Soluble (pro)renin receptor induces endothelial dysfunction and hypertension in mice with diet-induced obesity via activation of angiotensin II type 1 receptor

Ziwei Fu1, Fei Wang2, Xiyang Liu1, Jiajia Hu1, Jiahui Su1, Xiaohan Lu2, Aihua Lu1, Jae Min Cho3,4, J. David Symons3,4, Chang-Jiang Zou2 and Tianxin Yang2 1Institute of Hypertension, Zhongshan School of Medicine, Sun Yat-sen University, Guangzhou 510080, China; 2Department of Internal Medicine, University of Utah and Veterans Affairs Medical Center, Salt Lake City, Utah, U.S.A.; 3Department of Nutrition and Integrative Physiology, University of Utah School of Medicine, Salt Lake City, Utah, U.S.A.; 4Division of Endocrinology, Metabolism, and Diabetes, Molecular Medicine Program, University of Utah School of Medicine, Salt Lake City, Utah, U.S.A.


Introduction
In 2002, Nguyen et al. cloned a novel receptor for prorenin and renin, termed it the (pro)renin receptor (PRR), and assigned a renin-regulatory function to this receptor [1]. On the basis of structure, PRR belongs to the type I transmembrane receptor family, consisting of a large N-terminal extracellular domain, a single transmembrane protein, and a short cytoplasmic domain [2]. The soluble (pro)renin receptor (sPRR) is generated by protease-mediated cleavage in the Golgi apparatus and is released to plasma or urine [3]. Although furin and ADAM19 were initially thought to be the cleavage enzyme, independent studies by Nakagawa et al. [4] and us [5] have consistently identified site-1 protease (S1P) as being the

Western blot analysis
HUVECs were lysed for 30-min on ice with a RIPA lysis buffer (Beyotime, Shanghai, China) including protease inhibitor cocktail (REF 04693116001, Roche, Basel, Switzerland). Next, whole cell lysates were collected, sonicated, and centrifuged for 10 min × 12000 rpm × 4 • C. The supernatant was used to detect protein (BCA Protein Assay Kit; cat. no.23225, Thermo, Rockford, U.S.A.). An equal amount of protein was loaded onto SDS-PAGE and transferred to polyvinylidene fluoride membranes (Millipore, Massachusetts, U.S.A.). After blocking with 5% skimmed milk for 1 h at room temperature, membranes were incubated with the appropriate primary antibodies overnight at 4 • C. Next, membranes were washed with TBST (NaCl 8.0 g, Tris 6.0 g, Tweeen-20 1.0%), and secondary antibodies including HRP-linked anti-mouse or HRP-linked anti-rabbit antibody were incubated at room temperature for 1 h. β-Actin (1:10,000, Sigma) was used as a loading control. Antibody labeling was visualized by addition of chemiluminescence substrate for detection of HRP (cat. no.WBKLS0500, Millipore) [5,26].

Quantitative RT-PCR
Total RNA was extracted using Trizol (Invitrogen) and then reverse transcribed to cDNA using the Transcriptor First Strand cDNA Synthesis Kit (REF 04897030001, Roche). Real-time PCR was performed on the AB Applied Biosystems using the Fast Start Universal SYBR Green Master (ROX) (REF 04913914001, Roche). Values were normalized using GAPDH content. The gene primers used in this study are shown in Supplementary Table S1 [25,26].

Measurement of H 2 O 2
H 2 O 2 was measured using the ROS-Glo H 2 O 2 Assay kit (cat. no. G8820, Promega, Wisconsin, U.S.A.) according to the manufacturer's instructions. The assay is based on a luminescent signal generated by a chemical reaction of H 2 O 2 and its substrate. Luminescence was measured using a FLUOstar OPTIMA microplate reader (BMG Labtech, Ortenberg, Germany) and the value was normalized to protein content [25].

Flow cytometry
Apoptosis was quantified with the Annexin V-FITC/PI Apoptosis Detection Kit (Beyotime, Shanghai, China) according to the manufacturer's instructions. At the end of the treatment, cells were trypsinized and then carefully transferred to a new centrifuge tube with corresponding supernatant. After centrifugation (1000 rpm) for 5 min, the cell pellet was washed with cold phosphate buffer saline (PBS), and resuspended in Annexin V-FITC/PI binding buffer to a concentration of 105 cells/195 μl. The cell suspension was stained with 5 μl Annexin V-FITC and 10 μl propidium iodide (PI) and then was incubated for 20 min at room temperature in the dark. Samples were collected for flow cytometry (Bechman Coulter Epics XL-MCL Flow Cytometer, Miami, Florida, U.S.A.) and data were analyzed using FlowJo (Ashland, Oregon, U.S.A.) software [26].

