E XP E RI ME N TAL C E L L R ES E ARC H

324 (2014 ) 92 – 10 4

Available online at www.sciencedirect.com

journal homepage: www.elsevier.com/locate/yexcr

Research Article

Lipid rafts are required for signal transduction by angiotensin II receptor type 1 in neonatal glomerular mesangial cells Adebowale Adebiyin, Hitesh Soni, Theresa A. John, Fen Yang Department of Physiology, College of Medicine, University of Tennessee Health Science Center, 894 Union Avenue, Memphis, TN 38163 , USA

article information

abstract

Article Chronology:

Angiotensin II (ANG-II) receptors (AGTRs) contribute to renal physiology and pathophysiology, but the

Received 18 December 2013

underlying mechanisms that regulate AGTR function in glomerular mesangium are poorly understood.

Received in revised form

Here, we show that AGTR1 is the functional AGTR subtype expressed in neonatal pig glomerular

10 March 2014

mesangial cells (GMCs). Cyclodextrin (CDX)-mediated cholesterol depletion attenuated cell surface

Accepted 13 March 2014

AGTR1 protein expression and ANG-II-induced intracellular Ca2þ ([Ca2þ]i) elevation in the cells. The

Available online 21 March 2014

COOH-terminus of porcine AGTR1 contains a caveolin (CAV)-binding motif. However, neonatal GMCs

Keywords:

express CAV-1, but not CAV-2 and CAV-3. Colocalization and in situ proximity ligation assay detected an

Lipid rafts

association between endogenous AGTR1 and CAV-1 in the cells. A synthetic peptide corresponding to

Caveolin-1 scaffolding domain

the CAV-1 scaffolding domain (CSD) sequence also reduced ANG-II-induced [Ca2þ]i elevation in the

Angiotensin II

cells. Real-time imaging of cell growth revealed that ANG-II stimulates neonatal GMC proliferation.

Glomerular mesangial cells Intracellular calcium Proliferation

ANG-II-induced GMC growth was attenuated by EMD 66684, an AGTR1 antagonist; BAPTA, a [Ca2þ]i chelator; KN-93, a Ca2þ/calmodulin-dependent protein kinase II inhibitor; CDX; and a CSD peptide, but not PD 123319, a selective AGTR2 antagonist. Collectively, our data demonstrate [Ca2þ]i-dependent proliferative effect of ANG-II and highlight a critical role for lipid raft microdomains in AGTR1-mediated signal transduction in neonatal GMCs. & 2014 Elsevier Inc. All rights reserved.

Introduction Glomerular mesangial cells (GMCs) and their surrounding matrix form the mesangium, an important component of the renal glomerulus. The glomerular mesangium serves a variety of functions in the kidney, including organization and support of the glomerular capillaries, endocytosis, and synthesis of vasoactive mediators and cytokines [55,56]. The mesangium is often deranged in kidney diseases. For example, diabetic nephropathy is characterized by mesangial cell n

Corresponding author. Fax: þ901 448 7126. E-mail address: [email protected] (A. Adebiyi).

http://dx.doi.org/10.1016/j.yexcr.2014.03.011 0014-4827/& 2014 Elsevier Inc. All rights reserved.

hypertrophy and matrix accumulation [64]. Glomerular dysfunctions in lupus nephritis, IgA nephropathy (IgAN), and membranoproliferative glomerulonephritis (MPGN) are also associated with mesangial pathology, including hyperproliferation, immune complex deposition, mesangiolysis, and matrix expansion [9,56,57]. Angiotensin II (ANG-II), a major effector of the reninangiotensin-aldosterone system activates multiple signal transduction pathways in the mesangium that induce GMC contraction, proliferation, and extracellular matrix accumulation [55,56,74].

E XP E RI ME N TAL CE L L R ES E ARC H

ANG-II regulates kidney development and neonatal renal functions, including regional renal blood flow, fluid and electrolyte reabsorption, glomerular filtration, and arterial pressure [10,25,26,62,75]. The physiological effects of ANG-II are mediated by its heterotrimeric G protein-coupled receptors (GPCRs): types 1 (AGTR1) and 2 (AGTR2) [32]. AGTR subtype expression patterns in the brain, liver, and kidney are dependent on developmental stage [13,20,79]. However, the expression and physiological regulation of AGTR functions in neonatal mesangium are poorly understood. Lipid rafts, the cholesterol-enriched plasma membrane microdomains localize and control many signaling proteins, including the GPCRs [11]. Caveolae, the specialized lipid rafts and caveolins, the caveolae/raft scaffolding proteins regulate AGTR functions in vascular smooth muscle cells [71]. In rat aortic myocytes, ANG-II stimulated AGTR1 translocation to caveolin-enriched plasmalemmal fraction [29]. Caveolae disruption also attenuated ANG-II-induced tachyphylactic contractions and epidermal growth factor receptor transactivation in rat aortic rings and cultured rat aortic myocytes, respectively [37,72]. Lipid raft microdomains are present in GMCs and are involved in growth factor-regulated signal transductions [47,68]. However, it remains unclear whether AGTR functions in neonatal GMCs are controlled by lipid rafts. Here, we tested the hypothesis that lipid rafts are essential for ANG-II-induced intracellular Ca2þ ([Ca2þ]i) elevation and proliferation in neonatal GMCs.

Materials and methods Animals Use of animals in this study was reviewed and approved by the Animal Care and Use Committee of the University of Tennessee Health Science Center (UTHSC). Male newborn pigs (1-2 days old) were purchased from Nichols Hog Farm (Olive Branch, MS USA) and maintained at the UTHSC animal core facility. Animals were used within the first week of life.

Isolation and propagation of neonatal pig GMCs Neonatal pigs were euthanized by injection of ketamine/xylazine (100/10 mg/kg; i.m.) followed by exsanguination (by severing the abdominal aorta). After euthanasia, the kidneys were removed and placed in Dulbecco's modified Eagle's medium (DMEM; Life Technologies, Grand Island, NY). Renal glomeruli were isolated from the kidneys by serial sieving of homogenates from kidney cortical strips using sterile stainless steel meshes (pore size: 300, 180, and 90 mm). Isolated glomeruli were then decapsulated by passing them through a syringe fitted with a 25-gauge needle. Decapsulated glomeruli were cultured at 37 1C in Dulbecco's modified Eagle's medium (DMEM; Life Technologies) supplemented with mesangial cell growth medium (ScienCell Research Laboratories Carlsbad, CA), 2% fetal bovine serum (FBS; Atlanta Biologicals, Lawrenceville, GA), and 1% penicillin/streptomycin (Sigma-Aldrich, St. Louis, MO). This procedure generated a pure population of GMCs after 3–4 weeks in culture. Cultured GMCs were characterized by negative immunostaining for podoplanin (PDPN), a podocyte marker [39] and cytokeratin 18 (KRT18), an epithelial cell marker [42] and positive staining for regulator of G-protein signaling 5 (RGS5), a pericyte marker [12] and alpha smooth muscle actin (α-SMA) [42], a myogenic marker (Fig. 1).

324 (2014 ) 92 – 10 4

93

Reverse transcription polymerase chain reaction (RT-PCR) Total RNA isolation and cDNA synthesis were performed using the RNA MicroPrep (Zymo Research Corp., Orange, CA) and QuantiTect RT (Qiagen, Valencia, CA) kits, respectively. PCR amplification was performed in an Eppendorf Mastercycler (Eppendorf, Westbury, NY) using the EconoTaq Plus Green PCR Master Mix (Lucigen Corporation, Middleton WI) and oligonucleotide primer pairs (Table 1). Reactions were run under the following conditions: an initial denaturation at 98 1C for 2 min, followed by 35 cycles (denaturation at 98 1C for 10 s, annealing at 57 1C for 30 s, and extension at 72 1C for 30 s), with a final extension at 72 1C for 10 min. PCR products were separated on 1.5% agarose gels stained with GelRed dye (Biotium, Hayward, CA) and documented using a Kodak In Vivo F Pro Imaging System (Carestream Molecular Imaging, Rochester, NY).

