSM6255908, GSM6255909, GSM6255910, GSM6255911, GSM6255912, GSM6255913, and GSM6255914 (25). Samples were merged and filtered to keep only cells expressing more than 200 genes. Mitochondrial genes and genes expressed in less than three cells were removed. In addition, cells with a fraction of mitochondrial gene counts >6% were excluded, as well as counts of long noncoding MALAT1 RNA, which is abundantly expressed (26) and a known bias of scRNA-seq analysis. Samples were batch integrated by canonical correlation analysis (CCA) using Seurat (version 4.0.6) (27). Log normalization, dimensional reductions, graph-based clustering, UMAP visualization, and differentially expressed gene (DEG) analysis were completed following Seurat v4 documentation. ECs formed separate clusters identified by their top DEGs: EMCN, MMRN1, VWF, ACKR1, and CLDN5. EC clusters were extracted and the final dataset was obtained after cell type identification and removal of contaminants followed by reintegration and reclustering as described earlier. Contaminating cell types formed separated clusters of fibroblasts (DCN, LUM, FBLN1, COL1A2) or displayed interferon pathway expression patterns (RSAD2, ISG15, IFI44L). All analysis was performed using Rstudio (desktop version 2022.07.1) in R (version 4.2.1). A web application for data searching and visualization was generated using the shiny package of Rstudio (https://shiny.rstudio.com), and the package ShinyCell for database creation (28). Upregulated DEGs of combined ED compared to normal ECs were obtained for each cluster individually and further processed by GO enrichment analysis. Additional analysis was performed for cluster pc-S-EC2 comparing ED_DM and ED_nDM ECs to normal ECs separately. Hypergeometric distribution test was performed using GO stats (version 2.64.0). A universal gene list was obtained from org.Hs.eg.db (version 3.16.0) and gene sets included in the analysis contained 5−1000 genes of the human genome. Pathways were considered significant when passing thresholds of P < 0.00001 and gene count/term >10. Additionally, relevant GO terms were selected by OddsRatios and classified in seven different annotation categories, visualized using ggplot2 (version 3.4.2). Trajectory analysis was performed (curve not shown) using SCORPIUS (version 1.0.8) (29) with default parameters and k = 5. Cells were mapped along the trajectory. Cell order along the trajectory was retained when visualizing selected genes in a heatmap.
Statistical analysis
GraphPad Prism 9 was used for data visualization and statistical testing. Data between two groups were compared using the unpaired t-test with Welch’s correction. Differences were considered statistically significant when P < 0.05 and indicated on the graphs with star symbols: *P < 0.05, **P < 0.01 and ***P < 0.001. Hypergeometric distribution was assessed for GO enrichment analysis. DEG analysis was performed using Wilcoxon’s rank-sum test.
Results
Transcriptomics of human penile cavernous vasculature identifies EC populations with hybrid vessel identity
To characterize the molecular features of penile cavernous (pc) vasculature, we extracted endothelial cell (EC) transcriptomes, identified based on their expression of pan-endothelial markers (PECAM1, CDH5), from single-cell RNA sequencing (scRNA-seq) data obtained from human corpus cavernosum tissue (25). The dataset included samples from men without erectile dysfunction (n = 3, normal), as well as from nondiabetic (n = 3, ED_nDM) and type 1 diabetic (n = 2, ED_DM) men treated for erectile dysfunction (ED) (Fig. 1A).
Single cell transcriptomics of human penile cavernous ECs in health and in erectile dysfunction. (A) Schematic presentation of human penile cavernous tissue used for single cell RNA sequencing analysis. Analyzed dataset (25) included samples from men without erectile dysfunction (ED) (n = 3, normal), as well as from nondiabetic (n = 3, ED_nDM) and type 1 diabetic (n = 2, ED_DM) men treated for erectile dysfunction. (B) The five penile cavernous EC (pc-EC) clusters, from normal and ED tissue samples, visualized in a UMAP landscape and colored by cluster assignment. (C) Dot plots showing selected EC subtype markers for the five EC clusters presented in (B) between normal and ED pc-ECs. Dot size illustrates percentage of cells presenting transcript sequence counts and color illustrates the average scaled gene expression level (log2 fold change) within a cluster. (D) GO analysis of genes enriched in the clusters of the ED pc-ECs. Dot size illustrates odds ratio and color illustrates superordinate categories, with corresponding terms ordered by P-value along the x-axis.
Citation: Vascular Biology 6, 1; 10.1530/VB-23-0014
After removal of contaminating cells identified as fibroblasts and mural cells, which are commonly found in scRNA-seq datasets of vascular cells (30, 31), we obtained in total 21,274 high-quality ECs. By applying the Harmony method for batch correction and Seurat graph-based clustering approach, the cells were distributed into five clusters (Fig. 1B). ECs from individual samples and health states contributed to each cluster (Supplementary Fig. S1A, see section on supplementary materials given at the end of this article). Based on known molecular signatures and in line with previous analyses (25, 32), we annotated one cluster of penile cavernous arterial ECs (pc-A-ECs) expressing arterial markers (e.g. HEY1, GJA5, VEGFC), and three clusters of penile cavernous sinusoidal ECs (pc-S-EC1-3) expressing venous markers (e.g. EMCN, PLVAP, NR2F2) (Fig. 1B and C, Supplementary Table S1). An additional cluster was characterized by heterogenous expression of EC identity markers, potentially representing transition along the venous–capillary–arterial trajectory (pc-VcapA-EC) (Fig. 1B and C, Supplementary Table S1). Ordering of cells based on similarities in their gene expression patterns generated a linear trajectory that placed pc-VcapA-EC cluster in between pc-S-EC1 and pc-A-EC clusters, and revealed zonated expression of arterial, capillary, and venous markers previously identified in the brain vasculature (30) (Supplementary Fig. S1B). Unexpectedly, we found that the pc-S-EC clusters also expressed varying degrees of PROX1, which is the master regulator of LEC fate, along with other markers of lymphatic and hybrid EC identity (e.g. FOXC2, FLT4, LYVE1) (Fig. 1C). However, they did not express the bona fide LEC marker PDPN (Fig. 1C), suggesting hybrid EC identity. Additional cluster markers are provided in Table S1, and the data are made available for querying at https://makinenlab.shinyapps.io/HumanPenileCavernousEndothelialCells/.
