Epithelial to mesenchymal transition (EMT) is a complex cellular and molecular program typical of cancer cells by which epithelial cells lose their differentiated characteristics and acquire mesenchymal features, such as motility and invasiveness, that contribute to metastatic dissemination.1 A new study suggests the possibility of a flexible transition between epithelial-to-mesenchymal (EM) and mesenchymal-to-epithelial (ME), for which the term “EM plasticity” has been coined to indicate a dynamic interchange between epithelial and mesenchymal phenotype.2
Recently, EM plasticity has been implicated in the formation of fibrotic lesions after inflammatory or toxic injuries in several organs, including the kidney. Upon glomerular injury, podocytes undergo activation with loss of specific markers and gain of transitional features.3,4 Because terminally differentiated podocytes developmentally derive from the metanephric mesenchyme through ME transdifferentiation, it is not surprising that in disease conditions a process of reverse embryogenesis can occur.4 Actually, the same EM plasticity reported in cancer might take place when podocytes become matrix-producing cells and acquire motile function, which then translates into detachment from the basement membrane and migration to glomerular crescents. Cultured podocytes exposed to TGF-β express fewer slit diaphragm–associated proteins but increased levels of the mesenchymal marker desmin, produce interstitial matrix components, and upregulate the transcription factor Snail, favoring migration.4 Cell lineage tracing studies show that in anti–glomerular basement membrane GN, genetically tagged podocytes lose their molecular signature and migrate so as to contribute to crescents. Such acquired behavior could explain their presence in hyperplastic lesions of rats with progressive renal injury as well as patients with different podocytopathies.5,6
Among multiple signals that can initiate EMT in cancer, recruitment of β-arrestin to the G protein–coupled endothelin-A receptor (ETAR) upon activation by endothelin-1 (ET-1) has been found to induce ovarian cancer cell invasion through the stabilization of β-catenin and Snail.7–9 As for cancer cells, podocytes possess a fully functional ET system10 and represent a target of ET-1. In cultured podocytes, excessive ET-1 synthesis induced by protein overload modifies the cell phenotype, causing downregulation of the podocyte marker synaptopodin.11 Moreover, renal ET-1, which is upregulated in experimental and human chronic nephropathies, promotes podocyte injury and glomerular remodelling, thereby contributing to the progression of renal damage.10,12 In harmony with these studies are data showing that blockade of ETAR protects at the glomerular level through the prevention of podocyte loss, suggesting a role for ET-1, via ETAR, in fostering podocyte motility and detachment from the glomerular basement membrane.13 However, mediators or mechanisms underlying podocyte dysfunction in progressive kidney disorders have been poorly studied so far.
The recognized role of β-arrestin in the migratory behavior of cancer cells was taken here as a paradigm to evaluate, in vitro, whether ET-1, by binding to ETAR, promoted podocyte motility through β-arrestin-1 activation. The proposed pathway was then validated in doxorubicin (Adriamycin [ADR]) nephrosis, in which we found rather surprisingly but very consistently that ADR-treated rodents do develop hyperplastic lesions strongly reminiscent of crescents normally seen in rapidly progressive GN among humans.14
Results
ET-1 Promotes Podocyte Dysfunction and Migration via ETAR
Expression of synaptopodin and α-smooth muscle actin (α-SMA), taken as podocyte and mesenchymal markers, respectively, in cultured mouse podocytes exposed to ET-1 was evaluated. ET-1 reduced synaptopodin and increased α-SMA expression at 6 hours and even more so at 24 hours (Figure 1A). This translated into increased cell motility, as revealed by wound healing assay. ET-1 time-dependently increased podocyte migration compared with control medium (Figure 1B); the effect occurred via ETAR because it was prevented by the ETAR selective antagonist BQ123 (Figure 1B). Addition of BQ123 to podocytes incubated with control medium did not modify the number of migrated cells (Figure 1B).
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Figure 1: ET-1, via ETAR, promotes dysfunction and migration of mouse podocytes in vitro. (A) Representative images of synaptopodin and α-SMA expression in podocytes exposed to control medium or ET-1 (100 nM) for 6 and 24 hours. Nuclei were stained with DAPI (blue). Original magnification, ×630. (B) Migration was assessed by wound healing assay in control podocytes or after exposure to ET-1 alone (100 nM) or in the presence of the ETAR antagonist BQ123 (1 μM). Mean values±SEM of podocytes migrated into the wound track after 2, 6, 10, and 15 hours (n=10 for control and ET-1; n=5 for BQ123. *P<0.05; **P<0.01 versus control; °P<0.05; °°P<0.01 versus ET-1). (C) Podocytes were exposed to control medium or ET-1 for 6 hours and then processed for ETAR mRNA expression by real time RT-PCR. Data are mean±SEM (n=3 experiments). *P<0.05 versus control. (D) Podocytes stimulated with control medium or ET-1 for 24 hours and then processed for β-arrestin-1 (β-arr1) mRNA expression by real-time RT-PCR. Data are mean±SEM (n=3 experiments). *P<0.05 versus control.