Western immunoblotting Cultured GMCs were washed with phosphate buffered saline (PBS) and homogenized in ice-cold RIPA buffer supplemented with protease inhibitor cocktails (Thermo Scientific, Rockford, IL). Following centrifugation to remove cell debris, protein concentrations were determined using a Bio-Rad protein assay kit and a SmartSpec 3000 Spectrophotometer (Bio-Rad, Hercules, CA). Protein lysates were mixed with lithium dodecyl sulfate (LDS) sample loading buffer containing 5% β-mercaptoethanol and boiled at 95 1C for 5 min. Proteins were then separated on 8% Bis-Tris Plus gels using MES SDS buffer in a Bolt Mini Gel Tank (Life Technologies) and transferred onto nitrocellulose membranes using a Semi-Dry Blotter (Thermo Scientific). Nonspecific binding sites on the membranes were blocked with a NAP blocking buffer (G-Biosciences, St. Louis, MO) for  1 h at room temperature. The membranes were probed with respective primary antibodies overnight at 4 1C. After wash in Tris buffered saline supplemented with 0.05% Tween 20 (TBS-T), the membranes were incubated in horseradish peroxidase-conjugated secondary antibodies for 45 min at room temperature and washed in TBS-T. Immunoreactive proteins were visualized using the femtoLUCENT PLUS-HRP chemiluminescent reagent (G-Biosciences).

Biotinylation of cell surface proteins Sub-confluent GMCs were washed with ice-cold phosphate buffered saline (PBS) and incubated at 4 1C for 2 h in a mixture of EZ-Link Sulfo-NHS-LC-LC-Biotin and EZ-Link Maleimide-PEG2Biotin reagents (Thermo Scientific; 0.5 mg/ml each) in PBS. Afterward, the cells were washed with PBS and incubated with 100 mM glycine in ice-cold PBS for 30 min at 4 1C to quench unbound biotin. The cells were then homogenized in RIPA buffer followed by spectrophotometric determination of protein concentration. Dynabeads streptavidin (60 mL; Life Technologies) were mixed with 800 mg total protein and incubated at 4 1C overnight using gentle rotation. A DynaMag magnet (Life Technologies) was used to separate proteins bound to the Dynabeads. The beads were then washed 6 times in PBS containing 0.1% BSA. Immobilized biotinylated proteins were eluted from the Dynabeads by boiling the samples for 5 min in LDS sample buffer containing 5%

94

EX P ER I ME NTAL C E LL RE S E ARCH

3 24 (2014) 9 2 –10 4

Fig. 1 – Characterization of primary neonatal pig GMCs. Representative immunostaining of neonatal pig GMCs demonstrating negative staining for PDPN and KRT18, and positive staining for RGS5 and α-SMA. Rabbit and mouse IgGs (negative controls) did not show fluorescence. Bar¼ 10 lm.

β-mercaptoethanol. Proteins were then resolved using Western immunoblotting.

Immunofluorescence staining and confocal microscopy GMCs sparsely grown on glass coverslips were fixed in 4% formaldehyde and permeabilized with 0.2% Triton X-100 for 20 min at room temperature. After 1 h of incubation in a blocking buffer (Olink Bioscience, Uppsala, Sweden), cells were incubated overnight at 4 1C with respective antibodies (1:10-40, each) in a PELCO Stain Saver (Ted Pella Inc., Redding, CA). Next day, cells were washed with PBS and incubated with DyLight 488conjugated pre-adsorbed anti-rabbit or DyLight 550-conjugated pre-adsorbed anti-mouse secondary antibodies (1:80, each) for 1 h at room temperature. Following wash and mount, images were acquired using a Zeiss laser-scanning confocal microscope. DyLight 488 and DyLight 550 were excited at 488 and 543 nm and emission collected at 505–530 and 4560 nm, respectively. Colocalization was examined using a Coloc_2 analysis tool in Fiji image processing software [54]. The degree of colocalization was quantified using the Mander's correlation coefficient [38].

In situ proximity ligation assay (PLA) Interaction between endogenous AGTR1 and CAV-1 in GMCs was investigated using the in situ proximity ligation assay kit (Duolink kit; Olink Bioscience) [1,60]. GMCs were sparsely cultured in a glass chamber slide (Thermo Scientific). The cells were fixed and permeabilized as described above. After blocking nonspecific sites with Duolink blocking reagent, GMCs were incubated with anti-AGTR1 and anti-CAV-1 antibodies overnight at 4 1C. Negative control slides were incubated with antiCAV-1 antibody only. Cells were washed and incubated with secondary antibodies conjugated with oligonucleotides (antimouse PLA probe Minus and anti-rabbit PLA probe Plus) in a humidity chamber (Thermo Scientific) for 1 h at 37 1C. Cells were then incubated with a ligation solution for 30 min at 37 1C. Ligation of the oligonucleotides probes was followed by a rolling-circle amplification reaction [60]. The amplification products were detected by a fluorescently (Cy3)-labeled complementary oligonucleotide detection probes. Slides were mounted with a mounting medium containing DAPI nuclear stain (Olink Bioscience). PLA signals were examined using a Zeiss confocal microscope.

E XP E RI ME N TAL CE L L R ES E ARC H

95

324 (2014 ) 92 – 10 4

Table 1 – Oligonucleotide primer sequences. Gene

Sequence

β-Actin Forward Reverse

50 -CTGGCCGCACCACTGGCATTGTC-30 50 -CGTGGTGGTGAAGCTGTAGCCCC-30

AGTR1 Forward Reverse

50 -CCCCAAAGCTGGAAGGCATA-30 50 -GAAGGCGGGACTTCATTGGA-30

AGTR2 Forward Reverse

50 -TGGCTGTGGCTGACTTACTG-30 50 -GAGGCTTGCCAGGGATTTCT-30

CAV-1 Forward Reverse

50 -CCAGCTGAATGAGGTCAGCA-30 50 -GTCTCACACAATGGCCTCCA-30

CAV-2 Forward Reverse

50 -GGACATCCTCCCTCATCCCT-30 50 -CATCCGCCTTCTCAGTCTCC-30

CAV-3 Forward Reverse

50 -TCGAGATCCAGTGCATCAGC-30 50 -TTCTTTCCGCAGCATCACCT-30

n

GenBank Accession Number

Length (bp)

DQ845171.1

176

XM_003132469.2n

362

XM_005673847.1n

226

NM_214438.2

181

NM_001123091.1

332

NM_001037149.1

116

Predicted.

Intracellular Ca2þ ([Ca2þ]i) concentration measurement GMCs seeded in glass-bottom dishes were washed with PBS and incubated with Fura-2-acetoxymethyl ester (Fura-2 AM; 10 μM), and 0.5% pluronic F-127 for  1 h at room temperature in modified Krebs’ solution (MKS; 134 mM NaCl, 6 mM KCl, 1.2 mM CaCl2, 1 mM MgCl2, 10 mM HEPES, and 5.5 mM glucose, pH 7.4). Cells were then washed for 30 min before imaging to deesterify Fura-2 AM. [Ca2þ]i imaging was performed at room temperature using a ratiometric fluorescence system (Ionoptix Corp., Milton, MA, USA). Fura-2 AM fluorescence was recorded from cells located in the same field by exciting at wavelengths of 340 and 380 nm using a hyperswitch light source (Ionoptix). Only one field was imaged per dish. Background-subtracted Fura-2 AM ratios were collected at 510 nm using a MyoCam-S CCD digital camera (Ionoptix). GMC [Ca2þ]i concentrations were analyzed with IonWizard software (Ionoptix) using the following equation [23]: [Ca2þ]i ¼ Kd [(R Rmin) / (Rmax  R)] δ where “R” is the 340/380 nm ratio, and Rmin and Rmax are the minimum and maximum Fura-2 ratios determined in Ca2þ-freeþ EGTA and Ca2þ-replete solutions, respectively. δ represents the ratio of the 380 nm excitation in Ca2þ-free and Ca2þ-replete solution, while Kd is the apparent dissociation constant for Fura2 (224 nM, [23]). Rmin, Rmax, and δ were determined at the end of each experiment by perfusing the cells with 10 mM ionomycin and Ca2þ-free (plus 10 mM EGTA) or 10 mM Ca2þ solution.