We observed no major differences in the expression of venous and arterial EC identity markers in combined ED compared to the normal pc-A-EC and pc-VcapA-EC clusters (Fig. 1C). In contrast, the two clusters with a hybrid EC identity displayed an increased expression of lymphatic markers. PROX1 expression was elevated in pc-S-EC2, and LYVE1 expression was increased in pc-S-EC3 in combined ED disease states compared to the normal state (Fig. 1C, Supplementary Fig. 1C and Supplementary Table 1). To investigate pathological gene expression changes in pc-ECs globally, we performed Gene Ontology (GO) analysis of the upregulated, differentially expressed genes (DEGs) between the normal state and combined ED disease states. This analysis revealed a distinct disease-associated profile in the different EC types, with the selective enrichment of biological processes related to coagulation in pc-A-EC cluster, and inflammation in pc-S-EC2 cluster for combined ED (Fig. 1D and Supplementary Table 2). Further investigation showed that all inflammation related GO terms, except one, were specific to the ED_DM cohort (Table S2), suggesting a pronounced involvement of inflammatory processes in individuals with ED associated with diabetes. Other pc-EC clusters shared a more similar response, characterized by enrichment of processed related to steroid response, cytoskeletal organization and endothelial development (Fig. 1D and Supplementary Table 2).
Taken together, these results reveal zonation of human corpus cavernosum vasculature, and identify cavernous penile sinusoidal endothelium as a hybrid vascular bed with a distinct ED-associated changes in gene expression.
Murine penile cavernous sinusoids are blood-perfused hybrid vessels
To investigate if the molecular features of pc-Ss are conserved in the structurally similar murine penile vasculature, we analyzed the expression of EC identity markers using genetic reporter mice and immunofluorescence staining in 60–90 µm coronal sections of the penile glans and body. The cavernous tissues analyzed included corpus cavernosum glandis (CCg) (33) and corpus cavernosum uretrae (CCug) in the penile glans, as well as the corpus cavernosum uretrae (CCu) and the bilateral corpus cavernosa (CC) in the penile body (15) (Fig. 2A).
Characterization of the murine penile vasculature. (A) Schematic presentation of the murine penile anatomy. Blood vessels (pink), penile cavernous sinusoids (pc-Ss) (red), lymphatic vessels (green), baculum (penis bone, gray), and ureter (brown) are displayed. CCg, corpus cavernosum glandis; CCug, corpus cavernosum urethrae glandis; CCu, corpus cavernosum urethrae; CC, corpus cavernosum. (B, C) Vascular blood perfusion (visualized by lectin perfusion) and expression of Prox1-GFP in the penile glans and body. A magnified view of the boxed region in (B) is shown in (C). Red arrowheads point to blood-perfused Prox1-GFP-positive sinusoids, and blood vessels (red arrows) and lymphatic vessels (green arrowheads) are indicated. (D–F) Imaris rendering from two-photon images of cavernous tissue from 12-month-old R26‑mTmG;Vegfr3-CreERT2 mice, treated with 1 mg of tamoxifen on three consecutive days at 11 months of age, displayed in colored depth coding and showing different angles of a coronal section of CC (arrow) (D). Note colocalization of GFP (Vegfr3 expression) with intravenously injected lectin (purple) in CC and CCu but not in lymphatic vessels (LV) (E). (F) CCu (arrowhead) and CC (arrow) are displayed. Scale bars, 50 μm (C), 80 μm (D), 100 μm (F (right panel)), 200 μm (B, E, F (left panel)).
Citation: Vascular Biology 6, 1; 10.1530/VB-23-0014
The blood vessels were identified via intravenous injection of biotinylated Lectin, while lymphatic vessels were simultaneously visualized by the LEC marker Prox1 using the Prox1-GFP transgene (Fig. 2B). Similar to the hybrid vessel identity identified in human pc-S-ECs, we also found lectin+, i.e. blood-perfused pc-Ss that expressed Prox1-GFP (Fig. 2B and C, Supplementary Movie S1). Pc-S-ECs expressed another key LEC marker Vegfr3, as detected by GFP expression in a tamoxifen-treated R26‑mTmG;Vegfr3-CreERT2 reporter mouse (Fig. 2D–F), and immunofluorescence staining for VEGFR3 (Fig. S2A). Three-dimensional rendering of two-photon z-stack reconstruction of GFP+ penile vasculature in R26‑mTmG;Vegfr3-CreERT2 mice revealed lectin+ CC (Fig. 2D and E, Supplementary Movies S2 and S3) and CCu (Fig. 2F, Supplementary Movie S4), positioned ventral to the urethral lumen and extending into the urethral flaps. These results show that murine pc-Ss are blood-perfused vessels expressing the key LEC signature genes Prox1 and Vegfr3, suggesting conservation of their hybrid identity between mouse and man.
Heterogeneity, acquisition and maintenance of penile sinusoid identity
All murine penile cavernous tissues analyzed expressed the expected pan-endothelial markers, including Cdh5, encoding the junctional VE-cadherin protein (Supplementary Fig. S2B, Table 1), as well as the venous EC marker EMCN (Fig. 3A, Supplementary Fig. S2A). While the majority of pc-S-ECs in CCug and CCg of the penile glans showed prominent Prox1-GFP fluorescence (Fig. 3A), a smaller fraction of pc-S-ECs in the CCu and CC of the penile body were GFP+ (Supplementary Fig. S2C). We also observed heterogenous expression of the lymphatic vessel endothelial hyaluronan receptor 1 (LYVE1), while PDPN, the bona fide marker of mature lymphatic vessels, was not expressed in pc-S-ECs (Fig. 3B, Supplementary Fig. S2D, Table 1).
Prox1 expression in penile cavernous sinusoids of juvenile and aged mice. (A) Prox1-GFP expression, blood perfusion (determined by lectin perfusion) and immunofluorescence for EMCN is displayed in transversal sections of the penile glans (CCug and CCg). Veins (red arrows), lymphatic vessels (green arrowheads), and pc-Ss (red arrowheads) are indicated. (B) Immunofluorescence for LYVE1 and PDPN in vibratome sections of the glans (CCug and CCg). Lymphatic vessels (green arrowheads) and pc-Ss (red arrowheads) are indicated. (C, D) Representative gating schemes (C) and summary graph (D) for the analysis of BECs, LECs, and Prox1+ hybrid ECs in the penile glans (left panels) and body (right panels) of mature and aged mice. Data are representative of n = 6 (mature) and n = 7 (aged) mice from two independent experiments. (E) Prox1-GFP expression and immunofluorescence for LYVE1 and EMCN in CCg of juvenile (P21) mice. (F) Prox1-GFP expression and blood perfusion (determined by lectin perfusion) in CCg of mature (8-week-old) and aged (12-month-old) mice. Note expression of Prox1 in pc-Ss at all stages (red arrowheads), as well as in lymphatic vessels (green arrowheads). (G) Quantification of median fluorescent intensity (MFI) of Prox1-GFP expression in hybrid ECs (PECAM1+ PDPN−Prox1‑GFP+) of the glans of mature (8−10-week-old) and aged (10–12-month-old) mice by flow cytometry. Data represent mean (n = 6 (mature) and n = 7 (aged) mice from two independent experiments, indicated by different symbols) ± s.e.m. Unpaired t-test with Welch’s correction, **P < 0.01. Scale bars, 25 μm (A, B), 50 μm (E, F).
