Abstract
Aging is a major risk factor for impaired cardiovascular health. The aging myocardium is characterized by electrophysiological dysfunctions such as a reduced heart rate variability. These alterations can be intrinsic within cardiomyocytes, but might be modulated by the cardiac autonomic nervous system, as well1. It is known that nerves align with vessels during development2, but the impact of aging on the cardiac neuro-vascular interface is unknown. Here, we report that aging reduces nerve density specifically in the left ventricle and dysregulates vascular-derived neuro-regulatory genes. Aging leads further to a down-regulation of miR-145 and de-repression of the neuro-repulsive factor Semaphorin-3A. miR-145 deletion increased Sema3a expression and reduced axon density, thus mimicking the observed aged heart phenotype. Removal of senescent cells, which accumulated with chronological age while nerve density declined, rescued from age-induced dennervation, reduced Sema3a expression and preserved heart rate variability. These data suggest that senescence-associated regulation of neuro-regulatory genes contributes to a declined nerve density of the aging heart and thereby to a reduced heart rate variability.
Main Text
The vasculature and nervous system form complex, highly branched networks, which are frequently interdependent and functionally linked. Vessel-nerve alignments are mediated by nerve-derived signals that act on endothelial cells or, conversely, the formation of nerve fibers along a preformed vessel template2. Thereby, guidance cues such as Semaphorins, Eph/ephrins and vascular endothelial growth factor (VEGF)/VEGFR regulate vessels and neurons, and extend or repel axonal growth3. Afferent and efferent cardiac neurotransmission via sympathetic and parasympathetic cardiac nerves modulates many physiological functions of the heart. Hence imbalances of either branches can lead to arrhythmias. For instance, impaired cardiac parasympathetic activity is a negative prognostic indicator and can lead to ventricular arrhythmia4,5, whereas both excessive6–8 and reduced9 sympathetic activity can lead to arrhythmias. Moreover, cardiac denervation lead to silent ischemia and lethal arrhythmia in diabetic hearts 5,9,10. A reduced heart rate variability is indicative of an impairment of both parasympathetic and sympathetic innervation in the elderly and has a negative prognostic value11. Beyond the control of electrical stability, innervation has additional functions and, for example, is essential for regeneration of the heart, as shown in postnatal post-infarction regeneration models in mice12,13. In the vasculature and in other organs innervation can control inflammation14,15. However, whether aging has an impact on cardiac innervation on a cellular and mechanistic level is unknown.
Here, we explored the impact of aging on nerve density in old mice. We used 18-20 month old male C57Bl/6J mice, which revealed diastolic dysfunction, while ejection fraction was preserved (Suppl. Fig. 1a, b). Pan-neuronal staining for Tuj1 showed a robust reduction of axon density in 18-month old mouse hearts compared to 12 week old young mice (Fig.1a). The age-induced reduction in nerve density was specifically detected in the left ventricle, while the right ventricular innervation was comparable between old and young mice (Fig.1b, c). The extend of age-dependent decline in the epicardial region was not as strong as in the subendocardial and myocardial regions, but high magnification images proofed a robust decline of sub-epicardial axon density in aged hearts as well (Fig.1b; Suppl. Fig. 2a). This observation was further confirmed in whole mount staining of old mouse hearts, which showed a decline in nervous fibers across the left posterior wall (Fig.1d). To assess the specific time-point when denervation starts, we performed a time course study showing a decline of nerve density already at 16 month of age with a further decline at 22 months (Fig.1e; Suppl. Fig. 2b).
Next, we assessed which types of nerves are affected by aging. The heart is innervated by sympathetic, parasympathetic and sensory fibers, which are commonly stained for tyrosine hydroxylase (TH)16,17, choline acetyltransferase (ChAT)18,19 and calcitonin gene-related peptide (CGRP)20, respectively. TH-positive nerves were present in both ventricles, but were selectively reduced in the left ventricle of aged mice (Fig.1f). ChAT-positive nerves were sparse in either ventricles and only occasionally detected in the cardiac base of young hearts (Fig.1g). Sensory neurons were exclusively detected in perivascular regions and were also significantly diminished by age (Fig.1h). Aging additionally resulted in a higher incidence of ventricular tachycardia and arrhythmias in Langendorff perfused hearts of 18 month old mice compared to young mice (Suppl. Fig. 3) documenting increased electrical instability. Together these data demonstrate a decline of cardiac innervation in the left ventricle of old aged mice.
