Detailed methods for cell culture are described in the Supplementary Methods section. Animal models–All mouse procedures are conformed to the Guide for Care and Use of Laboratory Animals of the US National Institutes of Health. The study respected European and French legislation on good practices in animal experimentation and was approved by the Animal Subjects Committee of Champagne Ardenne (CEEA-RCA-56). Male db/db mice (C57BL6/J as genetic background) are 6 months old in order to test the treatments on a severe diabetic model presenting significant cardiovascular alterations in accordance with the literature (Daniels Gatward et al., 2021; Gomes et al., 2021; Gomes et al., 2022). Metabolic parameters (food and water intakes, HbA1c, Glycemia, and body weight) were described in Table 1. We used 6-month-old non-diabetic control mice (C56BL6/J, n = 10) as a reference group. Mice were purchased from Janvier (Le Genest-Saint-Isle, France) All mice had ad libitum access to a standard diet (AIN-93 M rodent diet, Special Diet Service, United Kingdom) and water during the experimental period. The drinking water (renewed daily) of db/db mice contained (or not) minoxidil (20 mg/kg/day) or nebivolol (20 mg/kg/day) for 8 weeks. The concentrations and treatment times were adapted to previous data from the literature using minoxidil as a therapeutic molecule for ECM alterations.

Blood pressure assay–Preconditioning was performed within 7 days before the final measurement of blood pressure. For this purpose, the animals were acclimated to contention and to the pressure of the sphygmomanometer on their tails. The day of the final measure, five “empty” measurements were carried out within a 60 s interval. The final measurement was the average of five successive measurements. If the systolic and/or diastolic pressures of the mice (aged C57Bl/6 or treated or untreated young db/db) were statistically superior to the pressure of the control mice, we considered the mice to be hypertensive. Pulse wave velocity–Pulse wave velocity measurement was performed non-invasively using the ultrasound Doppler Flow Velocity System (Indus Instruments, Webster, United States) on anesthetized mice. The heart rate and ejection time between the femoral aorta and the aortic arch were measured, making possible to determine the speed of the pulse wave (pulse wave velocity) expressed in m/s. The latter was calculated by dividing the distance between the two Doppler probes by the ejection time between the two regions of the aorta. High frequency ultrasound analysis was performed using Vevo imaging systems (Fujifilm VisualSonics, Toronto, Canada) and under light isoflurane anesthesia (1.5%–2% in oxygen), with acquisition completed within a maximum of 10 min per animal, in accordance with ethical guidelines and to minimize cardiovascular depression in db/db mice. Heart rate was monitored to remain within the recommended physiological range (400–600 bpm), and animals were maintained on a heated platform to prevent hypothermia. As isoflurane can affect systolic parameters by reducing preload and contractility, we standardized all procedures to limit variability across animals and ensure comparable conditions across groups. At the level of the aortic arch, the diameter of the vessel was measured during the cardiac cycle (systole, Ds-diastole, Dd), as well as the pulse wave velocity (PWV) and the thickness of the tunica intima-media (h). Distensibility factor (DC) and Young’s modulus (E) were derived by Bramwell, (1922) equation and Moens-Korteweg equation, respectively. From the conclusions of Brands et al. (1999), the local variation of the pressure during the cardiac cycle (DP) and the compliance (CC) can then be deduced (Vanalderwiert et al., 2024a).

Detailed methods for histology and Immunofluorescent staining are described in the Supplementary Methods section. Atomic force microscopy (AFM) – Frozen 10 μm-thick aorta cross-sections were thawed in Krebs–Henseleit solution. After locating the zones of interest, analysis was performed using AFM (Bioscope Catalyst, Bruker, Billerica, United States, driven by Nanoscope Analysis 1.8 software) coupled to a Nikon Eclipse Ti inverted microscope (Nikon, Tokyo, Japan). The young modulus at each point of the EFs or of the interfiber spaces was calculated using a value of the Poisson ratio of 0.5 for our samples considered incompressible. For each condition, at least 5,000 force curves were treated to obtain the mean YM values for the EFs and the interfiber spaces. The analyses were performed at three different locations in each cross-section, for a total of nine cross-sections obtained from three different mice.

