miR-181b regulates vascular stiffness age dependently in part by regulating TGF-β signaling

Background Endothelial dysfunction and arterial stiffening play major roles in cardiovascular diseases. The critical role for the miR-181 family in vascular inflammation has been documented. Here we tested whether the miR-181 family can influence the pathogenesis of hypertension and vascular stiffening. Methods and results qPCR data showed a significant decrease in miR-181b expression in the aorta of the older mice. Eight miR-181a1/b1-/- mice and wild types (C57BL6J:WT) were followed weekly for pulse wave velocity (PWV) and blood pressure measurements. After 20 weeks, the mice were tested for endothelial function and aortic modulus. There was a progressive increase in PWV and higher systolic blood pressure in miR-181a1/b1-/- mice compared with WTs. At 21 weeks, aortic modulus was significantly greater in the miR-181a1/b1-/- group, and serum TGF-β was found to be elevated at this time. A luciferase reporter assay confirmed miR-181b targets TGF-βi (TGF-β induced) in the aortic VSMCs. In contrast, wire myography revealed unaltered endothelial function along with higher nitric oxide production in the miR-181a1/b1-/- group. Cultured VECs and VSMCs from the mouse aorta showed more secreted TGF-β in VSMCs of the miR-181a1/b1-/- group; whereas, no change was observed from VECs. Circulating levels of angiotensin II were similar in both groups. Treatment with losartan (0.6 g/L) prevented the increase in PWV, blood pressure, and vascular stiffness in miR-181a1/b1-/- mice. Immunohistochemistry and western blot for p-SMAD2/3 validated the inhibitory effect of losartan on TGF-β signaling in miR-181a1/b1-/- mice. Conclusions Decreased miR-181b with aging plays a critical role in ECM remodeling by removing the brake on the TGF-β, pSMAD2/3 pathway.


Methods and results
qPCR data showed a significant decrease in miR-181b expression in the aorta of the older mice. Eight miR-181a1/b1 -/mice and wild types (C57BL6J:WT) were followed weekly for pulse wave velocity (PWV) and blood pressure measurements. After 20 weeks, the mice were tested for endothelial function and aortic modulus. There was a progressive increase in PWV and higher systolic blood pressure in miR-181a1/b1 -/mice compared with WTs. At 21 weeks, aortic modulus was significantly greater in the miR-181a1/b1 -/group, and serum TGF-β was found to be elevated at this time. A luciferase reporter assay confirmed miR-181b targets TGF-βi (TGF-β induced) in the aortic VSMCs. In contrast, wire myography revealed unaltered endothelial function along with higher nitric oxide production in the miR-181a1/b1 -/group. Cultured VECs and VSMCs from the mouse aorta showed more secreted TGF-β in VSMCs of the miR-181a1/b1 -/group; whereas, no change was observed from VECs. Circulating levels of angiotensin II were similar in both groups. Treatment with losartan (0.6 g/L) prevented the increase in PWV, blood pressure, and vascular stiffness in miR-181a1/b1 -/mice. Immunohistochemistry and western blot for p-SMAD2/3 validated the inhibitory effect of losartan on TGF-β signaling in miR-181a1/b1 -/mice. PLOS

