Vascular Pharmacology 63 (2014) 88–96

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Vascular Pharmacology journal homepage: www.elsevier.com/locate/vph

Notch signaling governs phenotypic modulation of smooth muscle cells Cho-Hao Lin, Brenda Lilly ⁎ Nationwide Children's Hospital, The Heart Center, Department of Pediatrics, The Ohio State University, Columbus, OH, United States

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Article history: Received 5 May 2014 Received in revised form 22 August 2014 Accepted 16 September 2014 Available online 23 October 2014 Keywords: Smooth muscle cells Endothelial cells Notch signaling Coculture Phenotypic modulation

a b s t r a c t A feature of vascular smooth muscle cells is their unique ability to exist in multiple phenotypes permitting a broad range of functions that include contraction, proliferation, or synthesis and secretion of extracellular matrix. Although it is known that these phenotypes can be overlapping, the mechanisms that regulate phenotypic modulation are still unclear. Given that endothelial cells are known to convey signals to smooth muscle cells that govern their activities within the vasculature; we sought to better define how endothelial cells regulate phenotypic changes of smooth muscle cells in coculture conditions. Using human aortic smooth muscle cells, we show that endothelial cells promote an increase in a differentiated/contractile phenotype while decreasing proliferation. Analysis of the synthetic phenotype demonstrates that endothelial cells also increase collagen synthesis and secretion. Characterization of pathways important for these endothelial cell-dependent phenotypes reveal that Notch signaling plays an important role in the establishment of these smooth muscle properties. These data highlight the ability of endothelial cells to control phenotypic modulation in a unique and previously undefined manner. © 2014 Elsevier Inc. All rights reserved.

1. Introduction Phenotypic modulation is a hallmark of smooth muscle cells, and although it is well described in vascular injury and with in vitro models, the mechanism of its regulation is poorly understood. In particular, smooth muscle cell phenotypes are considered mutually exclusive, however careful characterization of these cells demonstrated that they exist in a range of overlapping phenotypes [21,23,26]. This ability to exist in between or in shared phenotypes seems a valuable strategy for smooth muscle cells to perform a multitude of functions throughout their lifespan. The major role of mature differentiated vascular smooth muscle cells is to maintain blood vessel tone and to regulate blood pressure through constriction or relaxation. This is achieved through the expression of a complement of regulatory and contractile genes that provide the machinery for this response [21,22]. The differentiated contractile phenotype is largely characterized by expression of coordinately regulated smooth muscle-specific markers that include smooth muscle (SM) α-actin (ACTA2), smooth muscle myosin heavy chain (MYH11), Calponin (CNN1) and SM22α (TAGLN) [21,22]. But adult smooth muscle cells exhibit a plasticity, which allows them to undergo extensive phenotypic changes in response to environmental cues and vascular injury [22,25]. For example, smooth muscle cells lose the expression of smooth muscle-specific contractile proteins and their quiescence, and produce excessive extracellular matrix (ECM) proteins DOI of original article: http://dx.doi.org/10.1016/j.vph.2014.10.003. ⁎ Corresponding author at: The Research Institute at Nationwide Children's Hospital, 700 Children's Drive, Columbus, OH 43205, United States. Tel.: + 1 614 355 5750; fax: +1 614 355 5725. E-mail address: [email protected] (B. Lilly).

http://dx.doi.org/10.1016/j.vph.2014.09.004 1537-1891/© 2014 Elsevier Inc. All rights reserved.

in atherosclerosis or vessel stenosis [12]. This synthetic phenotype is classically characterized by an increase in the proliferation index together with an increase in rough endoplasmic reticulum, golgi, and ribosomes that facilitate the ECM production and secretion [7,26]. The proliferative and synthetic phenotypes are strongly associated in vessel pathology, but these particular smooth muscle characteristics are considered distinct, and can be found independent from one another [7,22]. The current belief is that smooth muscle cells exist in a continuum somewhere between the extremes of these defined phenotypes, and the surrounding environment dictates their current state. During blood vessel development, smooth muscle cells contribute to the structural integrity of the blood vessel by proliferating and supplying ECM components for the vessel wall and basement membrane [8,9]. These same features are also essential in adult blood vessels for repair and remodeling associated with maintenance and growth of the normal vasculature. Several studies have identified signaling pathways and genetic factors that are able to govern phenotypic modulation of smooth muscle cells [2,22,23]. Yet, many mysteries remain as to the triggers that promote these phenotypic transitions in normal and diseased blood vessels. Developmental studies from our lab have focused on the interactions of endothelial and smooth muscle cells in vessel formation, and from these and previous studies by other groups it has become apparent that endothelial cells have a substantial influence on smooth muscle cell differentiation [6,8,18]. Moreover, the importance of endothelial cells in vascular injury has been well described; where disruptions or removal of the endothelial cell monolayer results in the loss of smooth muscle contractile proteins and a switch to a highly proliferative and synthetic phenotype [12,27]. Thus, the influence of endothelial cells on smooth