To shed light on the functional consequences that arise from the age-associated cardiac denervation, we assessed heart rate variability by time domain and frequency domain analyses in awake mice using telemetric ECG tracing. In line with a recent report21, we observed a reduced variation of the RR-intervals (SDNN) in aged versus young mice (Fig. 1i). Especially, frequency-domain analyses as assessed by LF/HF ratio, which can be considered as an indicator for the sympatho-vagal balance22, was reduced with age, suggesting a reduced sympathetic activity in aged animals (Fig. 1j). The day-night-rhythm was also impaired with age (Fig. 1i, j). Taken together, aging leads to left ventricle-specific decrease in neuronal density that correspond to decreased heart rate variability and arrhythmias.
The vasculature and the nervous system co-develop and remain aligned also in the mature heart (Fig. 2a). To address if the decline in nerve density may be secondary to age-related capillary rarefaction, we histologically assessed capillary density over 22 months (Suppl. Fig. 4). However, capillary density only decreased at 22 months (Suppl. Fig. 4). At 16 months, when the initial decline of nerve density was observed, no difference in capillary density was detectable excluding that the reduction of nerves is secondary to the loss of capillaries. However, vascular alignment of nerves is lost in 18-month old mouse hearts (Fig. 2b, c), which might have been caused by a dysregulation of neuro-guidance cues in the vasculature.
To address, if neuro-guidance cues might be dysregulated in the cardiac endothelium of the aging heart, we isolated endothelial cells from young and old mice and performed RNA sequencing. GO term analysis of significantly induced genes demonstrated pathways assigned to “neuronal death” and “axon injury”, (Fig. 2d). Genes in these pathways include Semaphorin-3a (Sema3a), which patterns the autonomic nervous system during development23. Semaphorin-3A is further essential to maintain normal heart rhythm through sympathetic innervation patterning, but induces vulnerability to arrhythmias if overexpressed in cardiomyocytes6 (Fig. 2e). In addition, we found upregulation of members of the Slit/Robo family, such as Slit3, which can mediate repulsive signals24, and Netrin-1 (Ntn1), a laminin-related secreted protein, which may switch attraction to repulsive responses in a dose-dependent manner25 (Fig. 2e). Interestingly, recent studies suggest that the combination of guidance factors (such as Slit-family members and Netrin-1) can act in concert to modulate cellular responses25. Validation of protein expression confirmed the upregulation of Semaphorin-3A in aged mouse hearts (Fig. 2f) and further showed that Semaphorin-3A is predominantly expressed by vascular cells (Fig. 2g).
Since both overexpression and deletion of Sema3a may lead to sudden cardiac death and ventricular fibrillation7, we investigated up-stream pathways, which may control age-induced induction of Sema3a. Sema3a-mRNA has two miR-145 binding sites in the 3’UTR26 (Fig. 2h), and miR-145-5p was significantly reduced in the aging heart (Fig. 2i). Therefore, we hypothesize that miR-145 might repress Sema3a in the young heart. Overexpression of miR-145 indeed repressed Sema3a in human umbilical cord vein endothelial cells in vitro (Fig. 2j). Furthermore, miR-143/145-/- mice showed increased levels of Semaphorin-3a among vessels (Fig. 2k) and reduced axon density (Fig. 2l) even at young age (10-15 weeks). Together, these data suggest that loss of miR-145 induced de-repression of Sema3a is sufficient to reduce cardiac nerve density.
Importantly, SEMA3A was also up-regulated in senescent endothelial cells, which were generated by continuous passaging to induce replicative senescence as evidenced by acidic β-galactosidase staining (Fig. 3a, b). Interestingly, cellular senescence is induced concomitantly in the aging mouse hearts when neuronal density declines at 16 months (Fig. 3c-e). Moreover, genetic models of premature senescence such as 4th generation Tert-/- mice that lack telomerase confirmed a decline in nerve density (Fig.3f, g). By applying a senescent score27 to our previously published single nuclei RNA sequencing data of young vs. old mouse hearts28, we identified endothelial cells to acquire the most senescent phenotype (Suppl. Fig. 5a). Bulk RNA sequencing data of isolated cardiomyocytes29, fibroblasts28 and endothelial cells confirmed the up-regulation of senescence marker genes predominantly in aged cardiac endothelial cells (Suppl. Fig. 5b). This suggests that endothelial senescence might contribute to neuronal repulsion or death. Indeed, the selective induction of endothelial cell senescence in young animals by endothelial-specific overexpression of progerin30 significantly reduced the density of Tuj1 positive nerves compared to wildtype littermates (Fig. 3h). Taken together, different models of premature senescence indicate that the induction of (endothelial) senescence is sufficient to induce cardiac sympathetic denervation.