ET-1 Induces Formation of Molecular Signaling Complex of ETAR, β-Arrestin-1, and Src in Podocytes
In cancer cells, upon ET-1 stimulation, ETAR associates with β-arrestin-1, leading to Src activation and the formation of ETAR/β-arrestin-1/Src kinase complex.9 We first assessed whether ET-1 affected ETAR and β-arrestin-1 mRNA expression in podocytes. Constitutive expression of ETAR and β-arrestin-1 was upregulated by ET-1 in podocytes (Figure 1, C and D). Of note, these cells did not express ETBR mRNA in resting conditions or after ET-1 exposure. Coimmunoprecipitation experiments using lysates from ET-1–treated or control podocytes showed that ET-1 promoted the association between ETAR, β-arrestin-1, and Src (Figure 2A), which was evident at 5 minutes and remained stable at 15 minutes. BQ123 prevented β-arrestin-1 and Src association, indicating that ETAR engagement by ET-1 is needed for β-arrestin-1 to scaffold a signaling complex with Src (Figure 2B).
Figure 2: ETAR and β-arrestin-1 form a molecular signaling complex with Src promoting EGFR transactivation in ET-1-treated podocytes. (A) The association of β-arrestin-1 (β-arr1) with ETAR and Src was studied in cell lysates from control and ET-1–-treated podocytes by immunoprecipitation (IP) assays using irrelevant IgG (negative control) or an anti–β-arr1 antibody followed by immunoblotting (IB) with anti-ETAR, anti-Src, and anti–β-arr1 antibodies. (B) The role of ETAR in the formation of the molecular signaling complex was assessed by IPs using irrelevant IgG or an anti–arr1 antibody in lysates from podocytes exposed to control medium, ET-1 (100 nM, 15 minutes) alone or in the presence of BQ123 (1 μM), followed by IB with anti–β-arr1 and anti-Src antibodies. (C) Cell lysates from podocytes incubated with control medium or ET-1 (100 nM) for 5, 15, or 30 minutes were immunoblotted with anti–phospho-Src (Y416), anti-Src, anti–phospho-EGFR (Y845), anti-EGFR, anti–phospho-AKT, anti-AKT, anti–phospho-p42/44 MAPK, anti-p42/44 MAPK, and anti-HSP70 antibodies. (D) IB analysis using anti–phospho-EGFR (Y845) and anti-EGFR antibodies in podocytes exposed to control medium or stimulated with ET-1 (100 nM, 15 minutes) in the presence or absence of PP1 (1 μM). EGFR phosphorylation status was also assessed in ET-1–treated podocytes silenced for β-arrestin-1 (si-β-arr1) by specific siRNA or transfected with scrambled siRNA (SCR). For all panels, data are representative of at least three experiments. Molecular mass is indicated in kilodaltons.
ETAR/β-Arrestin-1/Src Signaling Complex Promotes Epidermal Growth Factor Receptor Transactivation and Downstream Pathways
To explore whether the ETAR/β-arrestin-1/Src signaling complex induced epidermal growth factor receptor (EGFR) transactivation, the activation status of EGFR and its downstream signaling pathways were evaluated. Western blot analysis revealed that ET-1 induced an early increase in the phosphorylation of Src that was inhibited by BQ123 (Supplemental Figure 1A), as well as EGFR and its downstream effectors protein kinase B (AKT) and p42/44 mitogen-activated protein kinase (MAPK) (Figure 2C). Activation of Src led to EGFR transactivation as revealed by the marked reduction of phospho-tyrosine EGFR levels when the Src inhibitor PP1 was added to ET-1–treated podocytes (Figure 2D). β-Arrestin-1 was required for ET-1–induced EGFR transactivation because EGFR phosphorylation was suppressed in β-arrestin-1–silenced podocytes (Figure 2D). Eighty percent reduction of β-arrestin-1 mRNA and protein levels in transfected podocytes confirmed the specificity of small interfering RNA (siRNA) oligos (Supplemental Figure 1, B and C).