C.-H. Lin, B. Lilly / Vascular Pharmacology 63 (2014) 88–96

2. Materials and methods

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HAoSMC HAoSMC/HPAEC 48hr HAoSMC/HPAEC 96hr

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Primary cultures of human aortic smooth muscle cells (HAoSMCs) were purchased from Vasculife and maintained in DMEM (Thermo Fisher Scientific) supplemented with 5% FBS (HyClone), insulin (4 ng/ml), EGF (5 ng/ml), ascorbic acid (50 ng/ml), 2 mM glutamine, 1 mM sodium pyruvate, and 100 units/ml penicillin/streptomycin. Primary cells between passages 6 and 9 were used for all experiments. Human umbilical vein endothelial cells (HUVEC, Cascade Biologics), human microvascular endothelial cells (HMVEC, Lonza) and human pulmonary artery endothelial cells (HPAEC, Lifeline) were grown in EBM-2 supplemented with the BulletKit components (Lonza) as recommended by the manufacturer. Endothelial cells between passages 7 and 11 were used for all

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HAoSMC HAoSMC/HMVEC 48hr HAoSMC/HMVEC 96hr

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muscle cell phenotypes appears to be a primary mediator of phenotypic modulation. Extending this notion further, others and we have demonstrated an important role of Notch signaling between endothelial cells and smooth muscle cells for smooth muscle differentiation [3,13,18]. Four mammalian Notch receptors (Notch1-4) and five transmembrane ligands (Jagged1,2, Delta-like (Dll-1,3,4)) are expressed in vascular cells [1,11]. Studies by High et al., demonstrated that loss of Jagged1 on endothelial cells results in an absence of vascular smooth muscle differentiation [13]. In this study we show that endothelial cells impart a unique phenotype to smooth muscle cells. Cocultured endothelial cells promote a quiescent and contractile phenotype, and additionally activate a synthetic phenotype in smooth muscle cells. Our data demonstrate that Notch signaling is central to endothelial cell-dependent phenotypic modulation and reveal that NOTCH2 and NOTCH3 play critical roles in this regulation.

HAoSMC HAoSMC/HUVEC 48hr HAoSMC/HUVEC 96hr

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Fig. 1. Arterial, venous and microvascular endothelial cells promote smooth muscle differentiation. (A–C) Human aortic smooth muscle cells (HAoSMCs) were cultured with human umbilical vein endothelial cells (HUVEC), human microvascular endothelial cells (HMVEC), or human pulmonary artery endothelial cells (HPAEC) for 48 or 96 h. After being separated using anti-PECAM1-conjugated magnetic beads, mRNA was isolated from smooth muscle cells and measured by qPCR. (D) Smooth muscle specific protein expression was detected by immunoblotting following coculture and separation from endothelial cells as described. (E–G) Quantified protein expression of smooth muscle markers normalized to TUBULIN (TUBB2A). *P b 0.05 compared to control cells cultured alone.