To determine if interfering with cellular senescence might prevent cardiac denervation in the aged heart, we treated old mice with 5 mg/kg dasatinib and 50 mg/kg quercetin, a combination of senolytics, which was shown to reduce the number of senescent cells by targeting anti-apoptotic pathways and expands life span in vivo31,32. Treatment was applied via oral gavage to aged mice (18 months) on three consecutive days, every second week for a total duration of two months (Fig. 4a). At 2 months after start of the treatment, the number of senescent acidic β-galactosidase-positive cells was significantly lower as compared to placebo-treated controls (Fig. 4b, c). Importantly, the reduction in senescent cells was paralleled by a rescue of Tuj1-positive nerves by senolytic treatment (Fig. 4d). Consistently, senolytic treatment augmented heart rate variability as assessed by the LF/HF ratio already 2 weeks after start of the treatment (Fig. 4e, f). Two months of senolytics treatment further improved the LF/HF ratio in aged mice, and restored the characteristic day-night-rhythm, while old control mice further deteriorated in the autonomous function (Fig. 4f; Suppl. Fig. 6a, b). These data indicate that senolytics induce a re-innervation of the aging heart, which restores the sympatho-vagal balance. In addition, senolytics treatment improved cardiac function as evidenced by a normalized diastolic function at 4 and 8 weeks of treatment (Suppl. Fig. 7a, b) and reduced vulnerability to arrhythmia as assessed by Langendorff-perfused hearts 8 weeks after senolytics treatment (Suppl. Fig. 8).
To provide mechanistic insights into how senolytics rescue cardiac innervation, we performed single nuclei RNA sequencing of old mouse hearts treated with senolytics or placebo (Fig. 4g). Interestingly, senolytic treatment affected genes associated with “nervous system development” within the top-regulated genes of cardiac endothelial cells (Fig. 4h). Importantly, Sema3a, which we showed to be de-repressed in the aging heart, was significantly reduced in old heart endothelial cells after senolytic treatment (Fig. 4i).
Here, we demonstrate that aging reduces axon density in the heart. Aging induced decline in axon density was associated with reduced miR-145 levels and de-repression of its target, the neuronal repulsive signal Semaphorin-3A, which is well known to induce electrical instability in the heart. Interestingly, induction of cellular senescence, which is a hallmark of aging, was inversely correlated with the onset of axon decline. Targeting senescent cells pharmacologically was sufficient to prevent the decline in axon density and reduced Sema3a expression in the aging heart suggesting a key role of senescent cells in cardiac denervation. Senescent cells release numerous secreted factors, termed senescence-associated secretory phenotype (SASP), which profoundly alters the microenvironment in the aging heart. Although neuronal guidance factors have not been reported as general SASPs, Semaphorin-3A is induced in senescent endothelial cells and may represent a specific vascular SASP in the aging heart.
The question which cell type(s) further contributes to the observed effects will need further studies. We demonstrate that the selective induction of pre-mature aging in endothelial cells is sufficient to reduce nerve density. However, we cannot exclude the involvement of other cells in this process. Interestingly, there was a tendency that Sema3a was also de-repressed by senolytic treatment in other vascular cells, namely pericytes, whereas e.g. fibroblasts showed very low levels and no Sema3a regulation (Suppl. Fig. 9). Moreover, neuronal and axonal related pathways were found within the top-25-regulated GO terms in lymphatic endothelial cell and in some fibroblast clusters of mice treated with senolytics (Suppl. Tab. 1). These findings indicate that endothelial cells may play a critical role in age-related denervation, but other cells such as pericytes or fibroblasts may contribute as well to the observed phenotype.
Our study additionally demonstrates that senolytic treatment restores vulnerability to arrhythmia, heart rate variability and the circadian rhythm. A decline in heart rate variability is typically observed in the elderly, is indicative of impaired sympathetic and parasympathetic innervation and is associated with increased electrical instability leading to increased overall mortality11. Our finding that senolytics normalizes heart rate variability during aging, thus, supports a functional benefit of the treatment.