Ang II-I 125 radioactive ligand competitive binding assay
HUVECs were pretreated with 1 nM, 10 nM, 50 nM sPRR-His or 10 μM Losartan for 30 min followed by exposure to 1 μM Ang II-I 125 (NEX105050UC, perkinelmer, Massachusetts, Waltham, U.S.A.). To remove the unbound Ang II-I 125 , the cells were washed by PBS twice after removing cell medium. Next, HUVECs were lysed with a RIPA lysis buffer (Beyotime, Shanghai, China) including protease inhibitor cocktail (REF 04693116001, Roche, Basel, Switzerland). Whole cell lysates were collected, mixed, transferred to polystyrene tubes, and incubated at room temperature for 1 h, followed by the measurement of radioactivity using a gamma counter (SN-6105, Shanghai He Suo Ri Huan photoelectric instrument co., LTD, Shanghai, China). Non-specific binding (control group cells without Ang II-I 125 treatment) was subtracted from all measured values. The measured CPM represent Ang II-I 125 that is combined with cellular AT1R. All values were normalized to protein content of the sample [27].

Construction of plasmids and immunoprecipitation
The pcDNA3.1-sPRR-His plasmid was constructed by subcloning rat sPRR cDNA into the pcDNA3.1 vector. sPRR, a solubilized form of PRR (residues 17-274) lacking the transmembrane domain at the C terminus, was combined with a six-histidine tag in the C terminus (sPRR-His), was generated by PCR from the PRR expression construct, and then was cloned into pcDNA3.1 vector. pCMV6-AT1 plasmid was purchased from Origene, its four different mutants: pCMV6-AT1 S109A , pCMV6-AT1 K199A , pCMV6-AT1 N295A and pCMV6-AT1 S109A, K199A, N295A were also generated by PCR to introduce site-mutation or site-mutations into pCMV6-AT1 plasmid (the primers are shown in Supplementary Table S1). All generated plasmids were verified by DNA sequencing (DNA sequencing core, University of Utah). 35 S-labeled proteins were synthesized by T7 TNT in vitro transcription and in vitro translation system (Promega). These mammalian-based systems express soluble, functional proteins that are post-translationally modified. Expressed proteins can be used directly after expression; no additional protein purification needed. Following instructions from the manual, 1 μg of pcDNA3.1 vector, pcDNA3.1-sPRR-His, pCMV6-AT1 plasmid or its mutants with 2μl of 35 S-methionine (10 μCi/μl, PerkinElmer) were added to 40 μl of T7 TNT in vitro transcription and translation system at 30 • C for 2 h. To test the protein interaction, 0.5 μg of recombinant human AT1 protein (Novus Biologicals) or sPRR-His protein (XBIO, Shanghai, China) was incubated with the in vitro translation mixture at 4 • C overnight. The bound proteins to the unlabeled AT1 protein or sPRR-His protein were precipitated by incubating the mixture with 2 μg of the rabbit anti AT1 antibody (Santa Cruz Biotechnology, Inc.) pelleted with protein G agarose (Roche), or by incubating with 2 μg of PRR antibody (Sigma) against mouse PRR 32-45 AA (Thermo fisher Com.), antibody against mouse PRR 219-235 AA (Thermo fisher Com.) and then pelleted with protein A agarose (Thermo fisher Com.). The same amount of rabbit IgG was used as the control for these antibodies. Those pelleted products were dissolved in SDS loading buffer and separated by SDS-PAGE (polyacrylamide) gel. The radioactive signals were captured by a phosphor imager (LS6500 Multipurpose Scintillation Counter, Beckman Coulter, Indianapolis, U.S.A.).

S-sPRR radioactive ligand competitive binding assay
When HUVECs reached 50% confluence, 0, 5, 10 or 20 μM Losartan was added into the medium. One hour later, 35 S-sPRR-His was added to media at a dilution of 1:500. As mentioned earlier, 35 S-sPRR-His was obtained by in vitro translation. To eliminate free 35 S-methionine, the in vitro translation mixture was centrifuged with a 10 kD filter at 4200 Revolutions Per Minute (rpm) for 30 min. The free 35 S-methionine was filtered, and 35 S-sPRR-His was obtained in the unfiltered solution. Then, the unfiltered solution was diluted with culture medium (Lifeline) and added to the cultured cells. Thirty minutes, 3 or 15 h after adding the 35 S-sPRR-His, the medium was removed, cells were washed with PBS five times, and then treated with lysis buffer (Sigma Aldrich). The radioactivity of the cell lysate was measured with a multi-purpose scintillation counter (LS 6500 Beckman Coulter).