Live content microscopy of GMC growth GMCs were sparingly plated in sterile flat-bottom, tissue culture plates (Corning Inc., Corning, NY) and starved overnight by culturing in 0.1% FBS/DMEM. Thereafter, the cells were treated

with respective test substances and the culture plates were placed in a chamber apparatus of the IncuCyte ZOOM live content microscopy system (Essen Instruments, Ann Arbor, MI) in an incubator (5% CO2, 37 1C). GMC growth kinetics was monitored in real-time. Growth curves were determined from cell confluence percentage automatically acquired by the IncuCyte cell density detection software at two-hourly intervals.

Antibodies (catalog numbers are in parentheses) and chemicals Rabbit polyclonal anti-AGTR1 (SAB3500209) and rat monoclonal antiPDPN (P0085) antibodies were purchased from Sigma-Aldrich (St. Louis, MO). Mouse monoclonal anti-CAV-1 (sc-53564), mouse monoclonal anti-RGS5 (TA503075), and rabbit monoclonal antiCYK18 (AJ1223) antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, CA), Origene Technologies (Rockville, MD), and Abgent Inc. (San Diego, CA), respectively. Mouse monoclonal anti-actin (ab3280) and anti-α-SMA (ab7817) primary antibodies, DyLight pre-adsorbed donkey anti-mouse (ab98795) and anti-rabbit (ab96919) and HRP-conjugated anti-rabbit (ab96919) and anti-mouse (ab98795) secondary antibodies were purchased from Abcam (Cambridge, MA). DyLight goat anti-rat secondary antibody (405409) was purchased from BioLegend (San Diego, CA). PD 123319 and EMD 66684 were purchased from Tocris Bioscience (Bristol UK). Ionomycin and KN-93 were purchased from Cayman Chemical (Ann Arbor, MI). Antennapedia and FITC-conjugated antennapedia peptides and Pluronic F-127 were purchased from Anaspec (Fremont, CA). 236-TriO-Butyryl-myo-Inositol-145-Trisphosphate-Hexakis (acetoxymethyl) Ester (Bt-IP3), CAV-1 scaffolding domain peptide, Fura-2 AM, ANG II, cyclodextrin, and BAPTA-AM were purchased from A.G. Scientific (San Diego, CA), Enzo Life Sciences (Farmingdale, NY), Life Technologies, California Peptide Research Inc. (Napa, CA), Sigma-Aldrich, and Assay Biotechnology Co., Inc. (Sunnyvale, CA) respectively.

96

EX P ER I ME NTAL C E LL RE S E ARCH

Data analysis Statistical significance was determined using Student's t-tests for paired or unpaired data and analysis of variance with Bonferroni post-hoc test for multiple comparisons. All data are expressed as

3 24 (2014) 9 2 –10 4

mean7 standard error of mean (SEM). A P valueo0.05 was considered significant.

Results Neonatal pig GMCs express only AGTR1 AGTR expression in neonatal pig GMCs was investigated using RT-PCR. PCR amplification generated amplicons of expected sizes for AGTR1 (362 bp) and β-actin (ACTB; 176 bp), but not AGTR2 (226 bp) (Fig. 2). No band was observed in negative control samples probed with AGTR1 primers, but in which no reverse transcriptase was added during cDNA synthesis (Fig. 2). These data suggest that neonatal pig GMCs express only AGTR1.

Fig. 2 – Neonatal pig GMCs express AGTR1. A representative gel image showing RT-PCR amplification of AGTR1 (362 bp) and β-actin (ACTB; 176 bp), but not AGTR2 (226 bp) transcripts in neonatal pig GMCs. No band was seen in samples probed with AGTR1 primers, but in which no reverse transcriptase (RT) was added during cDNA synthesis.

Lipid rafts regulate AGTR1 cell surface expression and ANG-II-induced [Ca2þ]i elevation in neonatal pig GMCs At the plasma membrane, lipid raft microdomains localize and integrate a wide variety of signaling molecules, including GPCRs [11]. To examine whether lipid rafts are required for plasma membrane

Fig. 3 – Cholesterol depletion reduces cell surface AGTRI. (A) Western blot and (B) mean data (n¼ 4 each) illustrating that cell surface AGTR1 protein is reduced in CDX (10 mM; 2 h)-pretreated neonatal pig GMCs. (C) Western blot and (D) mean data (n¼ 4 each) demonstrating that total AGTR1 protein is unchanged in CDX-pretreated neonatal pig GMCs. nPo0.05 vs. control.

E XP E RI ME N TAL CE L L R ES E ARC H

expression of AGTR1 in GMCs, we depleted membrane cholesterol by incubating the cells in β-cyclodextrin (CDX; 10 mM) for 2 h at 37 1C. The concentration of CDX used has previously been shown to effectively reduce membrane cholesterol with insignificant cell damage [27,33,45]. Cell surface proteins were then biotinylated and examined for the expression of AGTR1. As shown in Fig. 3A and B, immunoreactive AGTR1 in GMC surface was reduced by  50% in CDX-treated cells. In contrast, CDX treatment did not alter immunoreactive AGTR1 in total cell lysate (Fig. 3C and D). These data suggest that lipid rafts are necessary for plasma membrane protein expression of AGTR1 in neonatal pig GMCs. Next, we measured ANG-II-induced [Ca2þ]i elevation in control and CDX-treated cells. ANG-II alone increased [Ca2þ]i concentration in the cells by  122 nM (Fig. 4A and B). ANG-II-induced [Ca2þ]i elevation was essentially abolished by EMD 66684, a selective AGTR1 antagonist [40]. In contrast, PD 123319, a selective AGTR2 antagonist [8] did not alter ANG-II-induced [Ca2þ]i elevation (Fig. 4A and B). These data indicate that AGTR1 mediates ANG-II-induced [Ca2þ]i elevation in neonatal pig GMCs. CDX did not change basal [Ca2þ]i concentration in the cells [untreated (n¼ 13) vs. CDX-treated cells (n ¼10): 3373 vs. 3975 nM; P40.05]. However, ANG-II increased [Ca2þ]i concentration in cells treated with CDX by  43 nM, indicating that CDX reduced ANG-II-induced [Ca2þ]i elevation by  65% (Fig. 4A and B). To examine the effect of cholesterol replenishment on ANG-IIinduced [Ca2þ]i elevation, CDX-treated cells were incubated at 37 1C in water-soluble cholesterol for 2 h. As shown in Fig. 4A and B, cholesterol repletion reversed the inhibitory effect of CDX on ANG-II-induced [Ca2þ]i elevation. Collectively, these data indicate that lipid rafts are required for ANG-II-induced [Ca2þ]i elevation in neonatal pig GMCs. To examine whether CDX treatment alters inositol 1,4,5-trisphosphate receptor (IP3R)-mediated endoplasmic reticulum (ER) Ca2þ release, we studied IP3-induced ER Ca2þ release in control

324 (2014 ) 92 – 10 4

97

and CDX-treated cells. In the absence of extracellular Ca2þ, Bt-IP3, a membrane permeant IP3 analog [1,3,77] increased [Ca2þ]i by 135.5717 nM (Po0.05; n ¼4), indicative of IP3R-mediated Ca2þ release from the ER store. However, Bt-IP3-induced ER Ca2þ

Fig. 5 – Lipid raft disruption does not alter IP3-induced ER Ca2þ release. (A) Exemplar traces and (B) mean data illustrating that Bt-IP3 (10 lM)-induced [Ca2þ]i elevation in the absence of extracellular Ca2þ is unchanged in CDX-treated cells. P40.05; n¼ 4 each.