Introduction
Hypertension is a major cause of morbidity and mortality worldwide. Increased aortic stiffness is one of the major factors that disproportionately increases central arterial pressure, creates systolic hypertension, and increases pulse pressure. [1][2][3][4] The remodeling of extracellular matrix (ECM) by decreased elastin or it's fracturing, increased collagen deposition, endothelial dysfunction, and cross-linking of ECM, are the major contributors of increased vascular stiffness. [1,5] Many miRNAs contribute to vascular dysfunction, but among these, miR-145 and miR-181b have been found to be key regulators of vascular inflammation in ApoE signaling pathways. [6,7] In the ApoE-deficient mouse model, delivering miR-181b was found to protect against vascular inflammation by directly binding at the 3'-UTR of importin-α3, a key regulator of the NF-κB signaling pathway. [7,8] From human plasma samples, it has been shown that in coronary artery disease, the circulating level of miR-181b expression is significantly lower compared to healthy controls. [7] Furthermore, miR-181a has been reported to be decreased in vascular smooth muscle cells (VSMCs) when stimulated by angiotensin II, whereas overexpression of miR-181a resulted in inhibition of adhesion of VSMCs to collagen and reduced expression of osteopontin, a multifunctional protein found in abundance in atherosclerotic plaques. [9] The miR-181 family has also been reported to play an essential role in early NKT cell development by targeting 3'-UTR of PTEN in the thymus. [10] Together, these results suggest that the miR-181 family is an important modulator of vascular inflammation. [11] However, the role of the miR-181 family in overall vascular function, including endothelial function and vascular stiffness, remains unknown.
We hypothesized that the chronic depletion of miR-181a/b will lead to vascular stiffening either by endothelial dysfunction or by ECM remodeling or their combination. To study this, we used miR-181a1/b1 -/mice, and compared their aortic stiffness and blood pressure with WT mice. Better understanding of the role of miR-181 in the development of vascular stiffness and systolic hypertension may lead to novel therapeutic approaches in the future.

Material and methods Animals
We used male mice deficient for the miRNA clusters miR-181a1b1 (containing the miR-181a-1 and miR-181b-1 cluster, located on chromosome 1) (miR-181a1/b1 -/-). [10] qPCR data confirmed that both miR-181a1 and miR-181b1 are knocked-out from the aortic tissue of the mice (Fig 1A and 1B). This mouse model was generated in the C57BL6/J genetic background. [10] Chimeric offspring were backcrossed up to ten generations with C57BL/6 mice. Germline transmission was confirmed by PCR of tail genomic DNA with the primers 5 0 -AAATGCTTAT TCCATGCACATT-3 0 and 5 0 -ATCAACGGTCGATGGTTTT-3 0 which amplifies 554 and 647base pairs products from the wildtype allele and floxed allele, respectively. The primers 5 0 -AA ATGCTTATTCCATGCACATT-3 0 and 5 0 -TGAGCCCCTGGATAACAAAG-3 0 amplify a 377-base pair product from the deleted allele. Consistent with the previous study using the same genotyped mice [10,12], we also used C57BL/6 mice as the control group. All mice had the same C57BL/6 background. 25 mice from both phenotypes were used for these experiments. They were provided with food and water ad libitum, and house them in 7 am to 9 pm light and rest of the time dark cycle facility. Mice were treated humanely and all experimental procedures were approved by the Institutional Animal Care and Use Committee of Johns Hopkins University. Animal care staffs regularly monitor the mice, and all the animals looked healthy during the experimental protocols. We used Ketamine-Xylazine to anesthetize the animal (check for total anesthesia by toe pinching) before we excise the organs.