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experiments. Human adenocarcinoma (HeLa) cells were purchased from American Type Culture Collection and cultured in DMEM supplemented as indicated with 5% FBS. For virus production, TN-293 cells were purchased from Stratagene and cultured in DMEM with 5% FBS, 2 mM glutamine and 1 mM sodium pyruvate. All cultures were maintained in humidified 5% CO2 at 37 °C. For coculture, 1.25 × 105 HAoSMC was plated in 6-well plates, and after adhesion, 1.25 × 105 endothelial cells were added. N-[(3,5-Difluorophenyl)acetyl]-L-alanyl2-phenylglycine 1, 1-dimethylethyl ester (DAPT; Calbiochem) was added to specified wells at the time of plating. To separate endothelial cells from smooth muscle cells, anti-PECAM1 conjugated Dynabeads (Invitrogen) were used following the manufacturer's instructions. Cell separation was determined to be greater than 95% efficient [16]. All cell coculture experiments were performed in medium consisting of EBM-2 supplemented with all BulletKit components. The separated cells were then processed for quantitative RT-PCR (qPCR) or Western blot analysis. 2.2. Quantitative RT-PCR (qPCR) Total RNA was isolated from cells using RiboZol reagent (Amersco) according to the manufacturer's instructions and reverse transcribed with M-MLV reverse transcriptase (Promega) to generate cDNA. Real-time PCR was performed using a StepOne PCR system (Applied Biosystems) with SYBR Green and 50 ng of cDNA as template. The fold difference in target gene mRNA levels was calculated using the ΔΔCT method and normalized to GAPDH mRNA from the same sample. Primer sequences were as follows: NOTCH3, 5′- GAG CCA ATG CCA ACT GAA GAG (forward) and 5′- GGC AGA TCA GGT CGG AGA TG (reverse); NOTCH2, 5′ - ACA GTT GTG TCT GCT CAC CAG GAT (forward) and 5′ -

GCG GAA ACC ATT CAC ACC GTT GAT (reverse); HEYL, 5′- CAT ACA ATG TCC TTG TGC AGT ACA CA (forward) and 5′- GCC AGG GCT CGG GCA TCA AAG AA (reverse); SMOOTH MUSCLE α-ACTIN (ACTA2), 5′CAA GTG ATC ACC ATC GGA AAT G (forward) and 5′- GAC TCC ATC CCG ATG AAG GA (reverse); SM22α (TAGLN), 5′- CAA GCT GGT GAA CAG CCT GTA C (forward) and 5′- GAC CAT GGA GGG TGG GTT CT (reverse); CALPONIN (CNN1), 5′- TGA AGC CCC ACG ACA TTT TT (forward) and 5′- GGG TGG ACT GCA CCT GTG TA (reverse); COL1A1, 5′- CAG ACA AGC AAC CCA AAC TGA A (forward) and TGA GAG ATG AAT GCA AAG GAA AAA (reverse); COL3A1, 5′- TGG TCA GTC CTA TGC GGA TAG A (forward) and 5′- CGG ATC CTG AGT CAC AGA CAC A (reverse); COL4A1, 5′CGT AAC TAA CAC ACC CTG CTT CAT (forward) and 5′- CAC TAT TGA AAG CTT ATC GCT GTC TT (reverse); COL5A3, 5′- GAC AGA GAC TCC AGC TCC AAA TC (forward) and 5′- TCT CTA GGA TCG TGG CAT TGA G (reverse); CYCLIND1 (CCND1), 5′- CGT GGC CTC TAA GAT GAA GGA (forward) and 5′- CGG TGT AGA TGC ACA GCT TCT C (reverse); CYCLIND2 (CCND2), 5′- CCC TCT GCT GAG CGG TAC TAA (forward) and 5′- TCT TAT CCT GCC AAT TCA GTG TGA (reverse); CALD1, 5′- ACC AGG AGA CGT ATC CAG CAA (forward) and 5′- GGG AAG TGA CCT TAT CCA CAG ATT (reverse); RBP1, 5′- GTG GCC TTG CGC AAA ATC (forward) and 5′CCG TCC TGC ACG ATC TCT TT (reverse); VIMENTIN (VIM), 5′- AAT GAC CGC TTC GCC AAC T (forward) and 5′- ATC TTA TTC TGC TGC TCC AGG (reverse); GAPDH, 5′- ATG GAA ATC CCA TCA CCA TCT T (forward) and 5′- CGC CCC ACT TGA TTT TGG (reverse). 2.3. Western blotting Equivalent amounts of protein from whole cell lysates were run on 10% SDS-polyacrylamide gels. Proteins were transferred to nitrocellulose membranes (GE Healthcare), blocked using 3% nonfat dry milk

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Fig. 2. Endothelial cells increase smooth muscle cell actin filaments and contractile ability. (A) HAoSMC prestained with a tracker dye (green) were cultured with HUVEC for 48 h and counterstained with phalloidin (red) to highlight actin filaments and DAPI (nuclei, blue). Scale bar = 50 μm. (B) Tracker dye-stained (green) HAoSMCs were immunostained to detect ACTA2 or CNN1 (red). Scale bar = 20 μM. (C) Smooth muscle cells were cultured alone or cocultured with HUVEC or HeLa adenocarcinoma in a collagen I gel. HeLa and HUVEC were cultured alone as controls. The gel images were captured immediately after gel release and one hour later. (D) Quantification of gel area was measured by ImageJ software. *P b 0.05, n.s., not significant.