Nuclear and cytosolic separation
Nuclear proteins were isolated using a commercially available kit (Nuclear Extraction Kit, item No.10009277, Cayman, Ann Arbor, Michigan, U.S.A.), according to manufacturer's instructions. Briefly, adherent cells were dislodged following the respective treatments, collected in pre-chilled tubes, and centrifuged at 300 × g for 5-min at 4 • C. The pellet was resuspended in 5 ml of ice-cold PBS/phosphatase inhibitor solution, centrifuged at 300 × g for 5-min at 4 • C, and this step was repeated. Next, 500 μl ice-cold 1× complete hypotonic buffer was added to the pellet, and the cells incubated on ice for 15 min to allow cells to swell. After adding 100 μl of 10% NP-40 assay reagent to the mixture, the solution was centrifuged for 30 s at 4 • C. The supernatant containing the cytosolic fraction was transferred to a new tube. The pellet was resuspended in 100μl ice-cold complete nuclear extraction buffer, vortexed vigorously for 15-30 s, placed on ice on a rocker for 15-min, and centrifuged at 14,000 × g for 10 min at 4 • C. The resulting supernatant contained the nuclear fraction. Protein concentration was quantified using Cayman's Protein Determination Kit (Item No.704002). The presence of the cytosolic reference protein β-tubulin or β-actin in the cytosolic but not the nuclear fraction was used to confirm successful fractionation [23].

NF-κB transcription factor activity assay
After the respective treatments, nuclear and cytosolic proteins were isolated as described earlier. Binding of NF-κB in the nuclear extract to the NF-κB response element was detected with NF-κB p65 Transcription Factor Assay Kit (Cayman, Item No.10007889). Briefly, 10 μl of nuclear extract was added to wells coated with a specific double-stranded DNA sequence. Next, 90 μl of complete transcription factor buffer was added to blank wells, a positive control, non-specific binding wells, and competitor dsDNA wells. After overnight incubation at 4 • C, wells were washed five times and incubated with NF-κB (p65) antibody (except for blanks) for 1 h at room temperature. Wells were subsequently washed five times and incubated with an HRP-conjugated secondary antibody (except for blanks) for 1 h at room temperature. Each well was subsequently washed, and 100 μl of developing solution was added for 45-min until 100 μl stop solution was added.

Measurement of intracellular ROS
The formation of intracellular ROS was measured via monitoring the increasing fluorescence of 2 ,7 -dichlorofluorescein (DCF). The cell-permeant 2 ,7 -dichlorodihydrofluorescein diacetate (H 2 DCF-DA, cat. no. D399, Thermo Fisher Scientific, Grand Island, U.S.A.) enters the cell where intracellular esterases cleave off the diacetate group. The resulting H 2 DCF is retained in the cytoplasm and oxidized to DCF by ROS. HUVECs were seeded into each well of a 96-well plate and allowed to attach overnight. Subsequently, the cells were washed once with HBSS and treated with 50 nM sPRR-His. After the incubation for different time periods from 30 min to 3 h, cells were washed with HBSS, loaded with 50 μM H 2 DCF-DA for 30 min and washed again with HBSS. After a further incubation in HBSS for 1 h the fluorescence intensities were measured at 492-495 nm excitation and 517-527 nm emission in the fluorescence reader (TECAN infinite M200 PRO, TECAN, Switzerland) [28].

Measurement of the intracellular free calcium concentration (Ca 2+ )
The intracellular Ca 2+ concentration was measured using Fluo-3-AM (Abcam, ab145254). HUVECs were washed three times with PBS and incubated with 10 μM Fluo-3-AM for 30 min at 37 • C in 5% CO 2 . After proper rinsing, Ca 2+ fluorescence was recorded and quantified by Automatic living cell imaging analysis system (BioTek lionheart FX). Images were captured every 10 s after sPRR-His stimulation [27].

Animal care
All animal protocols were approved by the Animal Care and Use Committee at the University of Utah. All animal work took place in the Tianxin Yang's laboratory at University of Utah. All procedures conform to the NIH Guide for the Care and Use of Laboratory Animals. All animals were cage-housed and maintained in a temperature-controlled room with a 12:12-h light-dark cycle, with free access to tap water and standard chow. The euthanasia for animals was done by inhalant isoflurane with anesthetic vaporizer.

Animal model
Male C57/BL6 mice were purchased from Taconic (Rensselaer, New York). At one month of age, animals were placed on a high-fat diet for 9 months, i.e. 60% kcal high fat irradiated diet (Cat#D12492, Research Diets, New Brunswick, NJ). At 33 weeks of high fat diet, all animals were anesthetized with isoflurane by using inhalation anesthesia machine (induction 3-5% in O 2 , maintenance 1.5-2% in O 2 ), radiotelemetric devices (DSI) were impanated through catheterization of left carotid artery and they were allowed to recover for 1 week. After collection of baseline data for 2 days, the mice were infused with sPRR-His at 30 μg/kg/d via jugular vein catheterization connected to a separate osmotic minipump (Alzet model 1002, Alza, Palo Alto, CA) alone or combined treatment with subcutaneous infusion of Losartan at 5 mg/kg/d or Captopril at 50 mg/kg/d [29][30][31] via another minipump for 14 days.