Fig. 4 – Lipid rafts are required for AGTR1-mediated [Ca2þ]i elevation in neonatal pig GMCs. (A) Exemplar traces, and (B) mean data showing ANG-II (10 lM)-induced [Ca2þ]i elevation in control (n ¼13), EMD 66684 (100 nM;  20 min; n¼ 7)-, PD 123319 (100 nM;  20 min; n ¼5)-, CDX (10 mM; 2 h; n¼ 10)-, and CDXþcholesterol (1 mg/mL; 2 h; n¼ 6)-treated cells. nPo0.05 vs. control.

98

EX P ER I ME NTAL C E LL RE S E ARCH

release was unchanged in CDX-treated cells, suggesting that plasmalemmal cholesterol depletion does not directly alter IP3Rmediated ER Ca2þ release in neonatal GMCs (Fig. 5).

AGTR1 interacts with CAV-1 in neonatal pig GMCs Most signaling proteins that interact with caveolins contain aromatic-rich caveolin-binding motifs (CBMs; ΦXΦXXXXΦ and ΦXXXXΦXXΦ, where Φ is an aromatic residue tryptophan, phenylalanine or tyrosine) [14,43]. Analysis of the predicted porcine AGTR1 protein sequence (NCBI Reference Sequence: XP_003132517.1) showed one potential CBM in the COOHterminus suggesting that AGTR1 may associate with caveolins in GMCs (Fig. 6A). To investigate whether AGTR1 interacts with caveolins in the cells we first examined the expression of caveolin isoforms. RT-PCR experiments indicated that CAV-1 (181 bp), but not CAV-2 (332 bp) and CAV-3 (116 bp), is expressed in the cells (Fig. 6B). Next, immunofluorescence labeling and in situ PLA were used to investigate colocalization and interaction between endogenous AGTR1 and CAV-1 in neonatal pig GMCs. Data suggest that AGTR1 colocalizes extensively with CAV-1 in the plasma membrane (percentage of colocalization: 90.272.4; n ¼25 cells; Fig. 6C). PLA signals (red florescent dots) are detected when two

3 24 (2014) 9 2 –10 4

interacting proteins are in close proximity (o 40 nm separation) [60]. As shown in Fig. 6D, PLA signals were absent in control GMCs labeled with anti-CAV-1 antibody alone. Conversely, PLA signals were detected in GMCs labeled with both anti-AGTR1 and anti-CAV-1 antibodies (Fig. 6D). These results suggest an association between endogenous AGTR1 and CAV-1 in neonatal pig GMCs.

Endogenous CAV-1 scaffolding domain regulates ANG-IIinduced [Ca2þ]i elevation in neonatal pig GMCs The N-terminal region of CAV-1 contains a scaffolding domain (CSD; amino acids 82-101) that interacts with CBMs on caveolinbinding proteins [43]. As shown in Fig. 6A, porcine AGTR1 protein sequence contains a CBM. AGTR1 also interacts with CAV-1 in neonatal pig GMCs (Fig. 6C and D). To test the hypothesis that endogenous CSD regulate ANG-II-induced [Ca2þ]i elevation in the cells, we inhibited the coupling between the CAV-1 and AGTR1 by treating the cells with a synthetic CSD peptide (sequence: DGIWKASFTTFTVTKYWFYR). The CSD peptide was conjugated with an internalization sequence of the Drosophila antennapedia homeodomain (AP; sequence: RQIKIWFQNRRMKWKK) to enhance cell membrane permeability [15,34]. The AP peptide was used as the control.

Fig. 6 – Endogenous AGTR1 associates with CAV-1 in neonatal pig GMCs. (A) Porcine AGTR1 protein sequence contains a CBM (highlighted in yellow) in the COOH-terminus. Bold letters indicate tyrosine or phenylalanine. (B) A representative gel image showing RT-PCR amplification of CAV-1 (181 bp) and β-actin (ACTB; 176 bp), but not CAV-2 (332 bp) and CAV-3 (116 bp) transcripts in neonatal pig GMCs. (C) Immunofluorescence staining of neonatal pig GMCs demonstrating that AGTR1 colocalizes with CAV-1. (D) In situ PLA detects an interaction between AGTR1 and CAV-1 in neonatal pig GMCs. Arrows in the panel point to PLA signals (red fluorescence). PLA signals were absent in control cells labeled with only anti-CAV-1 antibody. Bar ¼ 10 lm.

E XP E RI ME N TAL CE L L R ES E ARC H

324 (2014 ) 92 – 10 4

99

Fig. 7 – AP-CSD peptide inhibits ANG-II-induced [Ca2þ]i elevation in neonatal pig GMCs. (A) FITC-labeled AP peptide confirmed successful GMC uptake of the peptides. (B) Exemplar traces, and (C) mean data showing ANG-II (10 lM)-induced [Ca2þ]i elevation in control (AP; 20 lM;  24 h; n¼ 8) and AP-CSD (20 lM;  24 h; n¼ 7)-pretreated neonatal pig GMCs. nPo0.05 vs. AP.

FITC-labeled AP peptide confirmed successful cellular uptake of the peptides (Fig. 7A). AP-CSD treatment did not change basal [Ca2þ]i concentration compared with AP peptide [AP (n¼ 8) vs. AP-CSD (n¼ 7): 2273 vs. 2873 nM; P40.05]. Furthermore, AP peptide did not alter ANG-II-induced [Ca2þ]i elevation in the cells [untreated (n¼ 13) vs. AP-treated (n ¼8) GMCs: 122.1711.6 vs. 116.878.0 nM; P40.05]. In contrast, AP-CSD peptide attenuated ANG-II-induced [Ca2þ]i elevation by 69% in the cells (Fig. 7B and C). Taken together, these data indicate that CSD controls ANG-II-induced [Ca2þ]i elevation in neonatal pig GMCs.

CDX and AP-CSD peptide attenuate ANG-II-induced neonatal pig GMC proliferation To examine whether lipid rafts are required for ANG-II-induced neonatal GMC proliferation, we studied cultured neonatal pig GMC growth kinetics over 48 h periods. ANG-II caused a concentration- and time-dependent proliferation of the cells (Fig. 8A and B). ANG-II-induced proliferation of the cells was inhibited by EMD 66684, but not PD 123319 (Fig. 8C). As shown in Fig. 4, ANG-II increases [Ca2þ]i concentration in neonatal pig GMCs. Apart from stimulating acute cell contraction, [Ca2þ]i-dependent signaling through the Ca2þ/calmodulin-dependent protein kinases (CAMK) can also promote GMC growth [55,73]. To investigate the role of [Ca2þ]i in ANG-II-induced GMC proliferation, we studied the effect of BAPTA-AM, a [Ca2þ]i chelator and KN-93, a CAMKII inhibitor on ANGII-induced GMC growth. Both BAPTA and KN-93 reduced ANG-IIinduced proliferation in the cells, suggesting that an elevation in [Ca2þ]i concentration contributes to the mitogenic effect of ANG-II in neonatal pig GMCs (Fig. 8C). Between 10 and 48 h, CDX attenuated ANG-II-induced GMC proliferation by 460% (Fig. 9A). Repletion of membrane cholesterol almost completely reversed the inhibitory effect of CDX on ANG-II-induced GMC growth (Fig. 9A). AP-CSD

peptide also reduced ANG-II-induced proliferation of the cells (Fig. 9B). Collectively, these findings suggest that ANG-II-induced activation of AGTR1 stimulates [Ca2þ]i-dependent neonatal pig GMC proliferation and this effect is regulated by lipid rafts and CSD.