Detailed methods for immunoblotting, qPCR are described in the Supplementary Methods section. Extracellular elastin quantification was performed by ELISA test. After 48 h of incubation (with potassium channel modulators), MOVAS cell cultures (no impermeabilized) were incubated at 37 °C (2 h) with anti-Elastin antibody (BA41/1000, Sigma), then secondary antibody coupled to HRP (1 h, 37 °C). 3,3′,5,5′-tetramethybenzidin (TMB) was added in each well (5 min of incubation), followed by sulfuric acid (0.3M, for 10 min) to stop the reaction. Absorbance was read at 450 nM. Plasma marker assays - Peripheral blood of mice was collected in heparinized tubes by retroorbital puncture, and plasma were stored at −80 °C. Evaluations of desmosine and elastin-derived peptides (EDP) concentrations were performed using commercially available kits (Cusabio Biotech Product, Houston TX, United States and Biocolor, County Antrim, United Kingdom, respectively). The NE and cathepsin activity assay commercialized by Abcam (Cambridge, United Kingdom) were used to measure NE and cathepsin S activities, respectively, according to the method described by Romier et al. (Sarafidis et al., 2017). Tissular total collagen quantity was determined using a QuickZyme Total collagen kit (QuickZyme Biosciences, Leiden, the Netherlands). Crosslink and elastin quantification- Protein analysis was performed as described by Romier et al. (Olawi et al., 2019). Briefly, for collagen crosslink analysis samples were reduced by sodium borohydride (Sigma-Aldrich, Germany) (25 mg NaBH4/ml in 0.05 M NaH2PO4/0.15 M NaCl pH 7.4, 1 h on ice, 1.5 h at room temperature) and digested with high purity bacterial collagenase (C0773; Sigma, Germany; 50 U/mL, 37 °C, 12 h). The soluble fractions containing collagen cross-links were hydrolyzed in 6 N HCl at 110 °C for 24 h. The hydrolysates were precleared by solid phase extraction. Dried eluates were analyzed with an amino acid analyzer (Biochrom 30, Biochrom, Great Britain). The nomenclature of the crosslinks used in the article refers to the reduced variants of crosslinks. The collagen content was analyzed in an aliquot of hydrolyzed samples of the collagenase soluble fraction prior to preclearence and calculated based on a content of 14 mg hydroxyproline in 100 mg collagen. For protein and elastin crosslinks analysis, samples were digested with bacterial collagenase. The soluble fraction containing collagen was subjected to hydrolysis and amino acid analysis. The residual fraction was extracted by hot alkali (0.1 N NaOH, 95 °C, 45 min). The supernatant containing non-collagenous/non-elastin proteins and the insoluble residue containing insoluble elastin were subjected to hydrolysis and amino acid analysis. The content of elastin crosslinks was analyzed in an aliquot of the NaOH-insoluble fraction containing elastin after CF-11 preclearence by amino acid analysis.

Data were prospectively collected and analyzed using StatView 5.0 software for Windows (SAS Institute, Berkley, CA). Results are presented as either boxplots (with minimum and maximum values) or bar graphs showing mean ± SEM, to reflect the precision of the estimated means in a multi-replicate experimental design. Group comparisons were performed using non-parametric ANOVA (Kruskal–Wallis test), followed by pairwise Wilcoxon–Mann–Whitney tests with Bonferroni–Dunn correction for multiple comparisons. All reported p-values, including those displayed in figures and tables, are adjusted (corrected) p-values. These are explicitly indicated in the figure legends. Each analysis was based on at least three independent experiments, each including a minimum of five animals per group. A corrected p-value <0.05 was considered statistically significant. In all figures and tables, “*” denotes a corrected p-value <0.05 between untreated db/db mice and those treated with minoxidil or nebivolol, while “c” denotes a corrected p-value <0.05 between untreated db/db mice and C57Bl6 control mice.

In order to evaluate its efficiency in limiting the premature aging of the aorta wall observed in db/db mice, we administrated minoxidil to diabetic mice for 8 weeks in conditions comparable to those used to stimulate neosynthesis/protection of arterial EFs in young rats and aged mice (Coquand-Gandit et al., 2017; Fhayli et al., 2019).