ETAR/β-Arrestin-1/Src Complex and EGFR Transactivation Are Required for β-Catenin Stability
Post-transcriptional modifications of β-catenin—as serine/threonine dephosphorylation and tyrosine phosphorylation accomplished by receptor tyrosine kinases, including EGFR—prevent its degradation and result in protein cytosol accumulation and nuclear translocation, leading to the transcription of genes related to cell motility.15–17 In ET-1–treated podocytes, β-arrestin-1 associated with β-catenin as early as 5 minutes (Figure 3A). Concomitantly, active protein accumulated, as detected by an antibody against non–serine-threonine phosphorylated nonubiquitinated β-catenin (Figure 3B). β-Catenin accumulation depended on the formation of ETAR/β-arrestin-1 complex because it was abolished by both BQ123 (Figure 3B) and β-arrestin-1 silencing (Figure 3C). ET-1–promoted serine/threonine dephosphorylation (Supplemental Figure 2A) was associated with increased levels of β-catenin Tyr phosphorylation (Figure 3D), further supporting the role of ET-1 in preventing degradation of β-catenin. Activation of β-catenin required Src and EGFR kinase activities as inhibitors of Src (PP1), and EGFR (AG1478) prevented tyrosine phosphorylation (Figure 3, D and E) and accumulation of active β-catenin (Figure 3F). Addition of AG1478 to ET-1–treated podocytes inhibited EGFR phosphorylation (Supplemental Figure 2B).
Figure 3: The complex ETAR/β-arrestin-1/Src and the EGFR transactivation are required for β-catenin stability. (A) Cell lysates from control or ET-1–treated podocytes were immunoprecipitated with irrelevant IgG (negative control) or anti–β-arrestin-1 (β-arr1) antibody and analyzed by immunoblotting (IB) using anti–β-catenin (β-cat) and anti–β-arr1 antibodies. (B) Active β-cat was assessed in podocytes exposed to control medium, ET-1 (100 nM) alone or with BQ123 (1 μM) by IB. Anti-HSP70 antibody was used to confirm equal protein loading. (C) Active β-cat in podocytes transfected with scrambled (SCR) or β-arrestin-1 (si-β-arr1) siRNA before exposure to ET-1 (100 nM) for 15 minutes. Anti-HSP70 antibody was used to confirm equal protein loading. (D) Lysates of podocytes treated with control medium, ET-1 (100 nM, 15 minutes) alone or in the presence of PP1 (1 μM) were immunoprecipitated with irrelevant IgG or anti–β-cat antibody and analyzed by IB using an antiphosphorylated tyrosine (pTYR) and anti–β-cat antibodies. (E) Lysates of podocytes treated with control medium, ET-1 (100 nM, 15 minutes) alone or in the presence of AG1478 (1 μM) were immunoprecipitated with irrelevant IgG or anti–β-cat antibody and analyzed by IB using anti-pTYR and anti–β-cat antibodies. (F) Accumulation of active β-cat was evaluated in podocytes exposed to control medium, ET-1 (100 nM) alone or supplemented with AG1478 (1 μM). Cell lysates were analyzed by IB using anti-active β-cat and anti-HSP70 antibodies. For all panels, data are representative of at least two experiments. Molecular mass is indicated in kilodaltons.
Podocyte Dysfunction in Response to ET-1 Is Dependent on β-Arrestin-1 and Requires Snail Activation
To understand whether β-arrestin-1 was crucial in mediating ET-1–induced podocyte phenotypic changes and motility, experiments with β-arrestin-1 siRNA were performed. β-Arrestin-1 knockdown almost normalized reduced synaptopodin mRNA levels (Figure 4A, left) and increased α-SMA expression (Figure 4A, right) of ET-1–treated podocytes as well as abrogated their migration (Figure 4B). Scrambled or β-arrestin-1 siRNA transfection did not significantly affect podocyte constitutive synaptopodin and α-SMA expression and migration (Supplemental Figure 3, A and B).
Figure 4: Podocyte dysfunction in response to ET-1 depends on β-arrestin-1 and requires Snail activation. (A) Real-time RT-PCR of synaptopodin mRNA expression (left) and quantification of α-SMA staining (right) in podocytes transfected with SCR or β-arrestin-1 (si-β-arr1) siRNA and then exposed to ET-1 for 6 hours. Data are mean±SEM (n=4 experiments for real-time RT-PCR, n=3–7 for immunofluorescence). **P<0.01 versus control; °P<0.05, °°P<0.01 versus ET-1+SCR. (B) Wound healing assay of podocytes transfected with SCR or si-β-arr1 siRNA and then exposed to ET-1. Mean values±SEM of podocytes migrated into the wound track after 2, 6, and 10 hours (n=4–7 experiments). **P<0.01 versus control; °°P<0.01 versus ET-1+SCR. (C) Real-time RT-PCR of Snail mRNA expression in podocytes transfected with SCR or si-β-arr1 siRNA and then exposed to ET-1 100 nM for 30 minutes. Data are mean±SEM (n=3 experiments). **P<0.01 versus control, °°P<0.01 versus ET-1+SCR. (D) Wound healing assay of podocytes transfected with SCR or Snail (si-Snail) siRNA and then exposed to ET-1 for 6 hours. Mean values±SEM of podocytes migrated into the wound track (n=6 for control and ET-1+SCR; n=10 for ET-1+si-Snail). **P<0.01 versus control; °°P<0.01 versus ET-1+SCR.