Discussion Data here show that: (1) neonatal pig GMCs express AGTR1, but not AGTR2; (2) a CBM exists in the C-terminus of porcine AGTR1 protein sequence; (3) neonatal pig GMCs express CAV-1, but not CAV-2 and CAV-3; (4) endogenous AGTR1 and CAV-1 exist in close molecular proximity in neonatal pig GMCs; (5) activation of AGTR1 by ANG-II increases [Ca2þ]i concentration in neonatal pig GMCs; (6) disruption of lipid rafts reduces AGTR1 cell surface protein expression and ANG-II-induced [Ca2þ]i elevation; (7) a synthetic CSD peptide inhibits ANG-II-induced [Ca2þ]i elevation in neonatal pig GMCs; (8) ANG-II-induced AGTR1 activation promotes neonatal pig GMC growth; (9) [Ca2þ]i elevation contributes to ANG-II-induced neonatal pig GMC proliferation; and (10) lipid raft disruption attenuates ANG-II-induced neonatal pig GMC proliferation. Collectively, these new findings suggest that plasma membrane lipid raft microdomains are requisite for AGTR1 surface expression and hence, ANG-II-induced [Ca2þ]i elevation and proliferation in neonatal GMCs. Adult mouse, rat, and human GMCs express both AGTR1 and AGTR2 [13,24,35,78]. In contrast, AGTR1 is predominantly expressed in cultured human fetal GMCs [50]. PCR experiments in the present study indicate that only AGTR1 is expressed in neonatal pig GMCs. Furthermore, ANG-II-induced increase in [Ca2þ]i concentration and proliferation was attenuated by selective AGTR1, but not AGTR2 antagonist. These findings indicate that AGTR1 is the functional AGTR subtype expressed in neonatal pig

100

EX P ER I ME NTAL C E LL RE S E ARCH

3 24 (2014) 9 2 –10 4

Fig. 8 – AGTR1-mediated [Ca2þ]i elevation promotes neonatal pig GMC proliferation. (A) Phase contrast images (segmentation mask illustrated in goldenrod), and (B) cell growth curves measured over 48 h showing time- and concentration-dependent proliferative effect of ANG-II (n¼ 4 each) in neonatal pig GMC. (C) Graphs illustrating that ANG-II-induced (n¼6) neonatal pig GMC proliferation is inhibited by EMD 66684 (100 nM; n¼ 6), BAPTA-AM (2 lM; n ¼4), and KN-93 (2 lM; n¼ 4), but not PD 123319 (100 nM; n¼ 6). “n” denotes number of wells. Each time point represents four independent scans per well of the culture flask. #Po0.05 vs. control; n Po0.05 vs. ANG-II; bar¼ 300 lm.

GMCs. ANG-II regulates renal perfusion in neonates and adults by constricting pre- and post-glomerular microvessels [4,62]. ANG-IIinduced [Ca2þ]i elevation also stimulates GMC contraction [66]. Given that GMC contraction may alter glomerular hemodynamics [30,65], it seems likely that activation of AGTR1 localized in GMCs will contribute to ANG-II-induced alteration in neonatal glomerular microcirculation. Lipid rafts are involved in AGTR trafficking, membrane localization, and physiological functions in a wide variety of cells. For example, beta-arrestins, the adapter proteins that regulate G-proteinindependent functions of AGTR1 [58,59] are localized in lipid rafts of human umbilical vein endothelial and HEK293-APP695 cells [61,69]. AGTR1 and AGTR2 exist in macromolecular protein complexes with CAV-1 in cultured human internal mammary artery smooth muscle cells [41]. AGTR1 density was less in the kidneys of mice lacking CAV-1 when compared with their wild type counterparts [76]. CBM was also required for AGTR1 plasma membrane

expression in HEK and COS-7 cells expressing AGTR1 [36,76]. The presence of caveolae invaginations in GMCs has been demonstrated [68]. CAV-1 and CAV-2, but not CAV-3, are expressed in adult mouse GMCs [31]. In contrast, our data show that neonatal pig GMCs express only CAV-1. These contradictory observations could be due to species variability in caveolin expression. It is also possible that caveolin isoform expression in GMCs is dependent on kidney maturation. The presence of a CBM in porcine AGTR1 COOH-terminus suggested a possible physical and functional interaction between AGTR1 and CAV1. Indeed, the results of our colocalization and PLA experiments signify an association between endogenous AGTR1 and CAV-1 in the plasma membrane of neonatal pig GMCs. Cholesterol is an important component required to maintain lipid raft structural and functional integrity [51]. Caveolins act as scaffolding proteins or molecular chaperones in lipid rafts [14,44,51]. Hence, treatment with CDX, a sterol-binding agent that depletes membrane cholesterol, disrupts lipid rafts and alters

E XP E RI ME N TAL CE L L R ES E ARC H

Fig. 9 – CDX and AP-CSD peptide attenuate ANG-II-induced neonatal pig GMC proliferation. (A) Cell growth curves showing ANG-II-induced proliferation in control (n¼4), CDX (10 mM; 2 h; n¼ 6)-, and CDXþcholesterol (1 mg/mL; 2 h; n ¼6)treated cells. (B) Graphs demonstrating ANG-II-induced proliferation in AP (20 lM;  24 h; n¼ 4)- and AP-CSD peptide (20 lM; 24 h; n¼4)-treated neonatal pig GMCs. nPo0.05 vs. control; δPo0.05 vs. CDXþcholesterol; #Po0.05 vs. AP.

signal transduction by caveolin-associated proteins in a variety of cells, including GMCs [44,47,72]. We observed that about half of cell surface AGTR1 protein was lost in neonatal pig GMCs treated with CDX, suggesting that lipid raft association is essential for plasma membrane localization of AGTR1 in the cells. To study the physiological impact of lipid raft disruption on AGTR1 activity, we examined the effects of CDX on ANG-II-induced [Ca2þ]i elevation. Unlike a previous report in rat tail artery smooth muscle [16], treatment of neonatal GMCs with CDX did not alter basal [Ca2þ]i concentration, suggesting that GMC membrane permeability was not compromised by CDX. However, ANG-II-induced increase in [Ca2þ]i concentration was reversibly attenuated by cholesterol depletion, indicating that intact lipid rafts are necessary for AGTR1 function. CSD binds and regulates several lipid raftassociated signaling proteins, including ion channels, endothelial

324 (2014 ) 92 – 10 4

101

nitric-oxide synthase, and G-proteins [14,21,34,46,67]. Hence, a synthetic CSD peptide can competitively block the binding of CAV-1-interacting proteins to endogenous CAV-1 [34,67]. Here, a CSD peptide did not alter basal [Ca2þ]i concentration in neonatal GMCs, but reduced ANG-II-induced [Ca2þ]i elevation in the cells to a similar degree as CDX treatment. These data suggest that through its scaffolding domain, CAV-1 regulates ANG-II-induced increase in neonatal GMC [Ca2þ]i concentration. ANG-II-induced [Ca2þ]i elevation involves phospholipase C (PLC) hydrolysis of phosphatidylinositol-4,5-bisphosphate [PI(4,5)P2] to IP3 [70]. IP3 stimulates Ca2þ release from the ER store via IP3Rs [6,70]. PI(4,5)P2 is localized in the caveolae of human dermal fibroblasts and mouse smooth muscle cells [19]. Whether PI(4,5)P2 is concentrated in GMC lipid rafts is unclear. However, if lipid raft disruption alters GMC PI(4,5)P2, ANG-IIinduced generation of IP3, and hence [Ca2þ]i elevation will be impaired. ANG-II can also activate members of the transient receptor potential (TRP) cation channels leading to [Ca2þ]i elevation in GMCs [17,22]. Lipid rafts are known to regulate TRP channel functions [2,5]. Therefore, alterations in TRP channel signaling by lipid raft/caveolae disassembly may contribute to the reduction in ANG-II-induced [Ca2þ]i elevation. Unlike, ANG-IIinduced [Ca2þ]i elevation, IP3-mediated ER Ca2þ release was unaltered by CDX, suggesting that plasma membrane lipid raft disruption does not directly interfere with IP3R-mediated ER Ca2þ release in neonatal pig GMCs and that ER disruption does not contribute to the inhibitory effect of CDX on ANG-II-induced [Ca2þ]i elevation. ANG-II promotes proliferation of adult and fetal GMCs [49,74]. Here, we provide new evidence indicating that ANGII also stimulates neonatal GMC proliferation by activating AGTR1. Binding of Ca2þ to intracellular proteins stimulates cascades of signal transduction pathways that result in activation of cell cyclepromoting transcription factors [7]. For example, binding of [Ca2þ]i to calmodulin (CaM) activates CAMKII, leading to phosphorylation of the Ca2þ/cAMP response element-binding protein in GMCs [80]. Data here reveal that chelation of [Ca2þ]i reduces ANG-II-induced neonatal GMC growth. Furthermore, pretreatment of the cells with a CAMKII inhibitor attenuated ANG-IIinduced proliferation. These findings suggest that ANG-II-induced [Ca2þ]i elevation and sequential activation of CAMKII contribute to neonatal GMC growth. Since ANG-II-induced [Ca2þ]i elevation is diminished in cholesterol-depleted and CSD peptide-treated cells, we reasoned that GMC proliferation may also be altered by these agents. To test this hypothesis, we examined GMC growth in CDX- and CSD peptide-treated cells. Both CDX and CSD peptide reduced ANG-II-induced GMC proliferation. These findings suggest that lipid rafts and endogenous CSD are required for ANG-IIinduced neonatal GMC growth. ANG-II stimulates human GMC growth via activation of c-Jun NH2-terminal kinase [81]. Other members of the mitogen-activated protein kinase (MAPK) family are involved in the proliferative effects of ANG-II in many cell types [28]. Since MAPKs associate with lipid rafts [48], our study does not rule out the possibility that MAPKs may contribute to neonatal GMC proliferation. Although CAV-1 is involved in caveolae biogenesis, CAV-1 can exist in non-caveolar rafts [18,53]. Acute cholesterol depletion can disrupt both caveolar and non-caveolar lipid rafts [44,63]. Therefore, the methods used in this study cannot discriminate between signal transduction in caveolar and non-caveolar lipid raft