Minoxidil-reduced systolic and pulse blood pressures (not mean arterial pressure (MAP)), as well as aortic pulse wave velocity or compliance and distensibility measured using the high ultrasound method, were indicative of arterial stiffness (Figures 2A–C, respectively). This was associated with a decrease in expression of the smooth muscle cells (SMC) contraction markers α-SMA, SM-22α, MLCK (Myosin light-chain kinase), Calponin and MYH11 (smooth muscle myosin heavy chain) (Figures 2D,E; Supplementary Figure 1B). These data demonstrated the antihypertensive and anti-aortic stiffening efficacy of minoxidil in diabetic animals but did not affect glycemic parameters (Table 1). Figure 2C suggests that minoxidil administration might be sufficient to restore an almost normal aortic function (observed by the dashed red line obtained with C57Bl6 mice (n = 10)). Minoxidil also induced aorta wall remodeling, leading to decreased intima-media and adventitia thicknesses (Figure 3A and Supplementary Figure 1B). This is possibly due to a decrease in total wall collagen content, particularly type III and I collagen as we observed by red picrosirius staining (Figure 3B). The increase of the mRNA expression of these collagens (Figure 3C) together with the absence of increased total collagen (Figure 3D) contents and crosslinks (Figure 3E) suggest the degradation of neosynthesized collagens. Because chronic inflammation, through the production of cytokines such as TNF-α and IL-6, can promote tissue fibrosis, we sought to determine whether minoxidil treatment affects inflammatory factors that could explain changes in collagen expression. We found that minoxidil significantly reduces the production of pro-fibrotic cytokines both at the plasma level (Figure 3F) and in the aorta of db/db mice (Figure 3G).

Elastin autofluorescence level (Figure 4A) and elastin quantity (Figure 4B; Supplementary Figure 2A) are increased, suggesting an increase in elastic fiber content and/or a decreased elastic fiber degradation after minoxidil treatment, as shown by elastin autofluorescence (Figure 4A) and Hart’s staining (Figure 4C). Indeed, while the number of elastic lamellae remained unchanged by minoxidil treatment (Figure 4D). Likewise, the number of elastic lamellae ruptures (Figure 4E) is decreased by the treatment reflecting a restructuring of lamellae. This reduced fragmentation of the elastic networks is confirmed by a significant decrease in plasmatic elastin-derived peptides (EDP) and desmosine levels (Figure 4F). Regarding elastolysis, the activities of cathepsin S and NE were reduced by the treatment (Figure 4G), while the tissue proteinase mRNA levels were unaffected by minoxidil (Figure 4H). The minoxidil-induced decrease in elastolysis in the aorta was amplified by the stimulatory effect of the treatment on the increased tissular expression of natural proteinase inhibitor, such as Serine peptidase inhibitor (SERPIN) (Figure 4I). The fact that minoxidil treatment stimulated aortic elastogenesis is further suggested by elevated mRNAs levels for elastin, LTBP4, and LOXL1 (Figure 4J). Increased and efficient elastogenesis was demonstrated by the increase of cross-links following the treatment (Figure 4K), as the occurrence of these cross-links is an indicator of the maturity and functionality of EFs. Inhibition of elastolysis and induction of elastogenesis by minoxidil improved elastic fiber content and function, as indicated by the decrease of elastic lamellae and interlamellar space stiffnesses in diabetic mice (Figure 4L). Taken together, these observations show that minoxidil limits premature aging of the aortic wall in diabetic mice and partially restores aorta function. Nevertheless, the cardiac parameter data (Figure 2F) show that, following the administration of minoxidil, cardiac stroke volume and left ventricle end-diastolic volume increased in db/db mice. The data highlights the fact that problems associated with alterations in left ventricular (LV) function can eventually impact aortic function and structure over time. This could compromise the anti-vascular aging effects of the medications used in our study and others (Coquand-Gandit et al., 2017; Knutsen et al., 2018; Knutsen et al., 2022).