The role of β-arrestin-1 in activating Snail, the EMT key transcriptional factor in podocytes,4 was next assessed in Snail-silenced podocytes. Snail siRNA oligos reduced by 50% Snail mRNA compared with scrambled siRNA-transfected podocytes (data not shown). ET-1 caused the doubling of Snail mRNA levels in scrambled siRNA but not in β-arrestin-1 siRNA-transfected podocytes (Figure 4C). Increased Snail expression was responsible for ET-1–mediated podocyte migration as Snail silencing significantly reduced cell motility (Figure 4D, Supplemental Figure 3C).
ETAR Blockade Prevents Podocyte Abnormalities in Mouse ADR-Induced Nephropathy
Having established the pathways of ET-1–induced podocyte dysfunction in vitro, we asked whether or not they operated in ADR nephropathy. At 1 week, ADR-treated mice developed proteinuria (mean±SEM, 25.7±5.1 mg/d versus 4.62±2.3 mg/d in control; P<0.05) that remained elevated at 4 weeks (29.8±5.1 mg/d versus 4.13±1.7 mg/d; P<0.01). Sitaxsentan reduced proteinuria, although not significantly (17.6±4.9 mg/d). Morphometric analysis of renal specimens revealed that Wilms' tumor 1 (WT1)–positive cells per glomerulus were reduced in ADR-treated mice already at 1 week (Figure 5A, Supplemental Figure 4A) concomitant with increased body surface of the remaining podocytes (Figure 5A). Podocyte loss was confirmed by immunostaining of nestin, another podocyte marker, similarly reduced in glomeruli of ADR-treated mice compared with controls (Supplemental Figure 4, A and B). Sitaxsentan normalized both podocyte number and volume (Figure 5A, Supplemental Figure 4, A and B). Scanning electron microscopy images of podocytes of control mice showed several primary processes and numerous interdigitating foot processes that almost totally covered the capillary wall (Figure 5, B and C). In ADR-treated mice, interdigitating foot processes were only occasionally observed, and interdigitation area was dramatically reduced with foot process shortening and degradation. Concomitantly, an enlargement of podocyte body volume was detectable (Figure 5, B and C). Sitaxsentan ameliorated both podocyte foot process architecture and cell volume in ADR-treated mice (Figure 5, B and C).
Figure 5: ETAR blockade limits podocyte loss, appearance of hyperplastic lesions, and glomerulosclerosis in ADR-treated mice. (A) Podocyte number/glomerulus and the glomerular volume occupied by podocyte body were assessed by morphometric analysis of kidney sections stained for the podocyte marker WT1. Data are mean±SEM (n=4–6 animals/group). **P<0.01 versus control at 1 and 4 weeks; # # P<0.01 versus ADR at 1 and 4 weeks. (B and C) Representative scanning electron micrographic images of podocyte ultrastructure in control and ADR-treated mice at 4 weeks receiving or not receiving sitaxsentan. (D) The extent of synechiae, pseudo-crescents, and sclerosis was scored from 0 to 4 related to the percentage of glomerular tuft occupied by the lesions and then expressed as index. Data are mean±SEM (n=4–6 animals/group). **P<0.01 versus controls at 1 and 4 weeks; °P<0.05, °°P<0.01 versus ADR at 1 week; ## P<0.01 versus ADR at 4 weeks.
ETAR Blockade Prevents the Formation of Hyperplastic Lesions in ADR-Induced Nephropathy
One week after disease induction, mice developed bridges between parietal and visceral epithelium called synechiae (index: 0.65±0.12) that increased in number and extension after 4 weeks (index: 1.43±0.25) compared with age-matched controls (index: 1 week, 0.08±0.02; 4 weeks, 0.03±0.02) (Figure 5D, Supplemental Figure 4C, arrow). At 4 weeks, glomerular lesions evolved to pseudo-crescents with cell multilayers accumulating at the site of synechiae (index: 4 weeks, 0.30±0.09) (Figure 5D, Supplemental Figure 4C, boxed area). Lesions were characterized by cell proliferation assessed by increased Ki-67 expression (Supplemental Figure 5A) or extracellular matrix accumulation resulting in glomerulosclerosis (index: 1 week, 0.08±0.04; 4 weeks, 0.87±0.16) (Figure 5D). Sitaxsentan [4-chloro-3-methyl-5-(2-(2-(6-methylbenzo[d][1,3]dioxol-5-yl)acetyl)-3-thienylsulfonamido) isoxazole sodium salt] limited the number and extension of the lesions by reducing synechiae (index: 0.54±0.21), pseudo-crescents (index: 0.12±0.07), and glomerulosclerosis (index: 0.34±0.12) (Figure 5D), and normalizing cell proliferation (Supplemental Figure 5A).