Non-invasive Pulse Wave Velocity (PWV) Measurements
A high-frequency, high-resolution Doppler spectrum analyzer (DSPW, Indus Instruments, Houston, TX, USA) was used. Mice were anesthetized with 1.5% isoflurane, and were placed in a supine position on a heated pad equipped with EKG. The animals were allowed to stabilize to a physiologic heart rate following which a 20 MHz probe was used to measure the descending aortic and abdominal aortic flow velocities. The time between the R wave of EKG to the start of pulse wave form for each measurement location was calculated using a real-time signal acquisition and spectrum analyzer system. [14,15] Non-invasive blood pressure measurement Blood pressures (BP) were measured by tail-cuff non-invasive BP measurement system using volume pressure recording sensors (CODA, Kent Scientific Corp, Torrington, CT, USA). The animals were trained in a holder every day for 1 week. BP measurements were performed on a heated platform set at 37˚C. Occlusion cuff and volume pressure recording cuff was placed on the tail and 20 measurements were acquired. The average of at least five consecutive readings were used for the analysis. [16] Tensile testing Thoracic aortas were harvested and cut into 2mm rings. The aortic ring and a 0.5mm segment proximal to each ring was imaged at ×10 magnification to measure inner diameters (D i ), outer diameters (D o ), and the vessel length (L). The diameter and the length were measured with Image J software (National Institutes of Health [NIH], Bethesda, MD, USA). The aortic rings were mounted onto the pins of an electromechanical puller (DMT560; Danish Myo Technology A/S, Aarhus, Denmark). After calibration and alignment, the pins were slowly moved apart using an electromotor at a rate of 20 μm/sec to apply radial stress on the specimen until breakage. Displacement and force were recorded continuously. Engineering stress (S) was calculated by normalizing force (F) to the initial stress-free area of the specimen (S = F/2t×l; where t = thickness and l = length of the sample). Engineering strain (λ) was calculated as the ratio of displacement to the initial stress-free diameter (Di). The stress-strain relationship was represented by the equation S = α exp (βλ), where α and β are constants. α and β were determined by nonlinear regression for each sample using Excel (Microsoft, Redmond, WA, USA). [5] Decellularization Aortic rings were harvested from 21 week old miR-181a1/b1 -/mice and C57BL6 mice following a protocol described previously [15] with minor modifications. A 0.1% SDS solution was prepared in 50mM NH4OH. The aortic rings were placed in the microcentrifuge tubes containing the solution and were placed on the vortexer/shaker for 24 hrs following which the rings were rinsed with PBS for 15 minutes on the vortexer/shaker twice.

Endothelial function (vasoreactivity)
Thoracic aortas were isolated, and trimmed into 2mm rings and mounted on a wire myograph (DMT). One end of the aortic ring was connected to a transducer and the other to a micromanipulator. The vessels were immersed in a bath filled with constantly oxygenated Krebs buffer (NaCl 118.3 mM, KCL 4.7 mM, CaCl2 1.6 mM, KH2PO4 1.2mM, NaHCO3 25mM, MgSO4 1.2mM, and Dextrose 11.1mM) at 37˚C. The vessels were stretched to an optimal resting tension using the micromanipulator. 60 mM KCl was administered, and repeated after three washes with Krebs buffer. Vessels were then allowed to equilibrate for 1 hour. The vessels were pre-constricted with Phenylephrine (1 μM) for 15 minutes and ACh-induced dose response relaxations (1nM to 10uM) and SNP-induced dose response relaxations (1nM to 10uM) were measured. Relaxation responses were calculated as a percentage of tension after preconstriction. [14,17,18] No production measurement NIS-Element suite (Nikon Corporation, Shinagawa, Tokyo, Japan), with excitation and emission wavelengths set to 485 and 510 nm, respectively. Fluorescence intensity was recorded every 30 s for a period of 10 min. Acetylcholine (10uM) was administered on to the sample and the slope of intensity change was calculated for a period of 5 minutes to indicate rate of NO production. Data was then normalized to WT baseline measurement = 100%. [19] TGF-β and angeotensin II measurement TGF-β1 ELISA kit (R&D Systems Inc., Minneapolis, MN, USA) and Angiotensin II EIA kit (Sigma-Aldrich, St. Louis, MO, USA) were used to detect TGF-β and Angiotensin II from mice serum and cell culture media as directed by the manufacturer's instructions.