Vascular reactivity
Twenty-four hours after collecting blood pressure and HR data for 15 days, mice were anesthetized using 2-5% isoflurane, the chest was opened, and the heart was excised. Mesenteric arteries were dissected free from adherent tissue, and their reactivity were evaluated ex vivo using isometric procedures.
Segments of mesenteric artery (∼150 μm, internal diameter) were mounted on a wire myograph system while immersed in iced NPSS. After a 30-min equilibration period wherein the vessel chamber was heated to 37 • C, 100 mg tension was placed on the vessel at 10-min intervals until 400 mg tension was achieved. Next, a series of internal circumference-active tension curves was completed to determine the vessel diameter that evoked the greatest tension development (L max ) to 100 mM potassium chloride (KCl). Four dose-response curves then were performed, each separated by 30-min. Non-receptor mediated and receptor mediated vasoconstrictive responses to KCl (20-100 mM) and phenylephrine (PE, 10 −8 -10 −5 M), respectively, were assessed. After arteries were pre-contracted to ∼65% of maximal PE-induced contraction and tension was stable, responses to acetylcholine (ACh, 10 −8 -10 −6 M; to determine endothelium-dependent vasorelaxation) and sodium nitroprusside (SNP, 10 −9 -10 −4 M; to determine endothelium-independent vasorelaxation) were evaluated. All tension data were recorded continuously using an analog-to-digital interface card (Biopac Systems Inc., Santa Barbara, CA) that allowed for subsequent off-line quantitative analyses. Vascular reactivity was assessed in two segments of mesenteric artery per mouse per group [32][33][34].

Statistical analyses
Data are presented as mean + − standard error of the mean. Significance was accepted when P<0.05. A test to determine normality of distribution for each data set was performed by GraphPad Prism software. For comparison among three or more mean values, if the data were distributed normally, a one-way ANOVA was performed to determine whether significant differences exist among groups. If significance was obtained, a Tukey's post hoc test was used to identify the location of the differences. If the data were not distributed normally, a Kruskall-Wallis ANOVA was performed. If significance was obtained, a Dunn's post hoc test was used to identify the location of the differences. For comparison among two mean values, a paired or unpaired t-test was performed as appropriate [28,35].

sPRR-His activates NF-κB in HUVECs
In primary HUVECs, sPRR-His reduced IκBα expression (Supplementary Figure S1A), increased NF-κB p-p65 protein (Supplementary Figure S1B) and elevated nuclear NF-κB p65 transcription factor activity (Supplementary Figure  S1C) at 2 and 3 h. Each response was maximal at 2 h. Our ability to successfully separate the cytosolic and nuclear fractions is shown in Supplementary Figure S2A. Immunofluorescence staining supported these data, i.e. 50 nM sPRR-His activated NF-κB p65 maximally at 2 h (Supplementary Figure S1D).

sPRR-His activates NF-κB secondary to increasing intracellular Nox4-derived H 2 O 2
The formation of intracellular ROS was measured via monitoring the increasing fluorescence of 2 7 -dichlorofluorescein (DCF), and results showed that sPRR-His increased intracellular ROS generation at 30-min (Supplementary Figure S2C). Subsequently, this result has been validated by using an independent method, e.g. the ROS-Glo H 2 O 2 assay for measurement of H 2 O 2 (Supplementary Figure S2D). Evidence exists that ROS are capable of NF-κB activation [36]. Therefore, we investigated whether sPRR-His leads to NF-κB activation by increasing cellular H 2 O 2 . In support of this notion, the important role of ROS was confirmed by findings that exogenous H 2 O 2 recapitulated sPRR-His treatment concerning IκBα degradation and NF-κB p-p65 up-regulation (Supplementary Figure S3D). This is an evidence that H 2 O 2 is an important mediator of sPRR-His-induced NF-κB activation in HUVECs.
We next sought to determine the source of sPRR-His evoked H 2 O 2 production. Nox4 is an NADPH oxidase subunit capable of generating H 2 O 2 in endothelial cells [15]. sPRR-His only elevated Nox4 mRNA and protein expression at 1, 2 and 3 h, with a maximal response observed at 2 h (Supplementary Figure S3A and B). A pharmacological and genetic approach were used next to assess the involvement of Nox4 in sPRR-His-induced H 2 O 2 generation and NF-κB activation. The dual Nox1 and Nox4 inhibitor GKT137892 (GKT) attenuated sPRR-His evoked H 2 O 2 generation (Supplementary Figure S3C), IκBα degradation and NF-κB p-p65 up-regulation (Supplementary Figure S3D). Further, HU-VECs with efficacious knockdown of Nox4 mRNA (Supplementary Figure S2B) were refractory to sPRR-His-induced H 2 O 2 generation (Supplementary Figure S3E), IκBα degradation and NF-κB p-p65 up-regulation (Supplementary Figure S3F). Taken together, these results indicate sPRR-His-induced H 2 O 2 generation is derived from Nox4 and is capable of activating NF-κB.