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and subjected to incubation using primary antibodies to Notch3 (sc5593, Santa Cruz Biotechnology), Notch2 (C651.6DbHN, DSHB) βtubulin (TUBB2A;T7816, Sigma), smooth muscle α-actin (ACTA2;1A4, Sigma), smooth muscle myosin heavy chain (MYH11;BT562, Biomedical Technologies Inc.), Calponin (CNN1;C2687, Sigma), and Vimentin (VIM;V5255, Sigma). Secondary antibodies conjugated to HRP (Amersham Biosciences) were incubated for 2 h at room temperate and used for detection. Protein was detected by enhanced chemiluminescence (Thermo Fisher Scientific). β-Tubulin was used to assess total amount of protein in each sample. 2.4. Gel contractile assay For contractile assay, 70 μl of rat tail collagen I suspension (1 mg/ml) supplied with 1% FBS containing the 2.8 × 104 smooth muscle cells, 2.8 × 104 endothelial cells or 2.8 × 104 HeLa cells was cast in one well of a 96-well culture plate and allowed to polymerize at 37 °C for 30 min. After collagen polymerization, 140 μl of complete EBM-2 medium was added to each collagen gel. Cultures were incubated for two days and replaced medium with fresh 10% FBS DMEM before releasing the gels from the sides of the well. The relative changes of collagen gel were measures 1 h later to quantify contraction with using NIH ImageJ software. 2.5. Immunostaining HAoSMCs were incubated with cell tracker dye Green CMFDA (Invitrogen) at 10 μM of final concentration for 15 min before coculture with HUVEC. After 96 hour co-culture, cells were fixed with 4%

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PFA for 20 min, blocked and permeabilized in PBS containing 5% goat serum, 2% Bovine serum albumin and 0.3% Triton X-100 for 1 h. Primary antibodies, Ki67 (ab66155, Abcam), ACTA2 (1A4, Sigma), CNN1 (C2687, Sigma) and Alexa Fluor 594 phalloidin (Invitrogen) were used to incubate with cells. In Ki67 staining, the percentage of Ki67 positive cells was determined by costaining with a tracker dye and determining the ratio of fluorescently labeled cells. For smooth muscle myosin heavy chain (MYH11) and Vimentin (VIM) costaining, HAoSMC where cocultured with HUVEC for 96 h. After cell separation, HAoSMCs were plated in a chamber slide for 2 h and then fixed with 4% PFA for 20 min, blocked and permeabilized in PBS containing 5% goat serum, 2% Bovine serum albumin and 0.3% Triton X-100 for 1 h. Primary antibody used MYH11 (BT562, Biomedical Technologies Inc.) and Vimentin (VIM) (V5255, Sigma). Intensity of expression was quantified using NIH ImageJ software. Fluorescently tagged secondary antibody used: Alexa-Fluor 488 goat anti-rabbit, Alexa-Fluor 568 goat anti-mouse and Alexa-Fluor 594 goat anti-rabbit. (Invitrogen). 2.6. RNA interference HAoSMCs were plated in a 12-well plate at 6 × 104 cells/well. After 24 h, cells were transfected with siRNA using RNAiMAX (Invitrogen). Notch2 siRNA was purchased from QIAGEN (GS4853) and Notch3 siRNA was synthesized by IDT as follows: 5′-AAC UGC GAA GUG AAC AUU G. Control siRNA was purchased from Invitrogen. All siRNAs were transfected at 40 nM. After 24 hour transfection, cells were cocultured with 6 × 104 HUVEC for additional 96 h and collected for qPCR analysis and Western blotting.