Citation: Vascular Biology 6, 1; 10.1530/VB-23-0014
Endothelial cell marker expression in penile cavernous sinusoids (pc-Ss). The analyzed markers in ECs of pc-Ss in comparison to previously reported expression in hybrid vessels: Schlemm’s canal (SC) (2, 3, 4, 56, 57, 58, 59, 60, 61), ascending vasa recta (AVR) (5, 50, 62, 63), and remodeled spiral artery (rSA) (6, 64, 65, 66, 67).
| Marker | pc-Ss | SC | AVR | rSA | |||
|---|---|---|---|---|---|---|---|
| CCug | CCg | CCu | CC | ||||
| Pan-EC markers | |||||||
| PECAM1 | + | + | + | + | + | + | + |
| TIE2 | + | + | + | + | + | + | + |
| VE-cadherin | + | + | + | + | + | NRF | + |
| VEGFR2 | + | + | + | + | + | − | NRF |
| BEC markers | |||||||
| AQP1 | + | + | + | + | + | − | + |
| CD34 | + | + | + | + | + | + | NRF |
| PLVAP | + | + | + | + | + | + | NRF |
| EMCN | + | + | + | + | + | + | − |
| vWF | + | + | + | + | + | NRF | + |
| LEC markers | |||||||
| PROX1 | + | + | +/− | +/− | + | + | + |
| VEGFR3 | + | + | + | + | + | + | + |
| LYVE1 | + | + | + | + | − | − | +/− |
| CCL21* | − | − | − | − | + | NRF | − |
| PDPN* | − | − | − | − | − | − | − |
+: expression detected; −: not detected; +/−: transient or discontinuous expression.
*Bona fide LEC markers.
CCug, corpora cavernosa uretrae glandis (glans); CCg, corpus cavernosum glandis (glans); CCu, corpus cavernosa uretrae (penile body); CC, corpus cavernosa (penile body); NRF, no reporting found.
Flow cytometry analysis of the murine penile tissue confirmed the presence of three PECAM1+ EC populations with blood (Prox1-GFP−PDPN−), lymphatic (Prox1-GFP+PDPN+), and hybrid (Prox1-GFP+PDPN−) vessel identities (Fig. 3C). Interestingly, we observed a relative decrease in the Prox1+ hybrid EC population (P = 0.0031, unpaired Student’s t-test with Welch correction), with a corresponding increase in the relative Prox1−BEC population (P = 0.0024) in penile glans of aged (10−12 months of age, n = 7) in comparison to mature (2−3 months of age, n = 6) mice (Fig. 3D).
The murine penis develops postnatally, with well-defined cavernous tissues being distinguishable from postnatal day (P)10 in the penile glans and body (33). Analysis of penile vasculature of juvenile 3-week-old mice showed EMCN+ pc-Ss that were weakly positive for Prox1-GFP fluorescence, but did not yet express LYVE1 (Fig. 3E). To study the maintenance of pc-S-EC identity during ageing, we analyzed vessel architecture and Prox1 expression in the penile tissues of mature (2−3 months) and aged (10−12 months) mice. Immunofluorescence analysis did not reveal differences in the morphology or blood perfusion of pc-Ss between the developmental stages, but Prox1-GFP fluorescence appeared weaker in aged mice (Fig. 3F). This finding was confirmed by flow cytometry analysis of hybrid ECs showing lower median fluorescence intensity (MFI) of Prox1-GFP in penile glans of aged in comparison to young mice (Fig. 3G). Interestingly, decline in PROX1 expression was previously associated with age-related deterioration of Schlemm’s canal, the hybrid vessel of the eye (3).
To investigate pc-S-EC heterogeneity in man with normal erectile function, we processed EC transcriptomes from normal human corpus cavernosum only. Besides a cluster of arterial ECs, we obtained four clusters of sinusoidal pc-S-ECs (Supplementary Figures S3A and B). Molecular heterogeneity was observed within the pc-S-EC population, in line with previous studies (25, 32), which included variable expression of the tight-junction molecule CLDN5 (Supplementary Fig. S3C), and high expression of the water channel aquaporin 1 (AQP1) selectively in the pc-S-EC1 cluster (Supplementary Fig. S3D). A similar heterogeneous expression of Cldn5 and AQP1 was also observed in vivo in murine penile tissue (Figures S3E and F).
Collectively, these results show that mouse and human pc-Ss are molecularly heterogenous and exhibit an identity similar to that of the previously described hybrid vasculatures of the eye (2, 3, 4), renal medulla (5), and placental decidua (6). Prox1 expression in pc-Ss initiates prior to reaching sexual maturity at 6–8 weeks, and is largely maintained throughout life but decreases with age.
Ultrastructural and functional features of murine penile sinusoids
Pc-Ss control the rigidity of the penis by entrapping incoming arterial blood, engorging and containing the blood within the sinusoidal spaces, which exerts pressure on the tunica albuginea and the occlusion of traversing veins. The sinusoidal endothelium is thus exposed to a range of pressure, stretch and shear forces likely to impact on their cell–cell junctions. Staining of whole-mounted tissue for VE-cadherin revealed torturous cell borders in LYVE1 positive pc-S-ECs in CCu (Fig. 4A) and CC (Supplementary Fig. S4A). To characterize the ultrastructural features of pc-Ss, we performed scanning electron microscopy of vibratome sections (Fig. 4B). Analysis of the luminal surface of pc-S-ECs revealed interdigitating and overlapping cell–cell contacts and protrusions (Fig. 4C). While the overall cell shape of pc-S-ECs shared some resemblance to the ‘oak leaf’ shape of lymphatic capillary ECs, we observed primarily a continuous distribution of VE-cadherin+ junctions with rare intercellular gaps (Fig. 4A), which distinguishes them from the button-like junctions in lymphatic endothelium (34). Consistent with the sinusoidal nature of pc-Ss, we observed abundant fenestrations (0.17 fenestrations/µm2 with 0.15% porosity and a diameter of 40–400 nm (mean 92.15 ± 2.89 nm, n = 319)) (Fig. 4D) within the CC of the penile body. In addition, we detected the expression of the plasmalemma vesicle-associated protein (PLVAP) in pc-S-ECs of all cavernous tissues analyzed (Supplementary Fig. S4B), suggesting the presence of filter-like diaphragms, previously found in fenestrae and transendothelial channels that confer vascular barrier function. However, PLVAP is also expressed on ECs without diaphragms, such as liver sinusoidal ECs that maintain transcellular pores of similar diameters (50–250 nm, mean 90.7 ± 11.7 nm), but higher density (8.45 fenestrations/µm2) and porosity (5.93%) (35, 36) compared to pc-S-ECs.