sPRR-His evokes inflammation
We sought to determine whether sPRR-His activates NF-κB to an extent that evokes inflammation in HUVECs. sPRR-His increased IL-6, IL-8, VCAM-1 and ICAM-1 mRNA expression levels at 2, 3 and 6 h (Supplementary Figure  S4A). The maximal response was observed at 3 h, remained elevated at 6 h, but decreased to basal levels by 9 and 12 h (Supplementary Figure S4A). Moreover, sPRR-His dose-dependently evoked inflammation, the noticeable changes of cytokine expression were observed at as low as 1 nM and maximal responses were seen at 50 nM (Supplementary Figure S4B). Importantly, sPRR-His evoked mRNA indices of inflammation after 3-h treatment were attenuated by the dual Nox1 and Nox4 inhibitor GKT, Nox4 siRNA and NF-κB inhibition using PDTC (Supplementary Figure S4C-E). Likewise, while sPRR-His elevated protein indices of inflammation, i.e. IL-6 and IL-8 expression, these responses were inhibited by Nox1/4 inhibition, Nox4 siRNA and NF-κB blockade (Supplementary Figure S5A-C). Collectively, sPRR-His evokes inflammation that is secondary to Nox4/H 2 O 2 /NF-κB activation in HUVECs.

sPRR-His increases apoptosis
Oxidative stress contributes importantly to apoptosis [37] and this was evaluated in the context of the current experimental conditions using two approaches. sPRR-His elevated the protein abundance of cleaved caspase-3 at 3 h (Supplementary Figure S6A), a response that was recapitulated by exogenous H 2 O 2 (Supplementary Figure S6B). Of note, the ability of sPRR-His to increase cleaved caspase-3 was attenuated by GKT, siRNA-mediated knockdown of Nox4 and PDTC (Supplementary Figure S6B and D). These findings were substantiated by quantitative analysis of apoptosis using flow cytometry. In this regard, sPRR-His increased the apoptotic population of cells at 3 h, and this result was reiterated by administering exogenous H 2 O 2 (Supplementary Figure S7A). Importantly, sPRR-His was unable to evoke apoptosis in cells transfected with Nox4 siRNA, or pharmacological inhibition of Nox1/Nox4 and NF-κB (Supplementary Figure S7 A-C). Quantitative analysis of the percentage of total apoptotic cells is shown in Supplementary Figure S8. These results support the statement that sPRR-His increases apoptosis by activating the Nox4/H 2 O 2 /NF-κB pathway in HUVECs.

The AT1R mediates the functions of sPRR-His
To this point our data indicate that sPRR-His evokes inflammation and apoptosis by activating the Nox4/H 2 O 2 /NF-κB pathway. The neutralizing antibody against sPRR (Ab) markedly attenuated sPRR-His-evoked Nox4 activation, IκBα Statistical significance was determined by using one-way ANOVA with the Bonferroni test for multiple comparisons. *P<0.01, compared with the CTR or Scr siRNA; #P<0.01, compared with the sPRR-His or sPRR-His+Scr siRNA. Data are mean + − SEM.
degradation, NF-κB p-p65 up-regulation and cleaved caspase-3 up-regulation (Supplementary Figure S9A), confirming specificity of the action of sPRR-His. We next explored whether these responses are secondary to sPRR binding to the AT1R. First we observed that AT1R blockade using Los ( Figure 1A,B) or AT1R siRNA knockdown ( Figure 1C,D) attenuated sPRR-His evoked Nox4 activation, H 2 O 2 generation, IκBα degradation, NF-κB p-p65 up-regulation and cleaved caspase-3 up-regulation. Second, sPRR-His elevated mRNA (Figure 2A,B) and protein ( Figure 2C,D) indices of inflammation in the absence but not the presence of Los or AT1R siRNA knockdown. Finally, quantitative analyses using flow cytometry indicated that Los inhibited sPRR-His-induced apoptosis ( Figure 2E,F). These results support the statement that sPRR-His-induced inflammation and apoptosis are secondary to AT1R activation in HUVECs.