102

EX P ER I ME NTAL C E LL RE S E ARCH

microdomains. The nature of non-caveolar lipid rafts in GMCs is unclear. Thus, further studies are required to explore whether non-caveolar rafts contribute to AGTR1 membrane localization and AGTR1-mediated signal transduction in GMCs. In conclusion, we present new evidence here indicating that AGTR1 is the functional AGTR subtype expressed in neonatal pig GMCs. Our data also suggest that lipid raft microdomains are required for AGTR1 localization, and hence AGTR1-mediated [Ca2þ]i elevation and proliferation in neonatal pig GMCs. Given that ANG-IIinduced GMC proliferation contributes to renal dysfunction [52], lipid rafts and CAV-1 may constitute potential therapeutic targets in ANG-II-associated nephropathy.

Author contributions Conceived and designed the experiments: A. Adebiyi. Performed the experiments: A. Adebiyi, H. Soni, T.A. John, and F. Yang. Analyzed the data: A. Adebiyi and H. Soni. Wrote the paper: A. Adebiyi.

Conflict of interest None.

Acknowledgments This work was supported by a start-up fund from UTHSC to Dr. Adebiyi.

references [1] A. Adebiyi, RGS2 regulates urotensin II-induced intracellular Ca2þ elevation and contraction in glomerular mesangial cells, J. Cell Physiol. 229 (2014) 502–511. [2] A. Adebiyi, D. Narayanan, J.H. Jaggar, Caveolin-1 assembles type 1 inositol 1,4,5-trisphosphate receptors and canonical transient receptor potential 3 channels into a functional signaling complex in arterial smooth muscle cells, J. Biol. Chem. 286 (2011) 4341– 4348. [3] A. Adebiyi, G. Zhao, D. Narayanan, C.M. Thomas-Gatewood, J.P. Bannister, J.H. Jaggar, Isoform-selective physical coupling of TRPC3 channels to IP3 receptors in smooth muscle cells regulates arterial contractility, Circ. Res. 106 (2010) 1603–1612. [4] W.J. Arendshorst, K. Brannstrom, X. Ruan, Actions of angiotensin II on the renal microvasculature, J. Am. Soc. Nephrol. 10 (Suppl. 11) (1999) S149–S161. [5] D.J. Beech, Integration of transient receptor potential canonical channels with lipids, Acta Physiol. 204 (2012) 227–237. [6] M.J. Berridge, Inositol trisphosphate and calcium signalling, Nature 361 (1993) 315–325. [7] M.J. Berridge, Calcium signalling and cell proliferation, Bioessays 17 (1995) 491–500. [8] C.J. Blankley, J.C. Hodges, S.R. Klutchko, R.J. Himmelsbach, A. Chucholowski, C.J. Connolly, S.J. Neergaard, M.S. Van Nieuwenhze, A. Sebastian, J. Quin III, Synthesis and structure-activity relationships of a novel series of non-peptide angiotensin II receptor binding inhibitors specific for the AT2 subtype, J. Med. Chem. 34 (1991) 3248–3260. [9] J.S. Cameron, Lupus nephritis, J. Am. Soc. Nephrol. 10 (1999) 413–424.

3 24 (2014) 9 2 –10 4

[10] R.L. Chevalier, Developmental renal physiology of the low birth weight pre-term newborn, J. Urol. 156 (1996) 714–719. [11] B. Chini, M. Parenti, G-protein coupled receptors in lipid rafts and caveolae: how, when and why do they go there?, J. Mol. Endocrinol. 32 (2004) 325–338. [12] H. Cho, T. Kozasa, C. Bondjers, C. Betsholtz, J.H. Kehrl, Pericytespecific expression of Rgs5: implications for PDGF and EDG receptor signaling during vascular maturation, FASEB J. 17 (2003) 440–442. [13] G.M. Ciuffo, M. Viswanathan, A.M. Seltzer, K. Tsutsumi, J.M. Saavedra, Glomerular angiotensin II receptor subtypes during development of rat kidney, Am. J. Physiol. 265 (1993) F264–F271. [14] J. Couet, S. Li, T. Okamoto, T. Ikezu, M.P. Lisanti, Identification of peptide and protein ligands for the caveolin-scaffolding domain. Implications for the interaction of caveolin with caveolaeassociated proteins, J. Biol. Chem. 272 (1997) 6525–6533. [15] D. Derossi, A.H. Joliot, G. Chassaing, A. Prochiantz, The third helix of the Antennapedia homeodomain translocates through biological membranes, J. Biol. Chem. 269 (1994) 10444–10450. [16] K. Dreja, M. Voldstedlund, J. Vinten, J. Tranum-Jensen, P. Hellstrand, K. Sward, Cholesterol depletion disrupts caveolae and differentially impairs agonist-induced arterial contraction, Arterioscler. Thromb. Vasc. Biol. 22 (2002) 1267–1272. [17] J. Du, S. Sours-Brothers, R. Coleman, M. Ding, S. Graham, D.H. Kong, R. Ma, Canonical transient receptor potential 1 channel is involved in contractile function of glomerular mesangial cells, J. Am. Soc. Nephrol. 18 (2007) 1437–1445. [18] A.M. Fra, E. Williamson, K. Simons, R.G. Parton, De novo formation of caveolae in lymphocytes by expression of VIP21-caveolin, Proc. Natl. Acad. Sci. U.S.A 92 (1995) 8655–8659. [19] A. Fujita, J. Cheng, K. Tauchi-Sato, T. Takenawa, T. Fujimoto, A distinct pool of phosphatidylinositol 4,5-bisphosphate in caveolae revealed by a nanoscale labeling technique, Proc. Natl. Acad. Sci. U.S.A 106 (2009) 9256–9261. [20] J. Gao, J. Chao, K.J. Parbhu, L. Yu, L. Xiao, F. Gao, L. Gao, Ontogeny of angiotensin type 2 and type 1 receptor expression in mice, J. Renin–Angiotensin–Aldosterone Syst. 13 (2012) 341–352. [21] G. Garcia-Cardena, P. Martasek, B.S. Masters, P.M. Skidd, J. Couet, S. Li, M.P. Lisanti, W.C. Sessa, Dissecting the interaction between nitric oxide synthase (NOS) and caveolin. Functional significance of the nos caveolin binding domain in vivo, J. Biol. Chem. 272 (1997) 25437–25440. [22] S. Graham, M. Ding, S. Sours-Brothers, T. Yorio, J.X. Ma, R. Ma, Downregulation of TRPC6 protein expression by high glucose, a possible mechanism for the impaired Ca2þ signaling in glomerular mesangial cells in diabetes, Am. J. Physiol. Ren. Physiol. 293 (2007) F1381–F1390. [23] G. Grynkiewicz, M. Poenie, R.Y. Tsien, A new generation of Ca2þ indicators with greatly improved fluorescence properties, J. Biol. Chem. 260 (1985) 3440–3450. [24] M. He, L. Zhang, Y. Shao, H. Xue, L. Zhou, X.F. Wang, C. Yu, T. Yao, L.M. Lu, Angiotensin II type 2 receptor mediated angiotensin II and high glucose induced decrease in renal prorenin/renin receptor expression, Mol. Cell Endocrinol. 315 (2010) 188–194. [25] K.F. Hilgers, V.F. Norwood, R.A. Gomez, Angiotensin's role in renal development, Semin. Nephrol. 17 (1997) 492–501. [26] J.B. Hook, M.D. Bailie, Perinatal renal pharmacology, Annu. Rev. Pharmacol. Toxicol. 19 (1979) 491–509. [27] H. Hua, S. Munk, C.I. Whiteside, Endothelin-1 activates mesangial cell ERK1/2 via EGF-receptor transactivation and caveolin-1 interaction, Am. J. Physiol. Ren. Physiol. 284 (2003) F303–F312. [28] W.R. Huckle, H.S. Earp, Regulation of cell proliferation and growth by angiotensin II, Prog. Growth Factor Res. 5 (1994) 177–194. [29] N. Ishizaka, K.K. Griendling, B. Lassegue, R.W. Alexander, Angiotensin II type 1 receptor: relationship with caveolae and caveolin