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Fig. 3. Proliferation is repressed with endothelial cell coculture. (A) Prior to coculture, HAoSMCs were stained with a tracker dye (green), and following coculture with HUVEC for 96 h, HAoSMCs were stained with Ki-67-specific antibody (red) and DAPI (blue). Scale bar = 100 μm. (B) The ratio of proliferating smooth muscle cells was determined by counting the number of pre-stained smooth muscle cells that were Ki-67 positive. (C) Cell cycle regulators, CYCLIND1 and CYCLIND2 mRNA expression in HAoSMC was analyzed by qPCR. *P b 0.05 compared to control cells cultured alone.

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2.7. Collagen assay 1.25 × 105 HAoSMCs were cocultured with equal number of HUVEC in a 6-well plate for 96 h. HAoSMCs were separated from coculture and re-plated in a 12-well plate at a cell density of 5 × 104 cells per well for additional 12 h. Collagen concentration in cultured medium was measure by Sirius Red assay as described [15]. Briefly, 500 μl of cultured medium was collected and incubated with 1.5 ml 25% ammonium sulfate solution for 24 h at 4 °C. 50 μM Sirius red F3B (Sigma) was used to bind the side chain groups of collagen, and then 0.1 M potassium hydroxide was added to release Sirius Red from the precipitated collagen. Colorimetric detection was performed at 540 nm absorbance using standard controls to determine collagen concentration using a Molecular Devices SpectraMax M5 microplate reader.

for additional 96 h. The cultured medium was collected and mixed with Nano-Glo reagent to measure secreted luciferase activity using a LUMIstar Omega luminometer (BMG Labtech). To normalize the transfection efficiency, Hsp-β-galactosidase (LacZ) was cotransfected, and luciferase activities were normalized based on an equivalent amount of LacZ activity. 2.9. Statistical analysis Statistical analyses were done using GraphPad Prism (GraphPad Software). Data graphed with error bars represent standard error of the mean. Student's t test and ANOVA were used to determine the significant difference if P b 0.05. Data shown are representative of at least three independent experiments.

2.8. Luciferase assay

3. Results

For Notch/CBF-Luciferase assay, CBF-luciferase plasmid was generated as described [18]. To measure the luciferase activity, HAoSMCs were plated in a 24-well plate at high (2 × 104) or low density (1 × 104) and transfected with plasmids using PolyJet (SignaGen). After 24 h, cells were co-cultured with HUVEC, or HAoSMC for another 48 h. The promoter activity was measured by luciferase assays using Bright-Glo reagent (Promega). A CMV-SEAP plasmid (Addgene plasmid 24595) was used as a control for transfection efficiency. For measuring secreted luciferase, the CAG promoter [20] was clone into a NanoLuc luciferase plasmid pNL1.3 (Promega) fused to an IL-6N-terminal secretion signal sequence [24]. To measure the secreted luciferase activity, 3 × 104 HAoSMCs were plated in each well of 24-well plate and transfected with PolyJet (SignaGen) or cotransfected with siRNA using [17] (Invitrogen). After 24 h, 3 × 104 HUVEC were co-culture with HAoSMC

3.1. Endothelial cells from unique vascular beds regulate smooth muscle differentiation

B Relative concentration of secreted collagen

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Previous studies have shown that primary cultures of smooth muscle cells that are cocultured with endothelial cells exhibit an increase in smooth muscle contractile gene expression [14,18]. To extend these analyses, we first characterized the response of smooth muscle cells to endothelial cells derived from different vascular beds. Coculture of human aortic smooth muscle cells with endothelial cells from umbilical vein, microvasculature, and the pulmonary artery exhibited a similar profile of induced gene expression. Smooth muscle cells were cocultured for 48 and 96 h then isolated from the different endothelial cells using anti-PECAM-1-conjugated magnetic beads. RNA and protein extracted from the smooth muscle cells was subjected to qPCR and

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Fig. 4. Synthetic phenotype is induced by endothelial cells. (A) Expression of collagen genes in HAoSMC was measured by qPCR after coculture for 96 h and separation from HUVEC. (B) The concentration of total collagen secreted by HAoSMC into the culture medium was measured by Sirius Red assay after coculture with HUVEC. (C) HAoSMCs were transfected with a plasmid harboring a constitutively-expressed secreted luciferase and cocultured with HUVEC for 96 h. Luciferase assays were performed with culture media to assess secretion rates. (D) Expression of synthetic markers was analyzed by qPCR. *P b 0.05 compared to control cells cultured alone.