sPRR-His directly interacts with AT1R
We next sought to elucidate the mechanism whereby sPRR-His activates the AT1R. Ang II was assessed in cell medium before and after HUVECs were treated with sPRR-His. Because Ang II concentrations were refractory to sPRR-His treatment ( Figure 3A), we suspected that sPRR-His might act in a manner that was independent of the canonical RAS. As such, we explored whether sPRR-His directly interacts with the AT1R. To test this, Ang II-I 125 radioactive ligand competitive binding assays were completed in the presence or absence of different concentrations of sPRR-His  with or without Los, to determine the amount (i.e., counts per min; CPM) of Ang II-I 125 that combined with cellular AT1R in each condition. Notably, sPRR-His binding to the AT1R out-competed Ang II-I 125 binding to the AT1R in a dose-dependent manner that was equivalent to Los at 50 nM ( Figure 3B). These data indicate that sPRR-His is a competitive ligand of Ang II. To strengthen these findings, we conducted 35 S-sPRR radioactive ligand competitive binding assays in the presence or absence of Los. Findings indicate that Los dose-dependently inhibits the uptake of 35 S-sPRR by cultured HUVECs with a maximal effect at 20 μM ( Figure 3C). Subsequently, a direct protein-protein interaction was examined by radioisotope labeling coupled with immunoprecipitation. 35 S-AT1R or 35 S-sPRR was generated by in vitro translation in the presence of 35 S-methionine. 35 S-AT1R was incubated with sPRR-His, followed by immunoprecipitation with each of the three anti-PRR antibodies. Autoradiography revealed a 35 S-AT1R band with each of the three anti-PRR antibodies but the absence of a band with IgG control ( Figure 3D). Similarly, the interaction of 35 S-sPRR and full-length AT1R protein was confirmed following anti-AT1R antibody-mediated immunoprecipitation and autoradiography analysis of 35 S-sPRR ( Figure 3D).  We further attempted to identify potential amino acid residues involved in AT1R binding with the sPRR. Although multiple amino acid residues in the transmembrane domain of AT1R constitute Los binding site, the involvement of Serine 109 and Asparagine 295 is relatively better defined [38]. Similarly, Lysine 199 of AT1R is heavily involved in interaction with Ang II [38]. To this end, we conducted mutagenesis of each of these three residues of 35 S-AT1R to Alanine and examined the consequences via immunoprecipitation coupled with autoradiography. 35 S-AT1R S109A precipitated with anti-PRR antibody showed similar band intensity on autoradiography as wild-type 35 S-AT1R. In contrast, the band intensity of immunoprecipitated 35 S-AT1R L199A or 35 S-AT1R N295A was significantly reduced ( Figure 3E,F). In addition, no 35 S-AT1R and its mutants were precipitated with an IgG control. These results suggest that Lysine 199 and Asparagine 295 but not Serine 109 are required for the protein-protein interaction between AT1R and sPRR.

sPRR-His directly activates AT1R-Gq signaling
Next we explored the downstream signaling events subsequent to sPRR-His AT1R binding in HUVECs. sPRR-His phosphorylates ERK1/2 and PKC rapidly (i.e., within 10-min), and activates intracellular Ca 2+ signaling within 90 s, and all responses are sensitive to Los or AT1R siRNA knockdown (Supplementary Figure S10A-D).

AT1R activation by sPRR-His reduces NO generation
Nox4-mediated ROS production inhibits activating phosphorylation of serine 1177 on the eNOS enzyme, i.e. p-eNOS S1177 [39]. As would be expected based on results presented thus far, sPRR-His reduced p-eNOS S1177 : total eNOS and NO generation at 3 h ( Figure 4A). Los pre-treatment ( Figure 4B,C) or AT1R siRNA-mediated knockdown ( Figure 4D,E) prevented each response, substantiating the importance of the AT1R. Taken together, these results indicate sPRR-His reduces NO generation by activating AT1R.