E XP E RI ME N TAL CE L L R ES E ARC H

[30]

[31]

[32] [33]

[34]

[35]

[36]

[37]

[38]

[39]

[40]

[41]

[42] [43]

[44]

[45]

after initial agonist stimulation, Hypertension 32 (1998) 459–466. B.M. Iversen, F.I. Kvam, K. Matre, L. Morkrid, G. Horvei, W. Bagchus, J. Grond, J. Ofstad, Effect of mesangiolysis on autoregulation of renal blood flow and glomerular filtration rate in rats, Am. J. Physiol. 262 (1992) F361–F366. N.H. Jung, H.P. Kim, B.R. Kim, S.H. Cha, G.A. Kim, H. Ha, Y.E. Na, Y.N. Cha, Evidence for heme oxygenase-1 association with caveolin-1 and -2 in mouse mesangial cells, IUBMB Life 55 (2003) 525–532. E. Kaschina, T. Unger, Angiotensin AT1/AT2 receptors: regulation, signalling and function, Blood Press. 12 (2003) 70–88. E.P. Kilsdonk, P.G. Yancey, G.W. Stoudt, F.W. Bangerter, W.J. Johnson, M.C. Phillips, G.H. Rothblat, Cellular cholesterol efflux mediated by cyclodextrins, J. Biol. Chem. 270 (1995) 17250–17256. A.M. Kwiatek, R.D. Minshall, D.R. Cool, R.A. Skidgel, A.B. Malik, C. Tiruppathi, Caveolin-1 regulates store-operated Ca2þ influx by binding of its scaffolding domain to transient receptor potential channel-1 in endothelial cells, Mol. Pharmacol. 70 (2006) 1174–1183. K.N. Lai, L.Y. Chan, S.C. Tang, A.W. Tsang, F.F. Li, M.F. Lam, S.L. Lui, J.C. Leung, Mesangial expression of angiotensin II receptor in IgA nephropathy and its regulation by polymeric IgA1, Kidney Int. 66 (2004) 1403–1416. P.C. Leclerc, M. Auger-Messier, P.M. Lanctot, E. Escher, R. Leduc, G. Guillemette, A polyaromatic caveolin-binding-like motif in the cytoplasmic tail of the type 1 receptor for angiotensin II plays an important role in receptor trafficking and signaling, Endocrinology 143 (2002) 4702–4710. A.E. Linder, K.M. Thakali, J.M. Thompson, S.W. Watts, R.C. Webb, R. Leite, Methyl-beta-cyclodextrin prevents angiotensin II-induced tachyphylactic contractile responses in rat aorta, J. Pharmacol. Exp. Ther. 323 (2007) 78–84. E.M.M. Manders, F.J. Verbeek, J.A. Aten, Measurement of colocalization of objects in dual-colour confocal images, J. Microsc. 169 (1993) 375–382. K. Matsui, S. Breiteneder-Geleff, D. Kerjaschki, Epitope-specific antibodies to the 43-kD glomerular membrane protein podoplanin cause proteinuria and rapid flattening of podocytes, J. Am. Soc. Nephrol. 9 (1998) 2013–2026. W.W. Mederski, D. Dorsch, H.H. Bokel, N. Beier, I. Lues, P. Schelling, Non-peptide angiotensin II receptor antagonists: synthesis and biological activity of a series of novel 4,5-dihydro4-oxo-3H-imidazo[4,5-c]pyridine derivatives, J. Med. Chem. 37 (1994) 1632–1645. E. Mendez-Bolaina, J. Sanchez-Gonzalez, I. Ramirez-Sanchez, E. Ocharan-Hernandez, M. Nunez-Sanchez, E. MeaneyMendiolea, A. Meaney, J. Asbun-Bojalil, A. Miliar-Garcia, I. Olivares-Corichi, G. Ceballos-Reyes, Effect of caveolin-1 scaffolding peptide and 17beta-estradiol on intracellular Ca2þ kinetics evoked by angiotensin II in human vascular smooth muscle cells, Am. J. Physiol. Cell Physiol. 293 (2007) C1953–C1961. P. Mene, M.S. Simonson, M.J. Dunn, Physiology of the mesangial cell, Physiol. Rev. 69 (1989) 1347–1424. T. Okamoto, A. Schlegel, P.E. Scherer, M.P. Lisanti, Caveolins, a family of scaffolding proteins for organizing “preassembled signaling complexes” at the plasma membrane, J. Biol. Chem. 273 (1998) 5419–5422. R.S. Ostrom, X. Liu, Detergent and detergent-free methods to define lipid rafts and caveolae, Methods Mol. Biol. 400 (2007) 459–468. S. Parpal, M. Karlsson, H. Thorn, P. Stralfors, Cholesterol depletion disrupts caveolae and insulin receptor signaling for metabolic control via insulin receptor substrate-1, but not for mitogenactivated protein kinase control, J. Biol. Chem. 276 (2001) 9670–9678.