C.-H. Lin, B. Lilly / Vascular Pharmacology 63 (2014) 88–96

Western blot, respectively. These analyses showed that endothelial cells derived from different vascular beds had a similar ability to induce smooth muscle-specific contractile gene expression (Fig. 1). Furthermore, like our previous studies, the expression of NOTCH3 was also induced (Fig. 1), indicating activation of Notch signaling is a common component in endothelial and smooth muscle cell interactions. Because we observed a similar gene activation profile using the three different cell types, we focused our subsequent analysis of smooth muscle phenotypes using HUVEC. We next asked if the increase in contractile gene expression translated into an increase in contractile response of these cocultured smooth muscle cells. Examination of the actin fiber content by phalloidin staining revealed a robust increase in muscle fibers within the cocultured smooth muscle cells that were identified by prestaining with a tracker dye (Fig. 2A). Images of the smooth muscle cells cocultured with endothelial cells revealed strong interaction between the cell types, consistent with our previous findings {Lilly, 2009 #312;Liu, 2009 #16}. Immunostaining to detect SMOOTH MUSCLE α-ACTIN (ACTA2) and h1-CALPONIN (CNN1) demonstrated an increase in expression of both of these smooth muscle marker genes (Fig. 2B). We next performed contraction assays in which smooth muscle cells and endothelial cells were cocultured in a collagen gel matrix. Compared to smooth muscle cells cultured alone or cocultured with HeLa cells, which we used as a control, smooth muscle cells cocultured with

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endothelial cells showed a strong contractile response, indicating that the increase in contractile gene expression was promoting a more functional contractile smooth muscle cell (Fig. 2C, D). 3.2. Endothelial cells promote a quiescent and synthetic phenotype The increase in contractile response together with contractile gene expression suggested that endothelial cells were promoting a differentiated and quiescent phenotype. To determine if proliferation was decreased by cocultured endothelial cells we measured proliferation by immunostaining for Ki-67 and by examination of CYCLIND1 and CYCLIND2 gene expression. The data show that proliferation of smooth muscle cells is decreased by coculture with endothelial cells (Fig. 3). Ki67-positive smooth muscle cells were decreased by 57%, and both CyclinD1 and CyclinD2 showed a substantial decrease in expression (35% and 54%, respectively). Smooth muscle cells can also transition to a synthetic phenotype that is brought on by physiological stressors in pathological states [12]. To test if endothelial cells affect the synthetic phenotype, we first measured collagen gene expression and collagen secretion. The results indicate that cocultured endothelial cells significantly increase the expression of Collagen genes 1A1, 3A1, 4A1, and 5A3, as well as collagen secretion from smooth muscles following coculture for 96 h (Fig. 4A, B). The secretion rates of established secretory cells are

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Fig. 5. Contractile and synthetic markers are co-expressed in smooth muscle cells cocultured with endothelial cells. (A) Following coculture with HUVEC for 96 h, HAoSMCs were separated, replated and were co-immunostained for expression of MYH11(green) and VIMENTIN (red). DAPI (blue) highlights nuclei. Notch signaling inhibitor DAPT was added as indicated at time of plating. Scale bar = 10 μm. (B–C) Quantification of florescence intensity of MYH11 and VIMENTIN. *P b 0.05, n.s., not significant.

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data from our lab indicated that contractile marker gene expression is blunted by inhibition of Notch signaling. Consistent with this, addition of Notch inhibitor, DAPT caused a decrease in MYH11 staining, but interestingly failed to inhibit the expression VIMENTIN (Fig. 5).

typically measured using a constitutively active secreted-luciferase plasmid [17]. Using this strategy, we measured the amount of secreted luciferase from transfected smooth muscle cells in the presence or absence of cocultured endothelial cells. These results showed a robust increase in secreted luciferase in the cocultured smooth muscle cells (Fig. 4C). Furthermore, analysis of traditional synthetic markers, CALDESMON (CALD1), RETINOL-BINDING PROTEIN-1 (RBP1), and VIMENTIN (VIM) [2,23], showed an increase in expression consistent with a synthetic phenotype (Fig. 4D). Together, these data indicate that endothelial cells cause an increase in both a contractile and synthetic phenotype, while suppressing the proliferative phenotype. The results indicated that contractile and synthetic phenotypes are both induced by cocultured endothelial cells. To determine if the smooth muscle cells that are cultured with the endothelial cells coexpress these phenotypes or exist as a mixed population of cells exhibiting one or the other of these phenotypes, we performed immunostaining with a contractile marker (MYH11) and a synthetic marker (VIMENTIN) on smooth muscle cells cultured alone or with endothelial cells. Both markers were robustly expressed in cocultured smooth muscle cells compared to the cells cultured alone, and these markers were coexpressed within the smooth muscle cells, indicating the presence of both phenotypes within a single smooth muscle cell (Fig. 5). Previous