sPRR-His induces hypertension and endothelial dysfunction by activating AT1R in obese C57/BL6 mice
Next we sought to discern the translational relevance of our findings in HUVECs to intact animals. One-month old C57/BL6 mice consumed high-fat chow for 36 weeks to produce diet-induced obesity (DIO). At 33 weeks of feeding, radiotelemetry devices were implanted for the eventual measure of blood pressure and heart rate (HR) in conscious mice. At 34 weeks of feeding, mice were subdivided to receive sPRR-His (DIO+sPRR-His) or sPRR-His+Los (DIO+sPRR-His+Los) for the last 2 weeks while blood pressure and HR were assessed. High fat diet induced an increase in plasma sPRR levels from 3.86 to 6.16 ng/ml in C57/BL6 mice (P<0.01, N = 5-10 per group). Administering sPRR-His increased circulating sPRR concentrations from 6.16 to 13.38 ng/ml in obese C57/BL6 mice (P<0.01, N = 5-10 per group). The mean arterial pressure (MAP), systolic blood pressure (SBP) and diastolic blood pressure (DBP) were higher in DIO+sPRR-His versus DIO+sPRR-His+Los treated mice ( Figure 5A-C). As would be expected in mice with intact baroreceptor function, HR was lower in DIO+sPRR-His treated versus DIO+sPRR-His+Los treated animals ( Figure 5D). Moreover, we performed additional experiments to examine the effect of sPRR-His in lean C57/BL6 mice. Radiotelemetry demonstrated that sPRR-His infusion to C57/BL6 mice at the same dosage over 2 weeks didn't significantly affect blood pressure and HR (Supplementary Figure S11A-D). It is possible that sPRR-His may play a specific role in mediating obesity-induced hypertension.
After measuring blood pressure and HR for the final two weeks of the 36-weeks feeding regimen, mice were anesthetized and mesenteric arterial function was assessed. We sought to determine if hypertension observed in DIO+ sPRR-His treated mice was secondary to endothelial dysfunction, and whether AT1R blockade might be protective. Acetylcholine-evoked vasorelaxation was greater in mesenteric arteries from Los-treated versus sPRR-His-treated DIO mice, while responses to sodium nitroprusside were similar among groups ( Figure 6A,B). Moreover, acetylcholine-evoked vasorelaxation was similar in arteries from Los-treated vs. sPRR-His-treated lean mice, and responses to sodium nitroprusside were not different among groups ( Figure 6C,D). However, vascular smooth muscle responses were similar among groups (Supplementary Figure S12A-D). These data suggest that endothelial function is preserved by AT1R blockade, and it is not unreasonable to speculate that this contributed importantly to normalizing arterial blood pressure.

ACE inhibition using captopril does not affect sPRR-His responses in vitro or in vivo
Concern might be raised that sPRR-His facilitates Ang II generation, which may mediate the action of sPRR-His. To address this, experiments were completed using the angiotensin-converting enzyme inhibitor (ACEi) captopril (Cap). Cap did not attenuate sPRR-His-evoked Nox4 activation, IκBα degradation, NF-κB p-p65 up-regulation and cleaved caspase-3 up-regulation in HUVECs (Supplementary Figure S9B). Furthermore, radiotelemetry data concerning arterial blood pressure indicated that Cap did not significantly affect sPRR-His-induced hypertension in DIO mice ( Figure 5E-H). In order to test the effective dosage of Cap, we measured plasma Ang II and bradykinin levels by ELISA. The plasma Ang II level was significantly reduced from 7462 to 2492 pg/ml in DIO+sPRR-His versus DIO+sPRR-His+Cap treated mice (P<0.01, N = 5 per group), and the plasma bradykinin level was increased from 31.61 to 61.26 pg/ml in DIO+sPRR-His versus DIO+sPRR-His+Cap treated mice (P<0.01, N = 5 per group), confirming the effectiveness of Cap. Together, these data suggest that the prohypertensive action of sPRR-His infusion in DIO mice is mediated by direct activation of AT1R, independent of generation of Ang II.