Innervation is not only important for the control of heart rhythm but nerves were shown to provide important paracrine factors, which for example contribute to cardiac regeneration12,13. A decline in nerve density may consequently lead to depletion of such nerve-derived factors influencing the reparative function of the heart as it was demonstrated for myocardial infarction in adult mice33. Moreover, in other tissues, nerves interact with immune cells34 and can control vascular inflammation14. Since inflammation is a hallmark of aging (“inflamaging”), the relation of neuro-immune interactions in the heart may deserve further studies. Together, the presented findings may lay the ground to decipher neuronal cross-talks in the heart and their role in aging.
Methods
Laboratory animals
Isogenic male C57Bl/6J wildtype mice were purchased from Janvier (Le Genest SaintIsle, France) and from Charles River (Sulzfeld, Germany). Homozygosity of these inbred mice was controlled by Janvier and Charles River using exome sequencing.
miR143/145 gene cluster knockout mice were generated as previously described35. Male and female miR143/145 gene cluster knockout mice with an C57Bl/6J background and an age between 10 to 15 weeks were used.
Pre-mature senescence was studied in male Tert-knockout mice (4th generation, 10 to 15 weeks old) with an C57Bl/6J background as previously described36,37.
Endothelial-specific progeria mice were generated as previously described30. Male and female mice were use at the age of 28 to 29 weeks.
To obtain hearts, mice were sacrificed via cervical dislocation during isoflurane anesthesia and perfused with cold Hank’s buffered saline solution (HBSS; 14175-053, Invitrogen).
Mice were housed in individually ventilated cages in a specific pathogen-free facility according to national and institutional guidelines for animal care.
Senolytic treatment
To eliminate senescent cells from aged mice, a combination of the two senolytics drugs dasatinib and quercetin was used as proposed by Xu et al.38. In brief, 5 mg/kg dasatinib (SML2589-50MG; Merck) and 50 mg/kg quercetin (Q4951-10G, Sigma-Aldrich) were applied via oral gavage to aged mice (18 months) on three consecutive days, every second week for a total duration of two months. Young (12-16 weeks) and aged (18-19 months) mice receiving the solvent (Phosal 50PG (368315, Lipoid) containing 3.3% ethanol (32221, Sigma-Aldrich) and 10% polyethylene glycol 400 (807485, Merck)) served as control cohorts. Cardiac function was monitored during the experiment using echocardiography and ECG traces as described below. The animal experiment has been conducted as approved by the state of Hessen (animal application number FU/1269).
Echocardiography
To assess heart function via echocardiography, mice were anaesthetized (2-2.5% isoflurane) and monitored using the Vevo 3100 echocardiography system with the Vevo LAB software (Fujifilm VisualSonics).
Telemetric ECG measurement
To record long-term ECG traces remotely in awake mice, ETA-F10 transmitters (270-0160-002, DSI) were implanted subcutaneously as described by the provider’s instruction. In brief, buprenorphine (0.1 mg/kg) was injected i.p. to mice 30 minutes before starting the surgery. Then mice were anaesthetized (1.5% isoflurane) and an incision was made on the left anatomical side of the mouse. The transmitter was covered with polymyxin and placed subcutaneously. The electrodes were stitched to the pectoral muscles in Einthoven II position. The wound was closed and mice received metamizol on three consecutive after surgery as post-surgical treatment. The animal experiment has been conducted as approved by the state of Hessen (animal application number FU/1269). ECG traces were recorded and time and frequency domain were analyzed using the software Ponemah 6.