Cell culture
Aortic rings were harvested from 10 week old miR-181a1/b1 -/mice and C57BL6 mice following a protocol described by Kobayashi et al. [20] with minor modifications. The rings were cut open to make aortic strips and then were placed into microcentrifuge tubes containing 2 mg/ ml collagenase. The tubes were gently rocked for 30 minutes at 37˚C before centrifugation at 3000 rpm for 10 minutes. Supernatant was discarded and the pellets were resuspended in DMEM containing 20% serum and transferred into 30 mm cell-culture dishes. After 2 hours, non-adhered debris was removed and replaced with ECM media (ScienCell Research Laboratories, Carlsbad, CA, USA) for endothelial cell culture. The media were replaced every 48 hours until VECs reached confluence. Aortic strips were further transferred to 30 mm dishes containing DMEM supplemented with 20% serum for VSMC culture. Samples were maintained for 7-10 days to obtain confluent VSMCs. At passage 2, media was washed out and were replaced with serum free media. After 24 hours of incubation, media was collected, centrifuged and the supernatant was stored at -80˚C for TGF-β measurement by ELISA. Cells were lysed into 1xRadioimmunoprecipitation assay (RIPA) buffer containing protease and phosphatase inhibitors.

Immunoblot analysis
30 μg of protein were resolved by SDS-PAGE and electro-transferred to PVDF membranes for western blotting. Primary antibodies that recognize p-SMAD2/3 and SMAD2/3 were from Santa Cruz Biotechnology (Rabbit). The p-eNOS (Ser 1177) and eNOS antibodies were from Cell Signaling Technology (Danvers, MA, USA). The α-tubulin and Angiotensin II receptor 2 antibodies were from Abcam (Cambridge, MA, USA). VE-Cadherin and α-Actin were from Santa Cruz Biotechnology (Starr County, TX, USA). Immunoreactive proteins were visualized using an enhanced chemiluminescence analysis kit (EMD Millipore, Billerica, MA, USA).

Digital image analysis
Slides that were stained for collagen and pSMAD2/3 were digitized on an Aperio AT system (Leica Biosystems, Nussloch, Germany). Using Aperio ImageScope tools, the area of staining were measured. That area was divided by the total area of the vascular layer or the vascular media in the same aortic ring section.
Luciferase reporter assays 1.25 X 10 6 rat aortic vascular smooth muscle cells (A7r5) were transfected with 50 ng of TGFβi 3'-UTR reporter construct (GeneCopoeia, Rockville, MD, USA) or with mutated TGF-βi reporter construct (mutagenesis at positions 72,74,76 and 77 from "A to C", "A to C", "G to C", and "U to A", respectively) with dual luciferase, firefly and renilla, which were then cotransfected with either scramble sequence or by mature miR-181b (GE-Dharmacon, Lafayette, CO, USA), using an electroporator (Neon Transfection System, Thermo Fisher, Carlsbad, CA, USA) following 1475 mV for 20 ms and 2 pulses. 48 hours after transfection, cells were washed lysed and assayed for firefly and renilla (served as internal control) luciferase activity using the Dual-Luciferase Reporter Assay System (Promega, Madison, WI, USA). The primer sequences to performed mutagenesis experiment are provided in Table 1.

Losartan preparation protocol
Losartan concentration of 0.6 grams/liter was prepared to give an estimated losartan dose of 50-70 mg/kg/day in the drinking water. [21,22] 650 mg of losartan tablets were dissolved in 1080 ml of water to achieve a concentration of 0.6 g/L. The mixture was then double-filtered using a vacuum filtration system to remove the remaining excess sediment. [23] For cell culture purposes, we used 10 μM Losartan (Cayman Chemicals, Ann Arbor, Michigan, USA) for 48 hours. [24,25]

Statistical analysis
The results are presented as mean and standard error of measurement (mean ± SEM). Vascular relaxation for myograph was analyzed by t-test and LogEC50 by Extra sum-of-squares F test. For multiple comparisons, one-way analysis of variance (ANOVA) and the Bonferroni post hoc test were used. P<0.05 was considered as statistically significant. All analysis was performed using Prism 5 (GraphPad Software Inc., La Jolla, CA, USA).