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HAoSMC HAoSMC/HUVEC Relative mRNA expression

Relative Luciferase Activity

5

Others and we have demonstrated that Notch signaling via the Jagged1 ligand on endothelial cells contributes to the induction of Notch activity in smooth muscle cells [3,13,18]. To determine whether Notch activation in smooth muscle cells was occurring only through endothelial cell signaling, we used a Notch-sensor (5X-CBF) luciferase construct transfected into smooth muscle cells to measure activation by different cocultured cells. Compared to HAoSMC cultured alone, transfected HAoSMC cultured with endothelial cells exhibited a robust Notch response (Fig. 6A). The addition of untransfected HAoSMCs did not promote the same Notch activation. Moreover, culturing of transfected HAoSMC at either high (confluent) or low density had little effect on Notch activity, suggesting that Notch signaling is relatively low between neighboring smooth muscle cells, and requires initiation by endothelial cells.

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3.3. Notch signaling governs endothelial cell-dependent phenotypic modulation of smooth muscle cells

Relative mRNA expression

94

Fig. 6. Notch signaling plays a critical role in smooth muscle cell phenotypic modulation. (A) HAoSMCs were plated at low and high density (confluent) and transfected with a Notch sensor (CBF-Luciferase reporter). Cells were then cultured alone or HUVEC or HAoSMCs were added for 48 h, followed by luciferase assays to measure Notch activity. HAoSMC cultured alone or cocultured with HUVEC were treated with DAPT or vehicle. (B–D) mRNA expression of NOTCH3 and smooth muscle markers, Cyclin genes and collagen genes were assessed by qPCR in smooth muscle cells following cell separation. (E) Collagen secretion from smooth muscle cells into the media was measured by Sirius Red assay. (F) Rate of secretion by HAoSMC was measured using secreted luciferase assay. (G) Gene expression of synthetic markers was measured by qPCR following DAPT treatment and coculture. (H, I) Protein expression of contractile and synthetic markers was determined by immunoblotting. *P b 0.05, n.s., not significant.

C.-H. Lin, B. Lilly / Vascular Pharmacology 63 (2014) 88–96

To further explore the role of Notch signaling, we targeted two of the Notch receptors that are most abundant in smooth muscle cells, Notch2 and Notch3 [28]. The deletion of NOTCH2 and NOTCH3 by siRNA revealed that loss of either receptor was sufficient to block endothelial cell-induced smooth muscle differentiation gene expression and contractile ability (Fig. 7A–D). In contrast, knockdown of NOTCH2 revealed a specific requirement for regulation of the CYCLIN D1 and D2 genes and Ki67 within smooth muscle cells (Fig. 7E, F). The examination of the importance of the Notch receptors in the regulation of the synthetic phenotype revealed that both were required for collagen gene expression (Fig. 7G), but loss of either had no effect of the traditional synthetic marker gene expression profiles (Fig. 7H), consistent with Notch inhibition with DAPT (Fig. 6). Finally, using the luciferase secretion assay to measure cell secretion, the data show that NOTCH2 has no effect on secretion, while loss of NOTCH3 blocked the endothelial cell-induced secretion in smooth muscle cells (Fig. 7I). These data indicate that Notch signaling plays an important role in phenotypic modulation, and point to unique roles of the Notch receptors in governing these activities.

Notch signaling has been linked to smooth muscle differentiation both in vitro and in vivo [4,10,11,19]. By immunostaining for contractile/differentiation marker, MYH11, inhibition of Notch signaling selectively blocked endothelial cell-induced contractile differentiation, while having no effect on the synthetic phenotype as assessed by VIMENTIN staining (Fig. 5). To investigate this observation in more detail, we inhibited Notch signaling with DAPT and examined contractile, proliferative and synthetic phenotypes following endothelial cell coculture. Inhibition of Notch signaling resulted in an inability of endothelial cells to induce contractile gene expression as well as collagen gene expression, and also resulted in a failure of proliferative genes, CYCLIN D1 and D2 to be significantly downregulated (Fig. 6B–D). Collagen secretion and secretion rate was also blocked by addition of DAPT (Fig. 6E, F). In contrast, the expression of the synthetic markers, CALDESMON1, RBP1, and VIMENTIN were not affected by Notch inhibition (Fig. 6G–I). These data indicate that Notch signaling contributes to endothelial cell-induced differentiation and synthetic phenotypes, but is not responsible for controlling synthetic marker gene expression.