324 (2014 ) 92 – 10 4

103

[46] H.H. Patel, F. Murray, P.A. Insel, Caveolae as organizers of pharmacologically relevant signal transduction molecules, Annu. Rev. Pharmacol. Toxicol. 48 (2008) 359–391. [47] F. Peng, B. Zhang, D. Wu, A.J. Ingram, B. Gao, J.C. Krepinsky, TGFbeta-induced RhoA activation and fibronectin production in mesangial cells require caveolae, Am. J. Physiol. Ren. Physiol. 295 (2008) F153–F164. [48] L.J. Pike, Lipid rafts: bringing order to chaos, J. Lipid Res. 44 (2003) 655–667. [49] P.E. Ray, G. Aguilera, J.B. Kopp, S. Horikoshi, P.E. Klotman, Angiotensin II receptor-mediated proliferation of cultured human fetal mesangial cells, Kidney Int. 40 (1991) 764–771. [50] P.E. Ray, L.A. Bruggeman, S. Horikoshi, G. Aguilera, P.E. Klotman, Angiotensin II stimulates human fetal mesangial cell proliferation and fibronectin biosynthesis by binding to AT1 receptors, Kidney Int. 45 (1994) 177–184. [51] B. Razani, S.E. Woodman, M.P. Lisanti, Caveolae: from cell biology to animal physiology, Pharmacol. Rev. 54 (2002) 431–467. [52] C. Ruster, G. Wolf, Renin–angiotensin–aldosterone system and progression of renal disease, J. Am. Soc. Nephrol. 17 (2006) 2985–2991. [53] P. Scheiffele, P. Verkade, A.M. Fra, H. Virta, K. Simons, E. Ikonen, Caveolin-1 and -2 in the exocytic pathway of MDCK cells, J. Cell Biol. 140 (1998) 795–806. [54] J. Schindelin, I. Arganda-Carreras, E. Frise, V. Kaynig, M. Longair, T. Pietzsch, S. Preibisch, C. Rueden, S. Saalfeld, B. Schmid, J.Y. Tinevez, D.J. White, V. Hartenstein, K. Eliceiri, P. Tomancak, A. Cardona, Fiji: an open-source platform for biological-image analysis, Nat. Methods 9 (2012) 676–682. [55] D. Schlondorff, The glomerular mesangial cell: an expanding role for a specialized pericyte, FASEB J. 1 (1987) 272–281. [56] D. Schlondorff, B. Banas, The mesangial cell revisited: no cell is an island, J. Am. Soc. Nephrol. 20 (2009) 1179–1187. [57] S. Sethi, F.C. Fervenza, Membranoproliferative glomerulonephritis: pathogenetic heterogeneity and proposal for a new classification, Semin. Nephrol. 31 (2011) 341–348. [58] S.K. Shenoy, R.J. Lefkowitz, Angiotensin II-stimulated signaling through G proteins and beta-arrestin, Sci. STKE 2005 (2005) cm14. [59] A.K. Shukla, K. Xiao, R.J. Lefkowitz, Emerging paradigms of betaarrestin-dependent seven transmembrane receptor signaling, Trends Biochem. Sci. 36 (2011) 457–469. [60] O. Soderberg, M. Gullberg, M. Jarvius, K. Ridderstrale, K.J. Leuchowius, J. Jarvius, K. Wester, P. Hydbring, F. Bahram, L.G. Larsson, U. Landegren, Direct observation of individual endogenous protein complexes in situ by proximity ligation, Nat. Methods 3 (2006) 995–1000. [61] U.J. Soh, J. Trejo, Activated protein C promotes protease-activated receptor-1 cytoprotective signaling through beta-arrestin and dishevelled-2 scaffolds, Proc. Natl. Acad. Sci. U.S.A 108 (2011) E1372–E1380. [62] H. Soni, A. Adebiyi, Pressor and renal regional hemodynamic effects of urotensin II in neonatal pigs, J. Endocrinol. 217 (2013) 317–326. [63] G. Sowa, M. Pypaert, W.C. Sessa, Distinction between signaling mechanisms in lipid rafts vs. caveolae, Proc. Natl. Acad. Sci. U.S.A 98 (2001) 14072–14077. [64] M.W. Steffes, R. Osterby, B. Chavers, S.M. Mauer, Mesangial expansion as a central mechanism for loss of kidney function in diabetic patients, Diabetes 38 (1989) 1077–1081. [65] J.D. Stockand, S.C. Sansom, Regulation of filtration rate by glomerular mesangial cells in health and diabetic renal disease, Am. J. Kidney Dis. 29 (1997) 971–981. [66] J.D. Stockand, S.C. Sansom, Glomerular mesangial cells: electrophysiology and regulation of contraction, Physiol. Rev. 78 (1998) 723–744. [67] P.C. Sundivakkam, A.M. Kwiatek, T.T. Sharma, R.D. Minshall, A.B. Malik, C. Tiruppathi, Caveolin-1 scaffold domain interacts

104

[68]

[69]

[70]

[71]

[72]

[73]

[74]

EX P ER I ME NTAL C E LL RE S E ARCH

with TRPC1 and IP3R3 to regulate Ca2þ store release-induced Ca2þ entry in endothelial cells, Am. J. Physiol. Cell Physiol. 296 (2009) C403–C413. O. Tamai, N. Oka, T. Kikuchi, Y. Koda, M. Soejima, Y. Wada, M. Fujisawa, K. Tamaki, H. Kawachi, F. Shimizu, H. Kimura, T. Imaizumi, S. Okuda, Caveolae in mesangial cells and caveolin expression in mesangial proliferative glomerulonephritis, Kidney Int. 59 (2001) 471–480. A. Thathiah, K. Horre, A. Snellinx, E. Vandewyer, Y. Huang, M. Ciesielska, K.G. De, S. Munck, S.B. De, Beta-arrestin 2 regulates Abeta generation and gamma-secretase activity in Alzheimer's disease, Nat. Med. 19 (2013) 43–49. R.M. Touyz, E.L. Schiffrin, Signal transduction mechanisms mediating the physiological and pathophysiological actions of angiotensin II in vascular smooth muscle cells, Pharmacol. Rev. 52 (2000) 639–672. M. Ushio-Fukai, R.W. Alexander, Caveolin-dependent angiotensin II type 1 receptor signaling in vascular smooth muscle, Hypertension 48 (2006) 797–803. M. Ushio-Fukai, L. Hilenski, N. Santanam, P.L. Becker, Y. Ma, K.K. Griendling, R.W. Alexander, Cholesterol depletion inhibits epidermal growth factor receptor transactivation by angiotensin II in vascular smooth muscle cells: role of cholesterol-rich microdomains and focal adhesions in angiotensin II signaling, J. Biol. Chem. 276 (2001) 48269–48275. Y. Wang, R. Mishra, M.S. Simonson, Ca2þ/calmodulin-dependent protein kinase II stimulates c-fos transcription and DNA synthesis by a Src-based mechanism in glomerular mesangial cells, J. Am. Soc. Nephrol. 14 (2003) 28–36. G. Wolf, E.G. Neilson, Angiotensin II as a renal growth factor, J. Am. Soc. Nephrol. 3 (1993) 1531–1540.

3 24 (2014) 9 2 –10 4

[75] L.L. Woods, R. Rasch, Perinatal ANG II programs adult blood pressure, glomerular number, and renal function in rats, Am. J. Physiol. 275 (1998) R1593–R1599. [76] B.D. Wyse, I.A. Prior, H. Qian, I.C. Morrow, S. Nixon, C. Muncke, T.V. Kurzchalia, W.G. Thomas, R.G. Parton, J.F. Hancock, Caveolin interacts with the angiotensin II type 1 receptor during exocytic transport but not at the plasma membrane, J. Biol. Chem. 278 (2003) 23738–23746. [77] Q. Xi, A. Adebiyi, G. Zhao, K.E. Chapman, C.M. Waters, A. Hassid, J.H. Jaggar, IP3 constricts cerebral arteries via IP3 receptormediated TRPC3 channel activation and independently of sarcoplasmic reticulum Ca2þ release, Circ. Res. 102 (2008) 1118–1126. [78] H. Xue, P. Yuan, L. Zhou, T. Yao, Y. Huang, L.M. Lu, Effect of adrenotensin on cell proliferation is mediated by angiotensin II in cultured rat mesangial cells, Acta Pharmacol. Sin. 30 (2009) 1132–1137. [79] L. Yu, M. Zheng, W. Wang, G.J. Rozanski, I.H. Zucker, L. Gao, Developmental changes in AT1 and AT2 receptor-protein expression in rats, J. Renin–Angiotensin–Aldosterone Syst. 11 (2010) 214–221. [80] H. Zeng, Y. Liu, D.M. Templeton, Ca2þ/calmodulin-dependent and cAMP-dependent kinases in induction of c-fos in human mesangial cells, Am. J. Physiol. Ren. Physiol. 283 (2002) F888–F894. [81] A. Zhang, G. Ding, S. Huang, Y. Wu, X. Pan, X. Guan, R. Chen, T. Yang, c-Jun NH2-terminal kinase mediation of angiotensin II-induced proliferation of human mesangial cells, Am. J. Physiol. Ren. Physiol. 288 (2005) F1118–F1124.