Ultrastructural and functional features of murine penile cavernous sinusoids. (A) Whole-mount immunofluorescence of CCu. Note the presence of continuous VE-cadherin+ cell–cell junctions (red arrowheads) and intercellular gaps (red arrows). (B, C) SEM analysis of the luminal surface of sinusoidal endothelium (color-coded in cyan in B), showing interdigitating and overlapping cell–cell contacts and protrusion (C, red arrowheads), and intercellular gaps and fenestrations (C, red arrows). (D) Quantification of fenestration size in pc-Ss of CC. Data are representative of seven images taken from one section and represented as mean ± s.e.m. (E) Experimental plan for the analysis of the uptake of TRITC−dextran tracer (2 × 106 MW; left panel). Twenty minutes after subcutaneous (SC) tracer injection, the tracer was taken up and visualized in the pc-Ss of the glans (middle panel) and in the sacral lymph node (right panel). Scale bars, 1 μm (C), 25 μm (A (right panel), B), 50 μm (A, left panel), 250 μm (E).
Citation: Vascular Biology 6, 1; 10.1530/VB-23-0014
To functionally assess potential macromolecular uptake by pc-Ss, we injected high-molecular-weight dextran tracer (2 × 106 MW) subcutaneously into the penile glans. As expected (37, 38), the tracer was drained by lymphatic vessels to lumbar and sacral lymph nodes (Fig. 4E). Although too large to enter blood capillaries, the tracer was unexpectedly detected within pc-Ss (Fig. 4E). The ultrastructural and functional analyses thus indicate a potential role of penile sinusoids in fluid and macromolecule uptake, similar to that of lymphatic vessels.
VEGF-C is not required for development of the penile hybrid vasculature in mice
The development of all lymphatic and previously described hybrid vascular beds is dependent on vascular endothelial growth factor C (VEGF-C) signaling via its receptor VEGFR3 (2, 6). To test if the regulatory mechanisms are conserved in pc-S formation, we inhibited VEGF-C using an adeno-associated virus (AAV) encoding a soluble ligand binding domain of VEGFR3 fused to the IgG Fc domain (mVEGFR31–4-mFc) that acts as a ligand trap (VEGF-C trap) (39). Control mice were treated with AAVs encoding the non-ligand-binding region of the VEGFR3 extracellular domain (mVEGFR34–7‑mFc). AAVs were administered to P6 pups, prior to the development of the cavernous tissues, by intraperitoneal injection and the penile tissue was analyzed at 5 weeks of age (Fig. 5A). In accordance with previous studies (24), this led to a systemic production of the trap molecule (Supplementary Fig. S5A) and blocked VEGF-C-induced lymphangiogenesis in the ear (Supplementary Fig. S5B). We also observed inhibition of penile lymphatic vessel formation (Fig. 5B), which was associated with edematous tissue swelling (Supplementary Fig. S5C). VEGF-C trap-treated mice additionally displayed developmental defects of the penis (Supplementary Fig. S5D) and phenotypic characteristics of cryptorchidism, with testicles being positioned suprascrotally inside the abdomen (Supplementary Fig. S5E). However, no apparent differences in the morphology of pc-Ss or the expression of pc-S-EC markers were observed in mice treated with AAVs encoding the VEGF-C trap compared to control AAVs (Fig. 5C). This demonstrates that the development of penile cavernous sinusoids and acquisition of their identity is independent of VEGF-C signaling.
The lack of effect of VEGF-C inhibition on the development of murine penile cavernous sinusoids. (A) Treatment scheme for inhibition of VEGF-C signaling prior to cavernous tissue development using the soluble VEGF-C trap (AAV-sVEGFR3) or control trap (AAV-Ctrl). (B, C) Immunofluorescence of vibratome sections of penile glans showing absence of PDPN+ lymphatic vessels (green arrowheads) but normal appearance of EMCN+ LYVE1+ pc-Ss (red arrowheads) in AAV-sVEGFR3-treated animals. Scale bars, 25 μm (C), 50 μm (B).
Citation: Vascular Biology 6, 1; 10.1530/VB-23-0014
Discussion
ECs of blood and lymphatic vessels represent differentiated cell lineages defined by expression of unique identity markers. Recently, hybrid vasculatures that possess features and functions of both blood and lymphatic vessels have been identified in multiple tissues, including the Schlemm’s canal in the eye. This has enabled identification of new molecular targets for the treatment of glaucoma (40, 41, 42, 43), which is associated with defective Schlemm’s canal function (44, 45, 46). In this study, we identify the penile cavernous sinusoids as a new hybrid vasculature that is characterized by the expression of key LEC regulators, including Prox1, and show distinct transcriptional alterations in ED.
Considering that ED is predominantly a disease of vascular origin, there is a striking lack of understanding of the specialized penile vasculature. In the non-erected, flaccid state the pc-Ss are minimally blood perfused. During an erection, pc-Ss fill, engorge and retain blood, thereby facilitating the tissue rigidity required for successful mating. This uniquely exposes pc-Ss to highly varying interstitial pressure, stretching and shear stress, which is likely to impact their gene expression. Previous transcriptome studies of ECs of human penile cavernous tissue (30, 31) identified populations of arterial and venous/sinusoidal ECs, with the latter showing high heterogeneity as also observed by us. Unexpectedly, we additionally found that human and murine pc-S-ECs express the master regulators of LEC fate and lymphangiogenesis, Prox1 and Vegfr3, respectively. Moreover, similar to the previously identified hybrid vasculatures (1), they did not express the marker of mature lymphatic vessels, PDPN. We further found that murine pc-Ss acquire a hybrid vascular identity prior to sexual maturity, but the expression of Prox1 in pc-S-ECs of the penile glans declined in aged compared to young mice. Notably, an age−related and pathological decline in PROX1 expression was described in the Schlemm’s canal and correlated with its functionality (3). A tight control of PROX1 levels is also required for the maintenance of LEC identity and lymphatic vessel function (47). Hence, it will be of interest to understand if regulation of hybrid identity of pc-S-ECs correlates with the functionality of pc-Ss and erectile function during aging.