miR-181a/b is not associated with circulating levels of Angiotensin II and expression of Angiotensin II receptors
The Angiotensin II levels in the serum of the mice were tested by ELISA. (Fig 7A) There were no significant differences in Angiotensin II in the WT compared to the miR-181a1/b1 -/group. (WT: 4.62±0.22vs. a/b KO: 3.65±0.43, p = 0.09). (Fig 7A) Further, although there was a trend towards higher Angiotensin II in those mice treated with losartan, there were no significant differences between the WT and the miR-181a1/b1 -/group treated with losartan. (WT: 5.51±0.97 vs. a/b KO: 4.89±0.55, p = 0.60). (Fig 7A) Western blot of Angiotensin II receptor in the aortic vessels were also similar between the two groups. (Fig 7B) Losartan mitigates age-associated increase in BP and PWV in miR-181a/b deficient mice WT and a/b KO mice were treated with losartan (0.6 g/L) in drinking water starting at 8 weeks of age. Losartan significantly lowered the BP and PWVs (Fig 8A) in a/b KO mice compared to untreated age-matched mice from 10 weeks until 20 weeks of age. Mechanical modulus of the aorta as assessed by tensile testing also showed significant attenuation in stiffness in losartan treated miR-181a1/b1 -/mice ( Fig 8B). Interestingly, WT, WT + Losartan, and a/b KO + Losartan, showed identical mechanical properties and PWV profiles (Fig 8B and 8C). This further suggests that depletion of miR-181a/b promotes vascular stiffness via ECM remodeling by activating TGFβ signaling in VSMCs. Both IHC (Fig 8D) and western blot (Fig 8E)  and not treated with losartan (n = 11), miR-181a1/b1 -/-(a/b KO) mice treated (n = 11) and not treated with losartan (n = 10), decellularized aortic rings from control (WT) mice treated (n = 10) and not treated with losartan (n = 8), and decellularized aortic rings from miR-181a1/b1 -/-(a/b KO) mice treated (n = 13) and not treated with losartan (n = 9). The bar graph shows the strength of the vessel at 1.5mm in which the bar represents mean and the error bar represent SEM. D, Immunohistochemistry of p-SMAD2/3 staining of the aorta treated with losartan in miR-181a1/b1 -/-(a/b KO) and control (WT) mice. E, Western blot of p-SMAD2/3 compared to total SMAD2/3 in the aorta from miR-181a1/b1 -/-(a/b KO) and control (WT) mice treated and not treated with losartan (n = 5). Values are mean ±SEM, *p<0.05, **p<0.01, ***p<0.001.