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Fig. 7. NOTCH2 and NOTCH3 uniquely contribute to phenotypic modulation of smooth muscle cells. HAoSMCs were transiently transfected with control (−), NOTCH2, or NOTCH3 siRNA and then cultured with HUVEC for 96 h. Smooth muscle cells were separated from endothelial cells, prior to expression analysis. (A, B) Expression of NOTCH2 and NOTCH3 RNA and protein following (si)RNA knockdown. (C) Smooth muscle-specific gene expression and (D) gel contraction assay. (E) CYCLIND1 and CYCLIND2 RNA expression and (F) Ki67 quantification of proliferating smooth muscle cells. (G–H) collagen genes, and synthetic markers expression were determined by qPCR. (I) Cell secretion rate was determined by utilizing a constitutively-expressed secreted luciferase plasmid cotransfected with siRNA and cocultured with HUVEC. *P b 0.05, n.s., not significant.

96

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4. Discussion Phenotypic modulation is a unique feature of smooth muscle cells that permits these cells to exhibit a range of phenotypes that meet the current needs of the vascular system. Their ability to transition between phenotypes makes these cells multipurpose, but in disease conditions this can have adverse consequences. Because it is well known that endothelial cells signal to smooth muscle cells during development and in the adult vasculature [6,27], our goal was to determine what influence these cells might have on smooth muscle phenotypic modulation directly. Our data reveal that cocultured endothelial cells impart a contractile and quiescent phenotype that is consistent with a mature differentiated smooth muscle cell. These data are somewhat expected, as it is well established that endothelial cell injury in atherosclerotic lesions or direct removal during angioplasty promotes the transition of smooth muscle cells from a quiescent/contractile cell to a proliferative cell expressing limited differentiation markers [12,22]. Somewhat unexpected was that in addition to causing a differentiated phenotype, endothelial cell coculture also promotes a synthetic phenotype. The synthetic phenotype of smooth muscle cells has been largely associated with increased proliferation [12,22]. However, classical studies performed by Campbell et al., demonstrated that these phenotypes are not codependent [7,21]. Our data indicate that endothelial cells promote the synthetic phenotype, which is somewhat inconsistent with a fully differentiated phenotype. What might the reason be for these endothelial cells to promote this unique phenotype? Possibly these cultured endothelial cells are behaving as they would prior to tube formation, and are instructing the neighboring smooth muscle cells to differentiate and build a basement membrane. Once an intact blood vessel is formed the signals might be changed. Additional signaling mediators in replace of or in addition to Notch signaling could modify the conversation, and endothelial cells might modulate smooth muscle cells differently. Previously, analysis from our lab and others demonstrated a key role for Notch signaling in the communication of endothelial cells and vascular smooth muscle cells [13,18]. The data revealed that Notch signaling via Jagged1 ligand on endothelial cells activates Notch receptors on neighboring smooth muscle cells. Results presented here, indicate that activated Notch signaling is critical for the unique contractile, quiescent and synthetic phenotype that we observed in these cocultured cells. Thus, Notch signaling in smooth muscle cells extends beyond differentiation, and appears to have a more refined role in providing a unique phenotype. This phenotype might be important during certain windows of development or during repair and injury in adult blood vessels. Moreover, our results establish separate and distinct functions of the NOTCH2 and NOTCH3 receptors to regulate specific aspects of the proliferative and synthetic phenotypes. These data are consistent with previously published results [5], and indicate that receptor-specific functions contribute to the phenotypic decisions of a smooth muscle cell. Taken together, these data expand our understanding of the role of endothelial cells and the Notch signaling pathway in controlling phenotypic modulation, and provide additional insight into how vascular smooth muscle cells are regulated. Acknowledgments This work was supported by AHA (786513) grant-in-aid and Nationwide Children's Hospital funds to BL.

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