Minoxidil chronic treatment can compensate for elastolysis induced by diabetes. However, this treatment has several side effects on cardiac functions, which limit its use (see Figure 2F) (Hanton et al., 2008). For this reason, we sought to test other antihypertensive molecules that could have similar beneficial effects without these harmful side effects. Among antihypertensive molecules, pharmacological blockade of the β1-adrenergic receptor can extend the life span of mice and flies, independently of body weight or metabolic syndrome (Spindler et al., 2013). Therefore, we chose to evaluate the effect of nebivolol, which has been described as a β-blocker reducing heart rate (Figure 5A), and a potent vasodilator activating β2 adrenergic receptors in endothelial cells and SMCs (Olawi et al., 2019; Toblli et al., 2010). Figure 5B shows that treatment with nebivolol for 8 weeks is sufficient to reduce systolic, mean, and pulse arterial pressures. We also observe nebivolol-induced decreases in aortic pulse wave velocity (Figure 5C), which favors the compliance of thoracic aorta and decreases Young’s modulus (Figure 5D), and expression of vasoconstriction factors such as SM-22α, α-SMA, MLCK, Calponin and Myh11 (Figures 5E,F). Interestingly, Figures 5B–D demonstrate that nebivolol restores aortic functions, at levels observed in young, non-diabetic mice.

From an anatomical point of view, treatment with nebivolol also significantly reduced adventitia and media-intima thicknesses (Figure 6A and Supplementary Figure 1B). Importantly, collagen modifications cannot explain by itself thicknesses reduction after treatment. Indeed, we noticed that the mRNA expression of types I and III were significantly increased (Figure 6B) while collagen staining by red picrosirius, in the aortic adventitia was reduced (Figure 6C). Moreover, total collagen level (Figure 6D) and crosslinks quantities (Figure 6E) were comparable to those observed in untreated db/db mice. Similar to minoxidil, nebivolol treatment induces a decrease in pro-fibrotic inflammatory factors in the aorta of diabetic mice (Figure 6F) as well as in their plasma (Figure 6G) compared to untreated db/db mice. Using Western blotting (Supplementary Figure 2), elastin autofluorescence (Figure 7A) and extraction of insoluble elastin (Figure 7B), we demonstrated an increase of elastin quantity after nebivolol treatment. Using Hart’s staining (Figure 7C) or by elastin autofluorescence (Figure 7A), we observed that, although the number of elastic lamellae was unchanged in the media (Figure 7D), their integrity was improved by nebivolol treatment. This was evidenced by the dramatic decrease in elastic lamellae ruptures (Figure 7E), and the decreased plasma levels of EDP and desmosines (Figure 7F). In parallel, nebivolol treatment decreased plasma cathepsin S and NE activities (Figure 7G), while, surprisingly, the expression of cathepsin S within the aorta (Figure 7H) was increased. The expression of the natural inhibitors of MMP9 and NE elastases, say TIMP1 and SERPIN, was significantly increased by nebivolol treatment (Figure 7I). Nebivolol treatment also induced the expression of elastin and LOXL1 and decreased LOXL3 expression (Figure 7J), suggesting a positive effect on elastogenesis. Figure 7K shows that this neosynthesis of elastin (Supplementary Figure 2) was associated with an increase of all crosslink classes, suggesting the formation of mature and functional elastin. The consequence of the treatment was a spectacular drop in aortic rigidity (by 5–10 times), as measured by AFM (Figure 7L). Altogether, these findings suggest that chronic treatment with nebivolol can limit the EFs premature aging, with fewer cardiovascular side effects.