Assessment of ventricular arrhythmia inducibility
After anaesthesia by isoflurane and cervical dislocation hearts were rapidly excised by opening the thorax and immediately placed in ice-cold buffer solution (modified Krebs-Henseleit solution; mM: NaCl 119, NaHCO3 25, KCL 4.6, KH2PO4 1.2, MgSO4 1.1, CaCl2 2.5, C6H12O68.3 and Na-Pyruvate 2; pH 7.4)5. The ascending aorta was pulled over a cannula, hearts were transferred into a Langendorff apparatus and the cannula was rapidly attached to keep no flow time as short as possible. Hearts were electro-mechanical uncoupled by blebbistatin added to the perfusion buffer (5 – 10 µM, Hoelzel Biotech). Perfusion pressure and heart rate were continuously monitored (Powerlab 8/30 & Labchart, ADInstruments). Perfusion flow was manually regulated based on the perfusion pressure (80 – 100 mmHg) using a peristaltic pump (Regalo Masterflex Masterflex, Ismatec)39. An octopolar electrophysiology catheter (2.0 F, 0.5 mm electrode spacing; CIBer Mouse, NuMed) was placed in the right ventricle to stimulate the heart and to continuously obtain atrial and ventricular electrograms5,40. For equilibration the heart was paced at 600 bpm for 30 minutes. Perfusion buffer was continuously oxygenated using carbogen (95% O2/5% CO2). Hearts which presented relevant arrhythmias or visible ischemia after equilibration were excluded. To assess susceptibility to ventricular arrhythmias (VA) we used a stimulation protocol based on three maneuvers: (1) Programmed extrastimulation: train of eight S1-stimuli (cycle length (CL) 100 ms) followed by two or three extrastimuli with a decremental S2S3- or S3S4-interval with a stepwise (2 ms) reduction (60 – 20 ms). (2) Miniburststimulation: train of 20 S1-stimuli (CL 100 ms) followed by ten S2-stimuli with a decremental S2-interval with a stepwise (2 ms) reduction (60 – 20 ms). (3) Burststimulation: train of 20 – 100 S1-stimuli with a decremental S1-interval (50 – 10 ms). VAs were classified using an established scoring system40.
Single-nucleus RNA sequencing
To assess the cardiac transcriptome on single nuclear level, nuclei isolation from mouse hearts, single-nuclei separation, library preparation and sequencing were performed as previously described41.
Single-nucleus RNA sequencing data analyses
To analyze single-nucleus RNA sequencing mouse data of senolytic and the control treated mice, the samples were mapped to the mice reference genome (GRCm38) via STARsolo (version 2.7.9) with the parameter “-- soloFeatures GeneFull”. Data ingtegration, normalisation, scaling and UMAP clustering were performed with Seurat (version 4.1.1), according to the developer’s tutorial (https://satijalab.org/seurat/articles/pbmc3k_tutorial.html). After filtering of nuclei based on mitochondrial content (<5%) and genes per nucleus (<2500) a total 13541 single nuclei were analyzed from 6 different samples.
Differential expressed genes were tested using the FindAllMarkers function with the statistical test bimod in the Seurat package. Genes with adjusted p-values <0.05 were considered as differential expressed genes.
Whole mount immunofluorescence staining
After sacrificing mice, hearts were perfused with 4% PFA (28908, ThermoFisher Scientific) in PBS, harvested and incubated in 4% PFA for 4h at 4°C. Hearts were washed trice with PBS for 5 minutes and bleached overnight at room temperature using DMSO (A994.2, Carl Roth GmbH & Co. KG) and H2O2 (8070.2, Carl Roth GmbH & Co. KG) diluted 1:1:4 (vol/vol/vol) in PBS. Hearts were washed trice with PBS for 20 minutes each followed by antigen retrieval by incubating whole hearts in retrieval buffer (4% SDS (CN30.3, Carl Roth GmbH & Co. KG) and 200 mM boric acid (191411, MP Biomedicals)) for 1h at room temperature followed by overnight incubation at 54°C. Hearts were again washed trice in PBT (0.2% Triton X-100 in PBS) for 1h each and incubated in blocking solution (10% FBS (4133, Invitrogen), 1% BSA (A7030-10G, Merck), 5% donkey serum (017-000-121, Jackson Immuno) in PBT) for 1h at room temperature. Rabbit anti-Tuj1 antibody (ab18207, Abcam) was diluted 1:100 in blocking solution and incubated with the hearts for 3 days at room temperature. Hearts were then washed trice in PBS for 20 minutes and incubated with the secondary antibody (donkey anti-rabbit antibody conjugated to Alexa 555; A-31572, Invitrogen) that was diluted 1:100 for at least 2 days at room temperature. Hearts were again washed trice for 20 minutes in PBS and embedded in agarose (9012-36-6, Carl Roth GmbH & Co. KG). Hearts were dehydrated at room temperature in an ascending methanol series (30%, 50%, 75%, 30 minutes each) and incubated twice in 100% methanol at room temperature for 30 minutes each. Hearts were washed twice in ECI (112372, Sigma-Aldrich) for 5 minutes and cleared by incubating in 80% ECI and 20% PEGM (447943, Sigma-Aldrich) for 30 minutes at room temperature.