Both murine and human CCu and CC lack a lymphatic vascular system responsible for draining interstitial fluid in most other tissues. It is currently unknown to what extent they participate in this process through the tough fibroelastic covering of the erectile tissues, known as the tunica albuginea. Our findings suggest that pc-Ss may contribute to the drainage function by their ability to take up fluid and high-molecular-weight tracers. This is consistent with the presence of fenestrae and AQP1 water channels in pc-S-ECs. Interestingly, prehospital systemic vascular access and resuscitation via the corpus cavernosa has been established as an effective measures for hypovolemic male casualties (48, 49), but no further investigations have been conducted on the mechanisms of saline uptake by pc-Ss.
Unlike other lymphatic and hybrid vasculatures analyzed so far (2, 6, 50), we found that the development of the murine penile hybrid vasculature was not dependent on VEGF-C signaling. Neonatal inhibition of VEGF-C using AAV-encoded soluble VEGFR3, concomitant with the inhibition of penile lymphatic vessel formation, unexpectedly resulted in defective penile development, phenotypic characteristics of cryptorchidism, and penile edema. While the pronounced penile edema can be explained by the complete lack of lymphatic vessels, the defective penile development and phenotypic characteristics of cryptorchidism were unexpected and the underlying mechanisms remain unclear. Interestingly, cases of cryptorchidism were described in patients with Noonan syndrome, lymphedema distichiasis, and Hennekam syndrome (51, 52), suggesting a potential link between lymphatic and testicular development. It is also possible that VEGF-C targets non-endothelial cells in the male genital tract, as was previously reported for VEGF that is able to bind its receptors VEGFR1 and VEGFR2 in the Leydig cells of the testis (53). Studies on the mechanisms of pc-S development and cellular origin of pc-S-ECs using inducible Cre/loxP-based approaches are hampered due to sensitivity of penile development to the antiestrogen tamoxifen used as an inducing agent (16, 54). Tamoxifen treatment during neonatal period leads to defects in the formation of epithelial spines of the penile glans and the bone, and cavernous tissue atrophia ((55)).
In summary, our study establishes pc-Ss as a new specialized hybrid vasculature that exhibits distinct transcriptional alterations in patients with erectile dysfunction, characterized by an upregulation of lymphatic markers and altered expression of genes related to immune regulation. Our characterization of pc-Ss and the associated searchable Web application for exploring transcriptome data of pc-ECs (https://makinenlab.shinyapps.io/HumanPenileCavernousEndothelialCells/) may help identify new molecular targets for the treatment of conditions associated with dysregulated penile vasculature, such as ED.
Supplementary materials
This is linked to the online version of the paper at https://doi.org/10.1530/VB-23-0014.
Declaration of interest
The authors declare that there is no conflict of interest that could be perceived as prejudicing the impartiality of the study reported.
Funding
This work was supported by grants from Knut and Alice Wallenberg Foundation (2018.0218, 2020.0057) (TM), the Swedish Research Council (2020-0269) (TM), Göran Gustafsson foundation (TM), and the European Union’s Horizon 2020 research and innovation program under the Marie Skłodowska-Curie grant agreement No 814316 (TM, MK, HS).
Acknowledgements
We thank Geoffrey Daniel (Swedish University of Agricultural Sciences, Uppsala) for SEM imaging and support in sample preparation. We also thank Sagrario Ortega (CNIO, Madrid) for the Vegfr3-CreERT2 mice, Ralf Adams (Max Planck Institute for Molecular Biomedicine, Münster) for the Cdh5-CreERT2 mice, Christer Betsholtz (Uppsala University, Uppsala) for the Cldn5-GFP mice, Kari Alitalo and the HelVi-AAVC core facility (University of Helsinki, Helsinki) for the AAVs, and Marie Jeansson (Uppsala University, Uppsala) for tissues samples and discussion. BioVis facility (Uppsala University) is acknowledged for flow cytometer and microscopy usage and support and Sofie Segerqvist Lunell and Aissatu Mami Camara for technical assistance.
References
- 1↑
Pawlak JB, & Caron KM. Lymphatic programing and specialization in hybrid vessels. Frontiers in Physiology 2020 11 114. (https://doi.org/10.3389/fphys.2020.00114)
- 2↑
Aspelund A, Tammela T, Antila S, Nurmi H, Leppänen VM, Zarkada G, Stanczuk L, Francois M, Mäkinen T, Saharinen P, et al.The Schlemm’s canal is a VEGF-C/VEGFR-3–responsive lymphatic-like vessel. Journal of Clinical Investigation 2014 124 3975–3986. (https://doi.org/10.1172/JCI75395)
- 3↑
Park DY, Lee J, Park I, Choi D, Lee S, Song S, Hwang Y, Hong KY, Nakaoka Y, Makinen T, et al.Lymphatic regulator PROX1 determines Schlemm’s canal integrity and identity. Journal of Clinical Investigation 2014 124 3960–3974. (https://doi.org/10.1172/JCI75392)
- 4↑
Truong TN, Li H, Hong YK, & Chen L. Novel characterization and live imaging of Schlemm’s canal expressing Prox-1. Acott TS, editor. PLoS One 2014 9 e98245. (https://doi.org/10.1371/journal.pone.0098245)
- 5↑
Kenig-Kozlovsky Y, Scott RP, Onay T, Carota IA, Thomson BR, Gil HJ, Ramirez V, Yamaguchi S, Tanna CE, Heinen S, et al.Ascending vasa recta are angiopoietin/Tie2-dependent lymphatic-like vessels. Journal of the American Society of Nephrology 2018 29 1097–1107. (https://doi.org/10.1681/ASN.2017090962)
- 6↑
Pawlak JB, Bálint L, Lim L, Ma W, Davis RB, Benyó Z, Soares MJ, Oliver G, Kahn ML, Jakus Z, et al.Lymphatic mimicry in maternal endothelial cells promotes placental spiral artery remodeling. Journal of Clinical Investigation 2019 129 4912–4921. (https://doi.org/10.1172/JCI120446)
- 7↑
Hong YK, Harvey N, Noh YH, Schacht V, Hirakawa S, Detmar M, & Oliver G. Prox1 is a master control gene in the program specifying lymphatic endothelial cell fate. Developmental Dynamics 2002 225 351–357. (https://doi.org/10.1002/dvdy.10163)