ETAR Blockade Restores β-Arrestin-1 Glomerular Expression and Distribution
Since previous studies demonstrated that podocytes, by acquiring a migratory phenotype, contribute to the formation of crescentic lesions,5,6,18 we evaluated the role of β-arrestin-1 in podocyte dysfunction in ADR-treated mice. In control animals, β-arrestin-1 showed a fragmented signal localized in podocytes and along the Bowman capsule (Figure 6A, arrowhead). In ADR-treated mice, β-arrestin-1 staining considerably increased and colocalized with WT1 in podocytes present in the glomerular tuft (Figure 6A, arrowhead) and in hyperplastic lesions (Figure 6A, arrow). Sitaxsentan restored glomerular β-arrestin-1 expression to control levels (Figure 6A). Of note, β-arrestin-1 was also expressed by parietal epithelial cells of the Bowman capsule in control animals (Figure 6A). Costaining of β-arrestin-1 and the metanephric mesenchymal marker neural cell adhesion molecule (NCAM), which defines renal progenitors in the Bowman capsule, demonstrated the presence of such cells in the hyperplastic lesions of ADR-treated mice (Supplemental Figure 5B). Overexpression of β-arrestin-1 was associated with an intense Snail staining in glomeruli of ADR-treated mice that was almost absent in control mice. Treatment with sitaxsentan significantly reduced Snail expression (Figure 6B).
Figure 6: β-Arrestin-1 is overexpressed in podocytes of ADR-treated mice. (A) Immunofluorescence images of β-arrestin-1 (red) and the podocyte marker WT1 (green) in control and ADR-treated mice at 4 weeks receiving or not receiving sitaxsentan. Nuclei were stained with DAPI (blue). Arrowheads and arrows indicate β-arrestin-1 staining that colocalized with WT1 in podocytes of the glomerular tuft or in areas of hyperplastic lesions, respectively. (B) Representative images of Snail (red) and FITC-wheat germ agglutinin (WGA)-lectin (green) in control and ADR-treated mice at 4 weeks receiving or not receiving sitaxsentan. Nuclei were stained with DAPI (blue). The histogram shows Snail signal intensity graded on a scale of 0–3 (mean score±SEM) of at least 25 glomeruli/animal (n=4 animals/group). *P<0.001 versus control; °P<0.01 versus ADR at 4 weeks. For all panels, images are representative of three independent experiments. Scale bars are indicated.
Glomerular Expression of β-Arrestin-1 increases in Human Extracapillary GN
The relevance of the above findings to human disease was assessed in patients with extracapillary GN, characterized by crescentic lesions. Systolic and diastolic BP did not differ among patients (systolic BP: 131.7±3.5 mmHg; diastolic BP: 80.0±0.0 mmHg). Mean values of proteinuria and serum creatinine were 4.1±0.7 g/d and 5.5±1.0 mg/dl, respectively. Immunofluorescence analysis in control humans showed a fragmented pattern of β-arrestin-1 staining in the glomerular tuft and along the Bowman capsule (Figure 7A). In patients with extracapillary GN, β-arrestin-1 expression increased in the glomerular tuft and in the area of hyperplastic lesions (Figure 7A, arrowhead). Some cells in the lesions were podocytes, as revealed by the costaining of β-arrestin-1 with podocalyxin (Figure 7A, arrowhead) and α-actinin 4 in adjacent renal tissue sections of the same patient (Figure 7B, arrows).
Figure 7: β-Arrestin-1 is overexpressed in podocytes of patients with extracapillary glomerulonephritis. (A) Immunofluorescence images of β-arrestin-1 (red) and podocalyxin (green) in control subject and in patient with extracapillary GN. Representative pictures of colocalization (yellow) of β-arrestin-1 and podocalyxin at the sites of hyperplastic lesions (arrowheads). Nuclei were stained with DAPI (blue). (B) Representative photomicrograph of a renal section costained with FITC-WGA-lectin (green) and β-arrestin-1 (red). Alongside, a portion of glomerular tuft shows β-arrestin-1 (arrow) in podocytes as evidenced by the α-actinin 4 staining (red) assessed on serial renal sections of the same patient with extracapillary GN. Asterisks indicate capillary lumens and arrows indicate podocytes coexpressing both β-arrestin-1 and α-actinin 4. Images are representative of six control subjects and eight patients. Scale bars are indicated.