Whole hearts were assessed histologically using a light sheet microscope (Ultramicroscope II, LaVision BioTec, Bielefeld, Germany). Excitation was performed at 470/40 nm and emission 525/50 nm (autofluorescence tissue), excitation 545/30 nm and emission 595/40 nm(Tuj). Main laser power 95% and software laser power for 470/40 95% and 525/50 35%. Step size was set to 5 µm. Exposure time was 300 ms, 6,3× magnification (10× zoom body + 0.63× Objective). Sheet width 60%; Sheet NA 4,05um; two sided scan. Pictures were taken with a Neo 5.5 (3-tap) sCOMs Camera (Andor, Mod. No.: DC-152q-C00-FI). Images were analyzed using the Imaris software, version 9.
Immunofluorescence staining of cryopreserved heart sections
After sacrificing mice, hearts were flushed with cold HBSS and fixed in PBS containing 4% PFA (28908, ThermoFisher Scientific). After overnight incubation at 4°C, hearts were washed three times for 10 minutes in PBS. To cryopreserve cardiac tissues, three consecutive overnight washes in PBS containing increasing concentrations of sucrose (10%, 20%, 30%; S0389, Sigma-Aldrich) were applied at 4°C. Tissues were embedded in PBS containing 15% sucrose, 8% gelatin (G1890, Sigma-Aldrich), and 1% polyvinylpyrrolidone (P5288, Sigma-Aldrich). After the embedding solution was solidified, tissues were stored at −80°C. Hearts were sectioned at 50 µm-thickness using a cryostate (Leica CM3050 S). Sections were placed on adhesive glass slides (10149870, ThermoFisher Scientific) and stored at −20°C until use.
For immunofluorescence staining, cryo sections were brought to room temperature and re-hydrated in PBS (twice for 5 minutes). To permeabilize the tissue, sections incubated with PBS containing 0.3% Triton X-100 three times for 10 minutes and were blocked in PBS containing 0.1% Triton X-100, 3% BSA (A7030-10G, Merck) and 5% donkey serum (ab7475, Abcam) for 1h at room temperature. Primary antibodies were diluted in blocking solution and incubated with the sections overnight at 4°C. Sections were then washed three times for 5 minutes in PBS and incubated for 1h at room temperature with the respective secondary donkey antibodies that were diluted in PBS containing 0.1% Triton X-100. Nuclei were stained with DAPI (6335.1, Carl Roth GmbH & Co. KG) that was diluted 1:1000 in 0.1% Triton X-100. After washing trice in PBS for 5 minutes, slides were mounted with Fluoromount-G™ (00-4958-02, Invitrogen). Sections were histologically assessed using the Leica Stellaris confocal microscope and the LASX software.
Immunofluorescence staining of paraffin heart sections
To assess hearts histologically on paraffin sections, hearts were processed and embedded as previously described28. To immunolabel paraffin section, slides incubated for 1h at 60°C and were deparaffinized twice with xylene for 10 minutes and an ethanol series of 100%, 95%, 80%, 70%, and 50% ethanol (5 minutes each step). Sections were washed in water for 5 minutes and were boiled in 0.01 M citrate buffer (pH = 6) for 90 seconds. Slides were then washed for 5 minutes with PBS and blocked in PBS containing 0.1% Triton X-100, 3% BSA (A7030-10G, Merck) and 5% donkey serum (ab7475, Abcam) for 1h at room temperature. Primary antibodies were diluted in blocking solution and incubated with the sections overnight at 4°C. Sections were then washed three times for 5 minutes in PBS and incubated for 1h at room temperature with the respective secondary donkey antibodies that were diluted in PBS containing 0.1% Triton X-100. Nuclei were stained with DAPI (6335.1, Carl Roth GmbH & Co. KG) that was diluted 1:1000 in 0.1% Triton X-100. After washing trice in PBS for 5 minutes, slides were mounted with Fluoromount-G™ (00-4958-02, Invitrogen). Sections were histologically assessed using the Leica Stellaris confocal microscope and the LASX software.