- 8↑
Hsieh CH, Liu SP, Hsu GL, Chen HS, Molodysky E, Chen YH, & Yu HJ. Advances in understanding of mammalian penile evolution, human penile anatomy and human erection physiology: clinical implications for physicians and surgeons. Medical Science Monitor 2012 18 RA118–RA125. (https://doi.org/10.12659/MSM.883201)
- 9↑
Priviero FBM, Toque HAF, Nunes KP, Priolli DG, Teixeira CE, & Webb RC. Impaired corpus cavernosum relaxation is accompanied by increased oxidative stress and up-regulation of the rho-kinase pathway in diabetic (db/db) Mice. Ushio-Fukai M, editor. PLoS One 2016 11 e0156030. (https://doi.org/10.1371/journal.pone.0156030)
- 10↑
Keegan KA, & Penson DF. Vasculogenic erectile dysfunction. Vascular Medicine 2013 341–348. (https://doi.org/10.1016/B978-1-4377-2930-6.00028-8)
- 11↑
Maiorino MI, Bellastella G, Della Volpe E, Casciano O, Scappaticcio L, Cirillo P, Giugliano D, & Esposito K. Erectile dysfunction in young men with type 1 diabetes. International Journal of Impotence Research 2017 29 17–22. (https://doi.org/10.1038/ijir.2016.38)
- 12↑
Huang T, Wang G, Hu Y, Shi H, Wang K, Yin L, & Peng B. Structural and functional abnormalities of penile cavernous endothelial cells result in erectile dysfunction at experimental autoimmune prostatitis rat. Journal of Inflammation 2019 16 20. (https://doi.org/10.1186/s12950-019-0224-0)
- 13↑
Hashimoto D, Kajimoto M, Ueda Y, Hyuga T, Fujimoto K, Inoue S, Suzuki K, Kataoka T, Kimura K, & Yamada G. 3D reconstruction and histopathological analyses on murine corporal body. Reproductive Medicine and Biology 2021 20 199–207. (https://doi.org/10.1002/rmb2.12369)
- 14↑
Rodriguez E, Weiss DA, Yang JH, Menshenina J, Ferretti M, Cunha TJ, Barcellos D, Chan LY, Risbridger G, Cunha GR, et al.New insights on the morphology of adult mouse penis. Biology of Reproduction 2011 85 1216–1221. (https://doi.org/10.1095/biolreprod.111.091504)
- 15↑
Piao S, Ryu JK, Shin HY, Han JY, Lee HS, & Suh JK. The mouse as a model for the study of penile erection: moving towards a smaller animal. International Journal of Andrology 2007 30 452–457. (https://doi.org/10.1111/j.1365-2605.2006.00737.x)
- 16↑
Weiss DA, Rodriguez E, Cunha T, Menshenina J, Barcellos D, Chan LY, Risbridger G, Baskin L, & Cunha G. Morphology of the external genitalia of the adult male and female mice as an endpoint of sex differentiation. Molecular and Cellular Endocrinology 2012 354 94–102. (https://doi.org/10.1016/j.mce.2011.08.009)
- 17↑
Hennefarth MR, Chen L, Wang B, Lue TF, Stoller ML, Lin G, Kang M, & Ho SP. Physicochemical and biochemical spatiotemporal maps of a mouse penis. Journal of Biomechanics 2020 101 109637. (https://doi.org/10.1016/j.jbiomech.2020.109637)
- 18↑
Muzumdar MD, Tasic B, Miyamichi K, Li L, & Luo L. A global double-fluorescent Cre reporter mouse. Genesis 2007 45 593–605. (https://doi.org/10.1002/dvg.20335)
- 19↑
Madisen L, Zwingman TA, Sunkin SM, Oh SW, Zariwala HA, Gu H, Ng LL, Palmiter RD, Hawrylycz MJ, Jones AR, et al.A robust and high-throughput Cre reporting and characterization system for the whole mouse brain. Nature Neuroscience 2010 13 133–140. (https://doi.org/10.1038/nn.2467)
- 20↑
Choi I, Chung HK, Ramu S, Lee HN, Kim KE, Lee S, Yoo J, Choi D, Lee YS, Aguilar B, et al.Visualization of lymphatic vessels by Prox1-promoter directed GFP reporter in a bacterial artificial chromosome-based transgenic mouse. Blood 2011 117 362–365. (https://doi.org/10.1182/blood-2010-07-298562)
- 21↑
Zhang Y, Daubel N, Stritt S, & Mäkinen T. Transient loss of venous integrity during developmental vascular remodeling leads to red blood cell extravasation and clearance by lymphatic vessels. Development 2018 145. (https://doi.org/10.1242/dev.156745)
- 22↑
Wang Y, Nakayama M, Pitulescu ME, Schmidt TS, Bochenek ML, Sakakibara A, Adams S, Davy A, Deutsch U, Lüthi U, et al.Ephrin-B2 controls VEGF-induced angiogenesis and lymphangiogenesis. Nature 2010 465 483–486. (https://doi.org/10.1038/nature09002)
- 23↑
Martinez-Corral I, Stanczuk L, Frye M, Ulvmar MH, Diéguez-Hurtado R, Olmeda D, Makinen T, & Ortega S. Vegfr3-CreER (T2) mouse, a new genetic tool for targeting the lymphatic system. Angiogenesis 2016 19 433–445. (https://doi.org/10.1007/s10456-016-9505-x)
- 24↑
Mäkinen T, Jussila L, Veikkola T, Karpanen T, Kettunen MI, Pulkkanen KJ, Kauppinen R, Jackson DG, Kubo H, Nishikawa S, et al.Inhibition of lymphangiogenesis with resulting lymphedema in transgenic mice expressing soluble VEGF receptor-3. Nature Medicine 2001 7 199–205. (https://doi.org/10.1038/84651)
- 25↑
Zhao L, Han S, Su H, Li J, Zhi E, Li P, Yao C, Tian R, Chen H, Chen H, et al.Single-cell transcriptome atlas of the human corpus cavernosum. Nature Communications 2022 13 4302. (https://doi.org/10.1038/s41467-022-31950-9)
- 26↑
Zhang B, Arun G, Mao YS, Lazar Z, Hung G, Bhattacharjee G, Xiao X, Booth CJ, Wu J, Zhang C, et al.The lncRNA Malat1 is dispensable for mouse development but its transcription plays a cis-regulatory role in the adult. Cell Reports 2012 2 111–123. (https://doi.org/10.1016/j.celrep.2012.06.003)
- 27↑
Hao Y, Hao S, Andersen-Nissen E, Mauck WM, Zheng S, Butler A, Lee MJ, Wilk AJ, Darby C, Zager M, et al.Integrated analysis of multimodal single-cell data. Cell 2021 184 3573–3587.e29. (https://doi.org/10.1016/j.cell.2021.04.048)
- 28↑
Ouyang JF, Kamaraj US, Cao EY, & Rackham OJL. ShinyCell: simple and sharable visualisation of single-cell gene expression data. Bioinformatics 2021 37 3374–3376. (https://doi.org/10.1093/bioinformatics/btab209)
- 29↑
Cannoodt R, Saelens W, Sichien D, Tavernier S, Janssens S, Guilliams M, Lambrecht B, De Preter K, & Saeys Y. SCORPIUS improves trajectory inference and identifies novel modules in dendritic cell development. Bioinformatics 2016. (https://doi.org/10.1101/079509)
- 30↑