Discussion
Hypertension is one of the major health problems of our aging population. [26,27] Vascular stiffness is an important component of systolic hypertension and treatment to modulate vascular stiffness is suggested, as PWV has become a powerful predictor of cardiovascular events. [4,26,28,29] Over the past few years, there have been numerous studies that have pointed to a major role of miRNA in hypertensive heart disease. [30][31][32] For example, patients with coronary artery disease have reduced circulating levels of miR-181b, suggesting the potential role of miR-181b in the pathogenesis of arterial inflammation. [7,8] Accumulating evidence indicates that miR-181 modulates vascular function by targeting multiple key cell signaling pathways, including NF-κB signaling in the vascular endothelium as well as the PI-kinase pathway. [7,8,10,33] However, there is no evidence showing the role of miR-181 in baseline vascular function. We used miR-181a1/b1-deficient mice to study this phenomenon, and have identified a phenotype associated with vascular stiffening due to activation of the TGF-β signaling cascade.
[10] (Fig 2A) Chronic down-regulation of miR-181b expression with age was associated with activation of TGF-β signaling in the VSMCs (Fig 6D, 6E, 6F, 6G and 6H). Excessive TGF-β production from VSMCs has been shown to modulate ECM growth and function. [32,34,35] Activated TGF-β is found to be increased in aged aortas and contributes to elastic artery stiffening with age. [36,37] TGF-β can initiate multiple effects in the vessels including phenotypic modulation of the VECs and VSMCs. It is also associated with induction of gene expression such as collagen I and III, stimulating production of ECM. Furthermore, it also stimulates plasminogen activator-inhibitor production, which inhibits breakdown of ECM, resulting in increased ECM. [38] These cellular signaling mechanisms contribute to vascular remodeling as well as primary and secondary forms of hypertension in atherosclerosis. [38] A role for collagen deposition in the adventia of the aortic vessels has also been suggested to cause vascular stiffness potentially leading to systolic hypertension. [39] We have also demonstrated in this study that TGF-β can be the direct target of miR-181b in the VSMCs, where miR-181b binds directly to the 3'-UTR of TGF-βi. (Fig 6H) TGF-βi (transforming growth factor, beta induced) is one of the most important components of the TGF-β signaling pathway, and it is highly abundant in the cardiovascular system. This gene encodes an RGD-containing protein that binds to type I, II and IV collagens. Thus, this protein plays a role in cell-collagen interactions (NM_009369.4). In theory, the entire miR-181 family can bind to the same 3'-UTR region of TGF-β ligand. Among the six mature family members, miR-181 a1, a2, b1, b2, c, and d, it has been found that miR-181a1 and miR-181b1 are the most prevalent in the aorta of mice. [7,40,41] The mature sequence of miR-181a1 and miR-181a2, and miR-181b1 and miR-181b2 are identical, but they are encoded from two different genomic loci: the miR-181a1 and miR-181b1 cluster is located on chromosome 1, and the miR-181a2 and miR-181b2 cluster is located on chromosome 9. [42] To test the baseline vascular function of miR-181 family in vivo, we used mice deficient for the miR-181a1-miR-181b1 cluster. [10] The miR-181a1-miR-181b1 cluster is mainly expressed in blood vessels, and both may contribute to vascular stiffness by regulating TGF-β signaling in the VSMCs. Intriguingly, VECs of the miR-181a1/b1 -/mice were associated with increased eNOS activity (Fig 4), which likely blunts some effects of increased BP and vascular stiffness (Figs 2 and 4). This phenomenon might be explained by the proposed effect of TGF-β on eNOS activity reported previously. Saura, et al. [43] reported an alternative function of TGF-β which induces eNOS expression mediated via SMAD2. Further, Heger et al. [44] reported a role for TGF-β in eNOS phosphorylation and increased NO release independent of SMAD activation. It is also possible that the increase in NOS may actually be uncoupled NOS, which may be deterious to endothelial function and may also stimulate atherosclerosis. [45] There are several phosphorylation sites on eNOS. The majority of work has focused however on 2 residues, serine 1177 (Ser 1177) and threonine 495 (Thr 495). [46] It has been well documented that Akt specifically induces phosphorylation of Ser 1177 and that increases NO production. On the other hand, phosphorylation of Thr 495 has been reported to downregulate the generation of NO. Since we measured phospho-eNOS at serine 1177, it is likely that the increase in phospho-eNOS 1177 leads to an increase rather than a decrease in NO production, and not to uncoupling, reduced NO, and increased reactive oxygen species production. Furthermore, the myography experiment did not show any endothelial dysfunction nor did we find any noticeable atherosclerotic changes