Antibodies
Following primary antibodies have been uses:
Rb anti-Tuj1 (1:100, ab18207, Abcam), Rb anti-tyroxine hydroxylase (1:100, AB152, Merck), Gt anti-Choline Acetyltransferase (1:100, AB144P, Merck), Gt anti-calcitonin gene-related peptide (1:100, ab36001, Abcam), Ms anti-α-Smooth Muscle - Cy3™ (1:200, C6198-2ML, Sigma-Aldrich), Rb anti-Semaphorin-3A (1:100, ab23393, Abcam) and GSL I - isolectin B4 (biotinylated; 1:25, VEC-B-1205, Biozol).
Following secondary antibodies have been used:
Donkey anti-mouse IgG Alexa Fluor 647 (1:200, A-31571, Invitrogen), Donkey anti-rabbit IgG Alexa Fluor 555 (1:200, A-31572, Invitrogen), Donkey anti-rabbit IgG Alexa Fluor 488 (1:200, A-21206, Invitrogen), Donkey anti-Goat IgG Alexa Fluor 555 (1:200, A-21432, Invitrogen), Donkey anti-Goat IgG Alexa Fluor 647 (1:200, A-21447, Invitrogen), Streptavidin, Alexa Fluor™ 405 (1:200, S32351, Invitrogen) and Streptavidin, Alexa Fluor™ 647 (1:200, S32357, Invitrogen).
Quantification of immunofluorescence images
To analyze and quantify immunofluorescence images, the stained area was determined and normalized to IB4- or DAPI-positive area using the software Volocity 7 by Quorum Technologies Inc.
Acidic beta-galactosidase staining
Acidic beta-galactosidase positive cells were visualized on cryopreserved heart sections and in vitro using the Senescence β-Galactosidase Staining kit (9860, CST) according to the manufacturer’s instruction. β-galactosidase-positive areas were quantified using ImageJ.
Endothelial cell isolation from murine hearts
Cardiac endothelial cells were isolated from young (12 weeks) and old (20 months) mice. Under isoflurane anesthesia, mice were sacrificed and hearts were flushed with HBSS. The hearts were harvested, dissected in small pieces, transferred into a C-tube (130-096-334, Miltenyi Biotec) and incubated in HBSS, containing 600 U/mL collagenase type II (354236, Corning), at 37°C and 5% CO2 in a humidified atmosphere. After 30, 20 and 10 minutes of incubation, tissue particles were further dissected using the GentleMACS Dissociator (Miltenyi BioTec) with the pre-set program m_neoheart_01_01. Collagenase digestion was stopped with 500 µL fetal bovine serum (4133, Invitrogen). Cell suspension was applied on a 200 µm cell strainer (43-50200-03, pluri-Select), centrifuged at 80x g and 4°C for 1 minute to deplete cardiomocytes and applied on a 70 µm cell strainer (43-50070-03, pluri-Select). Cells were washed twice with HBSS containing 0.5% bovine serum albumin (T844.3, Carl Roth GmbH & Co. KG) and 2 mM EDTA (A4892, AppliChem) (referred to as wash buffer in the following) by centrifugation (300x g at 4°C for 10 minutes). Endothelial cells were isolated using rat anti-mouse CD144 antibodies (555289, BD Bioscience) and magnetic sheep anti-rat dynabeads (11035, Life Technologies). During tissue dissection, anti-CD144 antibody-bead mixture was prepared by washing 25 µL dynabeads twice with wash buffer and re-suspending them in 400 µL wash buffer. 1 µL of antibodies were added to the beads, incubated for 1h at room temperature and were washed trice with wash buffer. Antibody-bead mixture was re-suspended in 1000 µL wash buffer, added to the cardiac cell pellet and incubated for 40 minutes on a turning wheel. Cells were washed trice on a magnetic rack using 1000 µL of wash buffer and lysed with 700 µL of Qiazol.