As described in the literature (Inyard et al., 2009; Standley et al., 1991), diabetes promotes in vivo vascular smooth muscle contraction processes and the expression of elastolysis markers. In contrast, the use of the antihypertensive molecules minoxidil or nebivolol in diabetic animals induced a decrease in the markers of SMC contraction and an increase in the markers of elastogenesis. In this context, we aimed to focus on the effects of the treatments, Minoxidil and Nebivolol, on the behavior of insulin-resistant smooth muscle cells (SMCs). For this purpose, SMCs were pre-incubated with a medium enriched in glucose-palmitate. To maintain consistency between in vivo models (male db/db mice from the C57/Bl6J strain) and in vitro conditions, we opted to utilize the MOVAS cell line, derived from SMCs of male C57Bl6J mice. Similar to endothelial cells, SMCs play a crucial role in preserving the homeostasis of the ECM. In Figure 8, insulin was used as a positive control having the ability to induce hyperpolarization of SMC membranes (Zierler and Rogus, 1980; Zierler, 1966). Glucose-palmitate treatment decreased the phosphorylation levels of the insulin receptor and related pathway actors, Akt and transcription factor FOXO1 (Figure 8A). Palmitate-glucose condition also decreases glucose up-take and increases mRNA PEPCK expression, known to be under the control of FOXO1 activity (Supplementary Figure 3). Together, these data suggested that palmitate-glucose condition induced an insulin-resistance in MOVAS cells. The glucose-palmitate condition significantly increased α-SMA, MLCK, Calponin and Myh11 expressions (Figure 8B) whereas, when associated with insulin, minoxidil, or nebivolol, we observed a decrease in its expression, returning close to that of insulin alone (in the absence of insulino resistance). This suggests that in vitro insulino resistance could favor the contraction of MOVAS cell line, isolated from the smooth muscle of an adult mouse, by changing the levels of several genes involved in contraction. Interestingly, those expressions of “contractil genes” are associated to an increase of pro-inflammatory cytokines (TNFα, IL6 or IL1b) in insulin-resistance condition, while the presence of minoxidil or nebivolol reduces significantly the expressions of IL6 and IL1b (Supplementary Figure 3). Figures 8C, 9A show that insulin stimulates mRNA expression of elastogenesis markers (elastin, fibrillin 1, fibulin 5, LTBP4, and LOXL1) and elastin production (Figures 8D,E). Conversely, the induction of insulin resistance (despite the presence of insulin) does not elevate the expressions of these same markers or even decrease their expressions, with the possible exception of LTBP4. The addition of minoxidil or nebivolol in the presence of glucose-palmitate increases mRNA (Figures 8C, 9A) and protein (Figures 8F,G) levels of all elastogenesis markers except for LOXL1. Insulin alone has no effect on elastolysis markers, MMP-9 and cathepsin S (Figure 9B). Glucose-palmitate-induced insulin resistance significantly increased the expression of both elastases, and this effect was abolished by minoxidil or nebivolol. Surprisingly, minoxidil and nebivolol also reduced the expression of MMP-9 natural inhibitor, TIMP1 (Figure 9B). When we ratioed the expression of an elastase to that of its natural inhibitor (i.e., MMP9/TIMP1 and cathepsin S/cystatin C) (Figure 9C), we observed that insulin resistance increased the cathepsin S/cystatin C ratio, and this effect was abolished by insulin, minoxidil, and nebivolol. Conversely, glucose-palmitate reduced the MMP9/TIMP1 ratio, and neither insulin, minoxidil, nor nebivolol could modify this effect (Figure 9C). These data suggest that insulin resistance promotes MOVAS cell contraction and favors elastolysis, whereas smooth muscle relaxants, such as insulin, minoxidil, and nebivolol, are pro-elastogenic factors. In correlation studies conducted in db/db mice treated with minoxidil and nebivolol, a negative correlation was observed between contraction markers (αSMA and SM22) and elastogenic factors (elastin and LOXL1) (Supplementary Table 2).