Discussion
For many decades AT1R activation was thought to completely depend on its sole physiological ligand, Ang II. Findings from the present study challenge this notion. Here, we report for the first time that sPRR directly binds to and activates the AT1R in an Ang II-independent manner to suppress NO generation by endothelial cells and we provide compelling evidence for a novel mechanism that is responsible. In addition, we provide important translational evidence concerning the (patho) physiological relevance of this signaling pathway in the context of hypertension and arterial dysfunction that exists in obese mice. To our knowledge no prior studies have reported the biological function of sPRR in the vasculature or its signaling through AT1R. We believe AT1R mediates the functions of sPRR during and IL-6 and IL-8 protein expression. These data provide solid evidence that elevated sPRR-His promotes inflammation/adhesion in endothelial cells. Inflammation plays an important role in the development and pathophysiology of hypertension [43]. Substantial evidence exists that Ang II induces inflammation and apoptosis by activating the AT1R/NADPH/ROS/NF-κB pathway [44,45]. These signaling events are similar to those we report here for sPRR-His. Moreover, the preliminary evidence suggests that AT1R and PRR can form a heterodimer that is functionally active to enhance ERK phosphorylation in PC12W cells [46]. These findings provided rationale for our exploration into whether the AT1R directly mediates the signaling events precipitated by elevated sPRR. Of significant interest to us, results from immunoprecipitation experiments and radioactive ligand competitive binding assays indicated that sPRR directly interacts with the AT1R. In particular, the direct protein-protein interaction between sPRR and AT1R was validated in a cell-free system by immunoprecipitation coupled with autoradiography. Furthermore, site-directed mutagenesis was conducted to map the binding site. The key amino acid residues of AT1R are Serine 109 and Asparagine 295 for Los binding and Lysine 199 for Ang II binding [38,47]. We found that mutation of Lysine 199 and Asparagine 295 but not Serine 109 significantly reduced sPRR binding to the AT1R. Therefore, sPRR may share partial or complete binding sites of AT1R with Ang II or Los. In addition to Lysine 199 , other residues including Tryptophan 253 , Phenylalanine 259 , Asparagine 263 , and Arginine 167 also interact with Ang II [38,47] but their involvement concerning sPRR binding has not been investigated. What's more, we observed that sPRR-His stimulated AT1R mediated Gq signaling, confirming that sPRR can directly activate AT1R. The current study is the first to report the direct interaction between sPRR and AT1R in general, and in endothelial cells in particular.
To determine whether our findings in vitro are relevant to the intact animal, we studied the effects of sPRR-His on vascular function and blood pressure in obese mice. Results using isometric tension techniques revealed that endothelium-dependent vasodilation displayed by arteries from obese mice treated with sPRR-His was improved by Los. Because vascular smooth muscle responses were similar between groups, it is not unreasonable to suggest that Los improved an endothelium-dependent defect evoked by sPRR-His. Endothelial dysfunction plays an important role in the development and pathophysiology of hypertension [48]. Indeed, we observed elevated blood pressure in obese mice treated with sPRR-His, an effect that was reversed by concurrent treatment with Los but not ACEi using Cap. The distinct effects of Los versus Cap strongly support that sPRR induces hypertension via direct activation of AT1R as opposed to the generation of Ang II. Cap significantly reduced plasma Ang II level and increased plasma bradykinin level in DIO+sPRR-His treated mice, confirming the effectiveness of ACE inhibition. While captopril effectively inhibited the production of Ang II, the radiotelemetry data showed no significant effect of captopril on sPRR-His-induced hypertension in DIO mice, indicating that the increased Ang II was not the reason for sPRR-His-induced hypertension in DIO mice. Of note, in agreement with the present study, Gatineau et al. reported a similar prohypertensive action of sPRR in DIO mice [49]. However, discrepancy exists in that Gatineau et al. found no effect of losartan on the prohypertensive action of sPRR. The exact reason for the discrepancy is unclear but may be related to differences in experimental models (36 versus 21 weeks of high fat diet) and protocols. It might be possible that sPRR-mediated activation of AT1R in the vasculature may predominate during prolonged high fat diet treatment whereas baroreflex dysfunction may be relatively more important during regular high fat diet treatment [49]. Indeed, the mechanism of obesity-related hypertension is complex, involving endothelial dysfunction, Na + retention, increased leptin levels and baroreflex dysfunction [50]. Relative importance of each of the underlying mechanisms may vary at various stages of the hypertension development [51,52].
We have examined the property of sPRR-His in cultured vascular smooth muscle cells as well as isolated mesentery arteries from the normal control lean mouse. To our surprise, sPRR-His did not induce activation of AT1R signaling pathway such as Gq (data not shown). In agreement with this observation, while potent Ang II-induced vasoconstriction was observed in mesenteric arteries as examined ex vivo, sPRR-His was without effect (Supplementary Figure  S13). It is likely that within the vasculature sPRR-His acts specifically on endothelial but not vascular smooth muscle cells and such specificity may be determined by some unknown co-receptor (s) rather than AT1R. An example is that fibroblast growth factor 23 (FGF-23) via its receptor FGFR acts in a wide variety of tissues and its specific action in the kidney for regulation of tubular phosphate transport is rather determined by its co-receptor Klotho.
The AT1R is a core member in the RAS that promotes myriad signaling pathways contributing to the pathogenesis of cardiovascular diseases [53]. In the classic sense, Ang II has been considered as the exclusive ligand for the AT1R. Here, we provide compelling in vitro and in vivo evidence that sPRR serves as an alternative activator of the AT1R, and that activation via this pathway contributes to cardiovascular complications (e.g., arterial dysfunction and hypertension) associated with diet-induced obesity. Taken together, our results provide solid support for sPRR to be considered further as a predictive marker and/or therapeutic target for cardiovascular related pathologies.

Clinical perspectives
• Obesity is a major risk factor of endothelial dysfunction and hypertension with unclear mechanism.
• The present study for the first time demonstrates that sPRR directly interacts with AT1R to lead to NF-κB activation, inflammation, apoptosis, and attenuated p-eNOS S1177 in primary HUVECs. Besides, the interaction between sPRR and AT1R contribute to endothelial dysfunction and hypertension in DIO mice.
• These findings offer a novel mechanism of endothelial dysfunction and hypertension associated with obesity. sPRR can be considered further as a predictive marker and/or therapeutic target for cardiovascular disorders associated with endothelial dysfunction.