Kalucka J, de Rooij LPMH, Goveia J, Rohlenova K, Dumas SJ, Meta E, Conchinha NV, Taverna F, Teuwen LA, Veys K, et al.Single-cell transcriptome atlas of murine endothelial cells. Cell 2020 180 764–779.e20. (https://doi.org/10.1016/j.cell.2020.01.015)
- 31↑
Muhl L, He L, Sun Y, Andaloussi Mäe M, Pietilä R, Liu J, Genové G, Zhang L, Xie Y, Leptidis S, et al.The SARS-CoV-2 receptor ACE2 is expressed in mouse pericytes but not endothelial cells: implications for COVID-19 vascular research. Stem Cell Reports 2022 17 1089–1104. (https://doi.org/10.1016/j.stemcr.2022.03.016)
- 32↑
Fang D, Tan XH, Song WP, Gu YY, Pan JC, Yang XQ, Song WD, Yuan YM, Peng J, Zhang ZC, et al.Single-cell RNA sequencing of human corpus cavernosum reveals cellular heterogeneity landscapes in erectile dysfunction. Frontiers in Endocrinology 2022 13 874915. (https://doi.org/10.3389/fendo.2022.874915)
- 33↑
Murakami R. A histological study of the development of the penis of wild-type and androgen-insensitive mice. Journal of Anatomy 1987 153 223–231.
- 34↑
Baluk P, Fuxe J, Hashizume H, Romano T, Lashnits E, Butz S, Vestweber D, Corada M, Molendini C, Dejana E, et al.Functionally specialized junctions between endothelial cells of lymphatic vessels. Journal of Experimental Medicine 2007 204 2349–2362. (https://doi.org/10.1084/jem.20062596)
- 35↑
Auvinen K, Lokka E, Mokkala E, Jäppinen N, Tyystjärvi S, Saine H, Peurla M, Shetty S, Elima K, Rantakari P, et al.Fenestral diaphragms and PLVAP associations in liver sinusoidal endothelial cells are developmentally regulated. Scientific Reports 2019 9 15698. (https://doi.org/10.1038/s41598-019-52068-x)
- 36↑
Cogger VC, O’Reilly JN, Warren A, & Le Couteur DG. A standardized method for the analysis of liver sinusoidal endothelial cells and their fenestrations by scanning electron microscopy. Journal of Visualized Experiments 2015 52698 e52698. (https://doi.org/10.3791/52698)
- 37↑
Supersaxo A, Hein WR, & Steffen H. Effect of molecular weight on the lymphatic absorption of water-soluble compounds following subcutaneous administration. Pharmaceutical Research 1990 7 167–169. (https://doi.org/10.1023/A:1015880819328)
- 38↑
Bernard FC, Kaiser J, Raval SK, Nepiyushchikh ZV, Doan TN, Willett NJ, & Dixon JB. Multichromatic near-infrared imaging to assess interstitial lymphatic and venous uptake in vivo. Journal of Biomedical Optics 2021 26. (https://doi.org/10.1117/1.JBO.26.12.126001)
- 39↑
Fang S, Nurmi H, Heinolainen K, Chen S, Salminen E, Saharinen P, Mikkola HK, & Alitalo K. Critical requirement of VEGF-C in transition to fetal erythropoiesis. Blood 2016 128 710–720. (https://doi.org/10.1182/blood-2015-12-687970)
- 40↑
Thomson BR, Liu P, Onay T, Du J, Tompson SW, Misener S, Purohit RR, Young TL, Jin J, & Quaggin SE. Cellular crosstalk regulates the aqueous humor outflow pathway and provides new targets for glaucoma therapies. Nature Communications 2021 12 6072. (https://doi.org/10.1038/s41467-021-26346-0)
- 41↑
Brigell M, Withers B, Buch A, & Peters KG. Tie2 activation via VE-PTP inhibition with Razuprotafib as an adjunct to latanoprost in patients with open angle glaucoma or ocular hypertension. Translational Vision Science and Technology 2022 11 7. (https://doi.org/10.1167/tvst.11.1.7)
- 42↑
Thomson BR, Carota IA, Souma T, Soman S, Vestweber D, & Quaggin SE. Targeting the vascular-specific phosphatase PTPRB protects against retinal ganglion cell loss in a pre-clinical model of glaucoma. eLife 2019 8 e48474. (https://doi.org/10.7554/eLife.48474)
- 43↑
Li G, Nottebaum AF, Brigell M, Navarro ID, Ipe U, Mishra S, Gomez-Caraballo M, Schmitt H, Soldo B, Pakola S, et al.A small molecule inhibitor of VE-PTP activates Tie2 in Schlemm’s canal increasing outflow facility and reducing intraocular pressure. Investigative Ophthalmology and Visual Science 2020 61 12. (https://doi.org/10.1167/iovs.61.14.12)
- 44↑
Souma T, Tompson SW, Thomson BR, Siggs OM, Kizhatil K, Yamaguchi S, Feng L, Limviphuvadh V, Whisenhunt KN, Maurer-Stroh S, et al.Angiopoietin receptor TEK mutations underlie primary congenital glaucoma with variable expressivity. Journal of Clinical Investigation 2016 126 2575–2587. (https://doi.org/10.1172/JCI85830)
- 45↑
Thomson BR, Souma T, Tompson SW, Onay T, Kizhatil K, Siggs OM, Feng L, Whisenhunt KN, Yanovitch TL, Kalaydjieva L, et al.Angiopoietin-1 is required for Schlemm’s canal development in mice and humans. Journal of Clinical Investigation 2017 127 4421–4436. (https://doi.org/10.1172/JCI95545)
- 46↑
Thomson BR, Heinen S, Jeansson M, Ghosh AK, Fatima A, Sung HK, Onay T, Chen H, Yamaguchi S, Economides AN, et al.A lymphatic defect causes ocular hypertension and glaucoma in mice. Journal of Clinical Investigation 2014 124 4320–4324. (https://doi.org/10.1172/JCI77162)
- 47↑
Wigle JT, Harvey N, Detmar M, Lagutina I, Grosveld G, Gunn MD, Jackson DG, & Oliver G. An essential role for Prox1 in the induction of the lymphatic endothelial cell phenotype. EMBO Journal 2002 21 1505–1513. (https://doi.org/10.1093/emboj/21.7.1505)
- 48↑
Bradley M. Brief report: systemic vascular access and resuscitation via corpus cavernosum. Military Medicine 2016 181 e1491–e1494. (https://doi.org/10.7205/MILMED-D-16-00001)
- 49↑
Nicol D, Watt A, Wood G, Wall D, & Miller B. Corpus cavernosum as an alternative means of intravenous access in the emergency setting. Australian and New Zealand Journal of Surgery 2000 70 511–514. (https://doi.org/10.1046/j.1440-1622.2000.01859.x)
- 50↑
Young MD, Mitchell TJ, Vieira Braga FA, Tran MGB, Stewart BJ, Ferdinand JR, Collord G, Botting RA, Popescu DM, Loudon KW, et al.Single-cell transcriptomes from human kidneys reveal the cellular identity of renal tumors. Science 2018 361 594–599. (https://doi.org/10.1126/science.aat1699)
- 51↑
Zenker M, Edouard T, Blair JC, & Cappa M. Noonan syndrome: improving recognition and diagnosis. Archives of Disease in Childhood 2022 107 1073–1078. (https://doi.org/10.1136/archdischild-2021-322858)
- 52↑
Northup KA, Witte MH, & Witte CL. Syndromic classification of hereditary lymphedema. Lymphology 2003 36 162–189.