RNA isolation
Total RNA was purified from cultured and isolated cells by using the miRNeasy Mini kits (217004, Qiagen), combined with on-column DNase digestion (DNase Set, 79254, Qiagen) as described in the manufacturer’s instruction. To isolate RNA from solid hearts, tissue was combined with 700 µL Qiazol and ¼” ceramic spheres and were homogenized three times for 20 seconds (4 m/s). RNA isolation was then performed using the miRNeasy Mini kit (217004, Qiagen). The RNA concentration was determined by measuring absorption at 260 nm and 280 nm with the NanoDrop ND 2000-spectrophotometer (PeqLab).
cDNA synthesis and quantitative PCR
To quantify mRNA expression by qPCR, 100 ng to 1 μg of total RNA was reverse-transcribed using the reverse transcriptase M-MLV (28025013, ThermoFisher Scientific) and assessed using the SYBR™ Green PCR Master Mix (4385617, Applied Biosystems) as previously described28. The primers were customized and purchased from Sigma-Aldrich (now Merck): human Rpl0 fw (TCGACAATGGCAGCATCTAC); human Rpl0 rev (ATCCGTCTCCACAGACAAGG); human SEMA3A fw (TGTTGGGACCGTTCTTAAAGTAGT); human SEMA3A rev (TAGTTGTTGCTGCTTAGTGGAAAG).
Bulk RNA sequencing
Library preparation and whole transcriptome analysis of isolated cardiac endothelial cells were performed as previously described28.
Gene ontology term analysis
Gene ontology (GO-term) analyses were performed using the Enrichr online platform (ontology category GO Biological Process 2021; https://maayanlab.cloud/Enrichr/) by assessing significantly regulated genes of logFC > 1 and logFC < −1.
Micro Array
To assess micro-RNA expression in whole young and old mouse hearts, a published data set was used42.
Cell Culture
Human umbilical cord vein endothelial cells (HUVEC, CC-2935) were purchased from Lonza and cultured with endothelial basal medium (EBM, CC-3156, Lonza) supplemented with 10 % FBS (4133, Invitrogen), Amphotericin-B (CC-4081C, Lonza), ascorbic acid (CC-4116C, Lonza), bovine brain extract (CC-4092C, Lonza), endothelial growth factor (CC-4017C, Lonza), gentamycin sulfate (CC-4081C, Lonza), and hydrocortisone (CC-4035C, Lonza) at 37°C and 5% CO2, with humidified atmosphere. Short passage HUVEC (P2, P3) were used for in vitro studies. To resemble cellular senescence, HUVEC were cultured until at least passage 13 and were controlled by acidic beta-galactosidase staining.
Transfection experiments
70,000 HUVEC were seeded per well of a 12-well plate (665180, Greiner Bio-One GmbH) and rested for one day at 37°C and 5% CO2. Cells were transfected using the Lipofectamine RNAiMax (13778150, Invitrogen) according to the manufacturer’s protocol. Predesigned miR-145-5p precursors (4464066 (ID: MC11480), Ambion by Life Technologies) were used in a final concentration of 10 nM. Cells were cultured for 48h after transfection.
Statistical Analysis
Data are expressed as mean and error bars indicate the standard error of the mean (SEM). Normality distribution was assessed by using the Kolmogorov-Smirnov or Shapiro-Wilk normality test. For comparing two groups of Gaussian distributed data, statistical power was determined using the unpaired, two-sided student’s t-test. To compare more than two groups, an ordinary one-way ANOVA with a post-hoc Tukey comparison was used.
Data availability
All data are available within the article, within the supplemental material or from the corresponding author on reasonable request. Transcriptomic data are available from the cited publications or at the GEO (accession code will be provided as soon as the manuscript is accepted for publication).
Author contributions
JUGW and SD planned the project and wrote the manuscript. JUGW, LMK, JP conducted the majority of experiments. SG performed bulk RNA sequencing. LST and DJ performed bioinformatics. WTA performed snRNA sequencing. KAS and MC contributed to transcriptomic and histological analyses. AF, PFM and RPB contributed to telemetry studies. KS, GKB, MMR and GL contributed to histology and whole mount staining. SC, DS, SA, EA, NK, MK and CM contributed to electrophysiology and telemetry studies. GL, CM and AMZ provided conceptual input. TBo and TBr provided miR145 mice. CB provided Tert mice. EN and SOM provided Prog-tg mice.
Conflict of interest
The authors acknowledge grant support as listed, but otherwise do not have a conflict of interest
Acknowledgement
The study is supported by the German Research Foundation (SFB1366, Project B4; TRR267, Project B3 and the Cluster of Excellence Cardiopulmonary Institute Exc2026/1), the German Center for Cardiovascular Research (DZHK Shared Expertise (B22-014 SE)), the Dr. Rolf-M.- Schwiete Foundation (2021-002) and the European Research Council.