Minoxidil was described as a potassium channel opener that hyperpolarizes cell membranes, causing vascular muscle relaxation and a consequent increase in blood flow (Katsuumi et al., 2018; Tam et al., 2020). The vasorelaxant effect of nebivolol has been primarily attributed to endothelial-dependent mechanisms, including beta-adrenergic receptors. However, nebivolol present additional vasorelaxant properties. Thus, the involvement of the ATP-sensitive potassium channels (KATP) of the SMCs would be a second mechanism involved in the vasorelaxant response to nebivolol (Altunkaynak-Camca, 2020; Haeusler, 1990; Kaiser et al., 2006; Korkmaz et al., 2015; Slove et al., 2013; Yasui et al., 2008). Nebivolol might appear to act indirectly on the channel by decreasing cytoplasmic ATP concentrations by inhibiting mitochondrial ATP synthase and/or increase of extracellular efflux of ATP (Supplementary Figure 4 and (Kremastiotis et al., 2021; Lafarge et al., 2010; Lafarge et al., 2014; Le et al., 2014)). The opening of potassium channels by minoxidil or by nebivolol can be associated with the induction of elastogenesis and the inhibition of elastolysis. In contrast, Supplementary Figure 5 shows that the closure of the voltage-gated potassium channels by tetraethylammonium (TEA) or of KATP channel by glibenclamide (Lebrun et al., 1997) (Lee and Dong, 2017) is associated with a decrease in most markers (protein or transcripts) of elastogenesis while those of elastolysis are expressed. To determinate if contractile status of MOVAS cells is determinant factor, we induced membrane depolarization of cells by addition of KCl in culture medium. Prior to the study, we evaluated the cytotoxic effect of extracellular KCl addition on MOVAS cell survival (Supplementary Figure 6). Excess of extracellular potassium effectively induces cell contraction but does not seem to have any significant effect on elastogenesis and elastolysis (Supplementary Figure 5). On other hand, the presence of KCl (or glibenclamide) inhibits the effects of minoxidil or nebivolol on the expressions of transcripts or proteins such as elastin. Taken together, these results suggest that the configuration of the potassium channels (opened or closed) is a major element in the control of the elastogenesis-elastolysis balance. Therefore, to determine the signaling pathway linking potassium channel and transcript expressions is important. Several studies (Hu et al., 2008; Tinker et al., 2014) suggested that the decrease of KATP channel function leads to FOXO-1 repression.

We hypothesized that the KATP channel pathway could activate the transcription factor FOXO1, involved in premature aging. Interestingly, data from the literature (Castillero et al., 2013; Nakae et al., 2001) suggest that insulin signaling pathways modulate FOXO1 activity, in accordance with our findings regarding the insulin signaling pathway (Figure 8A). In addition, we have shown that variations in elastin expression follow variations in expression and phosphorylation of FOXO1 (Figures 8F,G) depending on whether the SMCs are insulin-resistant or not or treated or not with minoxidil or nebivolol. The increased phosphorylation of FOXO associated with elastin expression was also observed in the aortas of db/db mice treated with minoxidil or nebivolol (Supplementary Figures 2A,B). Thus, we initially undertook a transcriptomic study of FOXO1 and FOXO3 on MOVAS cells incubated in different media, stimulating the opening or closing of ATP-sensitive potassium channels (see Figure 10A). In cells incubated in a classical medium (DMEM), the presence of the KATP channel openers minoxidil and nebivolol drastically reduced the expressions of FOXO1 and, for nebivolol only, FOXO3. Conversely, three KATP channels closing conditions (culture medium with glibenclamide, KCl, or glucose-palmitate) significantly increased the expression of FOXO1, whereas only KCl and glucose-palmitate elevated FOXO3 mRNA levels. KCl- or glucose-palmitate-supplemented culture medium, minoxidil, or nebivolol caused a significant decrease in FOXO1 and FOXO3 expressions compared to glucose-palmitate or KCl alone, while they decreased FOXO1 and increased FOXO3 expressions in the presence of glibenclamide compared to glucose-palmitate or KCl alone (Figure 10A). Interestingly, FOXO1 has been described as a potential regulator of elastin (Shi et al., 2012) and MMP-9 expressions (Yang et al., 2020; Zhang et al., 2017). Therefore, to confirm the impact of FOXO1 on ELN and MMP-9 expression, we used the FOXO1 transcription inhibitor AS1842856 (Figure 10B). Inhibition of FOXO1 activity by AS1842856 in insulin resistance conditions (palmitate + glucose) increased elastin and decreased MMP-9 expression compared to palmitate and glucose alone. We observed similar effects of AS1842856 on elastin and MMP-9 expressions in a glibenclamide (or KCl)-supplemented culture medium compared to glibenclamide or KCl alone. Surprisingly, in conditions where KATP channels were opened by minoxidil or in the presence of nebivolol, the FOXO1 inhibitor AS1842856 reduced the expression of MMP-9 and elastin (Figure 10B).