- 53↑
Korpelainen EI, Karkkainen MJ, Tenhunen A, Lakso M, Rauvala H, Vierula M, Parvinen M, & Alitalo K. Overexpression of VEGF in testis and epididymis causes infertility in transgenic mice: evidence for nonendothelial targets for VEGF. Journal of Cell Biology 1998 143 1705–1712. (https://doi.org/10.1083/jcb.143.6.1705)
- 54↑
Simmons MN, & Jones JS. Male genital morphology and function: an evolutionary perspective. Journal of Urology 2007 177 1625–1631. (https://doi.org/10.1016/j.juro.2007.01.011)
- 55↑
Iguchi T, Irisawa S, Uesugi Y, Kusunoki S, & Takasugi N. Abnormal development of the os penis in male mice treated neonatally with tamoxifen. Acta Anatomica 1990 139 201–208. (https://doi.org/10.1159/000146998)
- 56↑
Kizhatil K, Ryan M, Marchant JK, Henrich S, & John SWM. Schlemm’s canal is a unique vessel with a combination of blood vascular and lymphatic phenotypes that forms by a novel developmental process. Hogan BLM, editor. PLoS Biology 2014 12 e1001912. (https://doi.org/10.1371/journal.pbio.1001912)
- 57↑
Perkumas KM, & Stamer WD. Protein markers and differentiation in culture for Schlemm’s canal endothelial cells. Experimental Eye Research 2012 96 82–87. (https://doi.org/10.1016/j.exer.2011.12.017)
- 58↑
Hamann S, Zeuthen T, Cour ML, Nagelhus EA, Ottersen OP, Agre P, & Nielsen S. Aquaporins in complex tissues: distribution of aquaporins 1–5 in human and rat eye. American Journal of Physiology 1998 274 C1332–C1345. (https://doi.org/10.1152/ajpcell.1998.274.5.C1332)
- 59↑
Herrnberger L, Ebner K, Junglas B, & Tamm ER. The role of plasmalemma vesicle-associated protein (PLVAP) in endothelial cells of Schlemm’s canal and ocular capillaries. Experimental Eye Research 2012 105 27–33. (https://doi.org/10.1016/j.exer.2012.09.011)
- 60↑
Hamanaka T, Bill A, Ichinohasama R, & Ishida T. Aspects of the development of Schlemm’s canal. Experimental Eye Research 1992 55 479–488. (https://doi.org/10.1016/0014-4835(9290121-8)
- 61↑
Fujimoto T, Inoue T, Maki K, Inoue-Mochita M, & Tanihara H. Vascular endothelial growth factor: a increases the aqueous humor outflow facility. Acott TS, editor. PLoS One 2016 11 e0161332. (https://doi.org/10.1371/journal.pone.0161332)
- 62↑
Nielsen S, Pallone T, Smith BL, Christensen EI, Agre P, & Maunsbach AB. Aquaporin-1 water channels in short and long loop descending thin limbs and in descending vasa recta in rat kidney. American Journal of Physiology 1995 268 F1023–F1037. (https://doi.org/10.1152/ajprenal.1995.268.6.F1023)
- 63↑
Pannabecker TL, & Dantzler WH. Three-dimensional architecture of inner medullary vasa recta. American Journal of Physiology. Renal Physiology 2006 290 F1355–F1366. (https://doi.org/10.1152/ajprenal.00481.2005)
- 64↑
Bulla R, Bulla R, Villa A, Bossi F, Cassetti A, Radillo O, Spessotto P, De Seta F, Guaschino S, & Tedesco F. VE-cadherin is a critical molecule for trophoblast-endothelial cell interaction in decidual spiral arteries. Experimental Cell Research 2004. (https://doi.org/10.1016/j.yexcr.2004.09.015)
- 65↑
Goldman-Wohl DS, Ariel I, Greenfield C, Lavy Y, & Yagel S. Tie-2 and angiopoietin-2 expression at the fetal-maternal interface: a receptor ligand model for vascular remodelling. Molecular Human Reproduction 2000 6 81–87. (https://doi.org/10.1093/molehr/6.1.81)
- 66↑
Tewari N, Kalkunte S, Murray DW, & Sharma S. The water channel aquaporin 1 is a novel molecular target of polychlorinated biphenyls for in utero anomalies. Journal of Biological Chemistry 2009 284 15224–15232. (https://doi.org/10.1074/jbc.M808892200)
- 67↑
Hirashima M, Lu Y, Byers L, & Rossant J. Trophoblast expression of fms-like tyrosine kinase 1 is not required for the establishment of the maternal–fetal interface in the mouse placenta. PNAS 2003 100 15637–15642. (https://doi.org/10.1073/pnas.2635424100)