Discussion
Once conceived as negative regulators of G protein–coupled receptors, β-arrestins have been shown to act as scaffold proteins that, by interacting with numerous signaling networks, favor peculiar biologic responses as cell proliferation and differentiation.19,20 Here we show that the molecular mechanism involving β-arrestin in cancer cell invasiveness and metastatic activity9,21 is a shared paradigm that dictates glomerular podocyte dysfunction. Our first observation was that ET-1 caused phenotypic changes in cultured podocytes that lose the synaptopodin differentiation marker and acquired the mesenchymal marker α-SMA. Concomitantly, ET-1 affected podocyte motility through the activation of the sole ET receptor expressed by cultured murine podocytes, the ETAR.11 Previous studies showed that ET-1 alters the podocyte contractile apparatus, causing F-actin redistribution through phosphatidylinositol 3-kinase/Rho kinase.22 Crucial for ET-1–induced phosphatidylinositol 3-kinase activation is β-arrestin-1 by virtue of its ability to hold together ETAR and the Src kinase c-Yes, resulting in the activation of the latter.23 In mouse podocytes, ET-1 instigated a similar cascade of events resulting in the formation of the trimeric functional complex of ETAR, β-arrestin-1, and Src kinase. This complex promoted EGFR transactivation, leading to β-catenin translocation into the nucleus,9,24 which favored transcription of Snail25 responsible for ET-1–dependent migratory phenotype. Finding that β-arrestin-1 knockdown impaired the above signaling pathway and prevented ET-1–mediated podocyte dysfunction and motility would indicate the key role of β-arrestin-1 in podocyte phenotypic changes.4 The present results might have important implications in proliferative disorders and indicate that the ETAR/β-arrestin-1/Src complex is a checkpoint controlling pathogenetic pathways as well as a possible target to prevent podocyte plasticity.
Results from in vitro studies were the rationale for assessing the effect of inhibiting ETAR in ADR nephropathy, one of the first animal models of nephrotic syndrome in the literature.26 Of interest, in ADR nephropathy, which exhibits increased renal ET-1,27 sclerotic lesions are consistently preceded by activation of parietal epithelial cells and pseudo-crescent formation.14 In the present study, podocyte depletion occurred early and induced the remaining podocytes to cover the denuded glomerular basement membrane, leading to an increase in their volume and shortening of foot processes. Normalization of podocyte number and volume by sitaxsentan was accompanied by a partial amelioration of primary and secondary interdigitating processes by scanning electron microscopy. This would possibly explain why despite the normalization of podocyte number, sitaxsentan did not significantly decrease proteinuria, and it highlights the importance of combining three-dimensional morphologic and morphometric analyses to actually visualize podocyte damage/recovery and estimate the degree of protection afforded by a given drug. Lack of a significant effect of sitaxsentan on proteinuria is not surprising. Previous data showed that ETAR blockade does not affect the dimension of pores perforating the glomerular barrier, which are increased in size in diabetic animals.13 Despite the presence of larger-than-normal pores, ETAR partially lowers urinary protein excretion because of an effect on renal hemodynamics.13 Indeed, proteinuria-lowering effects of ETAR antagonist reflect reduction in systemic and glomerular perfusion pressure due to a preferential constriction of renal efferent arteriole of ET-1 via ETAR.28 Furthermore, proteinuria decreases when ETAR antagonist is given in combination with an angiotensin-converting enzyme inhibitor, to which the antiproteinuric effect is attributable.13,29
Sitaxsentan limited the exaggerated proliferation of glomerular cells and migration to form pseudo-crescents, preventing the accumulation of extracellular matrix and the evolution toward glomerulosclerosis. Enhanced expression of β-arrestin-1 in glomerular cells in ADR nephropathy, conceivably induced by increased renal ET-1 production, is very reminiscent of the observation in podocytes exposed to ET-1. Restoration of β-arrestin-1 signal and podocyte motility by sitaxsentan can be taken to suggest that the mechanism underlying podocyte migration in vivo could operate through ET-1 viaβ-arrestin-1. NCAM+ cells, previously identified as a population of progenitor cells in the adult rat glomeruli, also overexpressed β-arrestin-1 in ADR-treated rats and localized in the Bowman capsule and in the hyperplastic lesions.5 Sitaxsentan lowered β-arrestin-1 expression in progenitor cells, which opens the possibility that migration of these cells in hyperplastic lesions might be regulated by ET-1 through β-arrestin-1 and can be pharmacologically modulated. The role of β-arrestin-1 on progenitor cell fate is beyond the scope of the present paper and would merit ad hoc investigation.
A different role of β-catenin in normal and pathologic conditions has recently been proposed. Although β-catenin is morphologically and functionally dispensable for normal adult podocytes, β-catenin–dependent pathways are instrumental to mediate podocyte dedifferentiation and glomerular scarring in ADR nephropathy through the canonical, Wnt-dependent activation.30,31 The present study supports the notion that β-catenin activation contributes to podocyte plasticity in ADR-induced nephrosis, although through an ET-1–driven, Wnt-independent pathway.