in the a/b KO vascular ring. Several studies have pointed out the important role of miR-181b in VECs. [7,[47][48][49] It has been validated that miR-181b regulates NFκB signaling by targeting importin-α3. [7,47] Thus loss of miR-181b can activate NFκB signaling in VECs. Several studies has pointed out that NF-κB is a negative regulator of eNOS expression in VECs. [50,51] Systemic administration of miR-181b reduced downstream NF-κB signaling by targeting importin-α3 in the vascular endothelium. [47] Recently, miR-181b has been shown to target caspase recruitment domain family member 10 (Card10) in ECs to prevent thrombin-mediated endothelial activation and arterial thrombosis. [49] This could explain why we have noticed higher p-eNOS in our miR-181a1b1 -/mice. As the role of miR-181b in VECs is a very well studied topic, and vascular stiffness is mainly due to functional changes of VSMCs, we focused our study on the underlying mechanism involving miR-181b in VSMCs rather than VECs.
Losartan, an angiotensin II-type 1 receptor blocker and a known TGF-β neutralizer, is a very effective anti-hypertension medication. [23] Our results showed that treatment with losartan (0.6 g/L), which not only lowers the BP, but neutralizes TGF-β signaling, was able to mimic the role of miR-181a1/b1 during the entire time span of the experiment (11 weeks to 20 weeks), and reversed the vascular stiffness phenotype of miR-181a1/b1 -/mice. [23,52,53] (Fig  8) Interestingly, our result showed no difference in the Angiotensin II levels in the serum of the WT and the miR-181a1/b1 -/mice. This indicates that miR-181b is associated with direct inhibition of TGF-β signaling independent of Angiotensin II. Several studies have suggested that TGF-β induces miR-181 expression. [54][55][56][57] However, our result have also shown that miR-181b can directly target the 3'-UTR of TGF-βi (Fig 6H and 6I). This suggests that miR-181b may have a role in a feedback loop of TGF-β signaling. Further, there were no significant alterations in TGF-β release after inhibition of TGF-β receptor with D4476 in the VSMCs. This may suggest that the role of miR-181b in the feedback loop of the TGF-β cascade is upstream of the TGF-β receptor. Although the vascular stiffness phenotype was reversed following treatment with the TGF-β neutralizer, losartan, (2,9,27) losartan did not alter miR-181b expression in the VSMCs, suggesting that reversal of vascular stiffness by losartan was due to inhibition of TGF-β signaling downstream of miR-181b. Although its mechanism remains to be fully elucidated, usage of Losartan has been reported to be associated with reduced expression of TGFβ ligands, receptors, and activators. (13) These results may suggest that the main role of losartan in the reversal of the phenotype was not only due to reduced expression of TGFβ ligands which may not have been significant as we did not see any changes in the expression of miRNA-181b, but by downstream inhibition of TFG-β targets.
Hypertension and increased vascular stiffness are common with aging. [28,58] Reducing vascular stiffness could be an important adjunct to controlling blood pressure to reduce the risk of cardiovascular events in the elderly. We have identified the potential role of miR-181a1/ b1 in the development of vascular stiffness, which ultimately was associated with higher systolic blood pressure via activation of TGF-β. Even though losartan is currently in widespread clinical use for the management of hypertension and stroke, miR-181b could potentially be a miR-181 family and vascular stiffness new or additive therapeutic target for intervention, targeting the TGF-β cascade independent of Angiotensin II.

Conclusion
More than 500 miRNAs have been identified in humans, with few of them being involved in regulating smooth muscle cell phenotype, and modulating inflammatory response in endothelial cells and macrophages in the process of arterial remodeling and atherosclerosis. [59] The present study shows a role of miR-181b in regulating TGF-β signaling, in the smooth muscle cells, which modulates vascular stiffness. Increased TGF-β expression is observed in aged aorta, which is associated with increased extracellular matrix deposition, increased cross-linking and ultimately vascular remodeling. [38,[60][61][62] Not only is lowering blood pressure an important target in clinical practice today, but central vascular stiffness as determined by pulse wave velocity, has become both a powerful predictor of cardiovascular events, and an important parameter for targeting therapy. [26] miRNA181b could be a potential therapeutic target in modulating vascular stiffness along with the treatment of hypertension.
Supporting information S1 File. Fig A. Transfection efficiency of rat aortic smooth muscle cells. Cell image represent transfection efficiency of A7r5 cell lines using electroporation. Red color represents mCherry tagged beta-actin. Green color represents FITC tagged at the 3'-end of miR-181b.