The translational relevance of this study rests on the similarity of β-arrestin-1 localization in the kidney of mice with ADR nephropathy and patients with extracapillary GN. Whether β-arrestin-1 activation is induced by ET-1 or other stimuli through their corresponding G protein–coupled receptors cannot be demonstrated in human samples because of the paucity of human trials using ET receptor antagonists. Our results underline novel analogies between podocytes and cancer cells and would explain why, until recently, the sole effective therapeutic option for reducing crescentic lesions in extracapillary GN is represented by the anticancer agent cyclophosphamide.
These data offer a new perspective for understanding undisclosed cell signaling that may be important for podocyte dysfunction, and they raise the possibility that ETAR antagonists may be of therapeutic value for treating crescentic and other types of inflammatory glomerulonephritides.
Concise Methods
Cell Culture and Incubations
Immortalized mouse podocytes (obtained from Dr. Peter Mundel, Department of Medicine, Albert Einstein College of Medicine, New York, NY) were grown and differentiated as described elsewhere,11 and then incubated with RPMI 1640 medium alone or with 100 nM ET-1 for 5, 15, or 30 minutes. The signaling pathways were investigated by adding the selective ETAR antagonist BQ123 (1 μM; Sigma-Aldrich, St. Louis, MO), the EGFR inhibitor AG1478 (1 μM; Calbiochem, Merck Chemicals Ltd., Nottingham, UK), or the Src inhibitor PP1 (1 μM; Calbiochem).
Immunofluorescence Analysis on Cultured Cells
Podocytes were fixed in 2% paraformaldehyde (Electron Microscopy Science, Hatfield, PA) and 4% sucrose (Sigma-Aldrich, Milan, Italy) and then permeabilized with 0.3% Triton X-100 (Sigma-Aldrich). After the nonspecific binding sites were blocked, cells were incubated with mouse anti-synaptopodin antibody (undiluted; Progen Immunodiagnostica, Heidelberg, Germany), followed by rabbit anti-mouse cy3-conjugated secondary antibody (Jackson ImmunoResearch Laboratories, West Grove, PA), or with mouse cy3-conjugated anti–α-SMA antibody (1:500; Sigma-Aldrich). Nuclei were stained with 4′,6-diamidino-2-phenylindole (DAPI) (Sigma-Aldrich), and negative controls were obtained by omitting primary antibodies. Fluorescence was examined by an inverted confocal laser scanning microscope (LS 510 Meta; Carl Zeiss, Jena, Germany). The α-SMA–positive areas were quantified on 15 fields per sample randomly acquired. Digitized images were binarized using a threshold for areas of α-SMA staining, and the fluorescent area, calculated in pixels using the analysis software ImageJ 1.38×, was then normalized for the number of cell nuclei identified by DAPI staining.
Wound Healing Assay
Migration of differentiated podocytes was assessed by wound healing assay.32 Cells were monitored by phase-contrast (original magnification, ×10) on time-lapse microscopy (Axio Imager.z2 microscope; Carl Zeiss). The number of cells that migrated into the wound track was counted (n=5 fields/well) at different time intervals.
Immunoblotting and Immunoprecipitation
Cell were detached, collected by centrifugation, and lysed in lysis buffer (250 mM NaCl, 50 mM HEPES [pH, 7.4], 1 mM EDTA, 1% Nonidet P-40, protease inhibitors). Equal amounts of whole cell lysates were separated by SDS/PAGE and transferred to polyvinylidene difluoride membranes that were blocked in Tris-buffered saline with 0.1% Tween 20 with 5% dry milk or BSA. Primary antibody incubations to β-arrestin-1 (K-16), β-catenin (E-5), ETAR, EGFR (Santa Cruz Biotechnology, Inc., Santa Cruz, CA), Src, phospho-Src (Tyr416), phospho-Akt (Ser473), AKT, pMAPK, MAPK, phospho-β-catenin (Ser33/37/Thr41) (Cell Signaling Technology, Danvers, MA), phospho-EGFR (Tyr845), active-β-catenin (antibody against non–serine-threonine phosphorylated, nonubiquitinated) (Upstate Biotech, Billerica, MA), HSP70 (Enzo Life Sciences, Lausen, Switzerland) were performed overnight at 4°C. The membranes were incubated with the appropriate secondary peroxidase-conjugated antibody, and immunoreactive proteins were visualized by using an enhanced chemiluminescence system (Amersham Pharmacia Biotech, Piscataway, NJ). For immunoprecipitation, precleared whole cell lysates were incubated with irrelevant IgG (Santa Cruz Biotechnology) or anti–β-arrestin-1, anti–β-catenin antibodies, and protein A-agarose beads (Amersham Pharmacia Biotech) at 4°C overnight. Agarose beads were washed three times in cold lysis buffer. Then, the precipitates were boiled for 5 minutes in SDS loading buffer, loaded onto 10% SDS-PAGE, transferred to nitrocellulose membrane, and immunoblotted with different antibodies as before. For detection of coimmunoprecipitated β-arrestin-1, horseradish peroxidase–conjugated protein A (Pierce, Rockford, IL) was used as secondary antibody.33
RNA Interference
Subconfluent mouse podocytes were transfected with ON-TARGET plus SMART pool siRNA (400 pmol) specific for mouse β-arrestin-1 or mouse Snail or control nontarget siRNA using Lipofectamine according to the manufacturer’s instruction. Forty-eight hours after transfection, podocytes were exposed to 100 nM ET1–1 for different time periods and used for the experiments.
Real-Time PCR
ETA receptor, β-arrestin-1, synaptopodin, and Snail mRNA expression in mouse podocytes were determined by real-time PCR. Total RNA was isolated as previously described.22 Amplification was performed with ABI7300 Real-Time PCR System using TaqMan Universal PCR Master Mix (Applied Biosystems, Monza, Italy) and inventoried TaqMan assays of β-arrestin-1 gene (FAM/MBG probe Mm00617540m1), synaptopodin gene (FAM/MBG probe Mm03413333m1), or mouse β-actin endogenous control (VIC/MGB probe) according to the manufacturer’s instructions. To amplify cDNA of ETA and Snail, SYBR Green PCR Master Mix and the following primers were used: mouse ETA receptor (300nM) forward 5′-CTTGCGGATCGCCCTTAGT-3′ reverse 5′-TTTGCCACTTCTCGACGCT-3′; mouse Snail (300nM) forward 5′-CGACCCGGTGACCCCGACTA-3′ reverse 5′-GGAAGGTGAACTCCACACACGCT-3′; glyceraldehyde 3-phosphate dehydrogenase (300 nM) forward 5′-TCATCCCTGCATCCACTGGT-3′ reverse 5′-CTGGGATGACCTTGCCCAC-3′. cDNA content was calculated by ΔΔCt technique in each sample using the cDNA expression in untreated podocytes as calibrator.
Experimental Model of ADR-Induced Nephropathy
Male Balb/c mice (Charles River Laboratories Italia, Calco, Italy), 7–8 weeks old, were used. Animal care and treatment were in accordance with institutional guidelines in compliance with national (Decreto Legislativo n.116, Gazzetta Ufficiale suppl 40, 18 febbraio 1992, Circolare n.8, Gazzetta Ufficiale 14 luglio 1994) and international (EEC Council Directive 86/609, OJL358–1, December 1987; Guide for the Care and Use of Laboratory Animals, US National Research Council, 1996) laws and policies. Animal studies were approved by the institutional animal care and use committees of Mario Negri Institute, Milan, Italy. Animals were housed in a constant-temperature room with a 12-hour dark/light cycle and fed a standard diet. Disease was induced by a single dose of ADR (10.5 mg/kg; Pfizer Italia s.r.l, Latina, Italy) by tail-vein injection, as described.34 Six ADR-treated mice were euthanized at 1 week. Two groups of mice were treated from 1 to 4 weeks with ETAR antagonist sitaxsentan (Pfizer, Tadworth, UK) (100 mg/kg, in the drinking water, n=6) or water (n=9) as vehicle. Healthy mice euthanized at 1 (n=4) and 4 (n=7) weeks served as controls. Proteinuria was determined by the Coomassie method using a Cobas Mira auto-analyzer (Roche Diagnostic Systems, Basel, Switzerland).
Renal Histology
Three-micrometer sections of paraffin-embedded kidney samples were stained with periodic acid-Schiff reagent and observed by light microscopy (BH2-RFCA; Olympus, Melville, NY). At least 50 glomeruli were examined for each animal, and the extent of synechiae, pseudo-crescents, and glomerulosclerosis was expressed by giving a score from 0 to 4 related to the percentage of glomerular tuft occupied by the lesions (0, no lesions; 1, lesions affecting <25% of the glomerulus; 2, lesions affecting >25%–50% of the glomerulus; 3, lesions affecting >50%–75% of the glomerulus; 4, lesions affecting >75%–100% of the glomerulus) and the average index was then calculated as weighted mean.5
Immunofluorescence Analysis on Renal Tissue
Acetone-fixed cryosections were incubated with goat anti-human/anti-mouse β-arrestin-1 (1:50; Santa Cruz Biotechnology), rabbit anti-mouse WT1 (1:50, Santa Cruz Biotechnology), rabbit anti-mouse Snail (1:200; AbCam, Cambridge, UK), rabbit anti-mouse Ki-67 (1:200; AbCam), mouse anti-human podocalyxin (1:250; gift from Prof. Robert Atkins, Department of Nephrology, Monash Medical Centre, Clayton, VIC, Australia), rabbit anti-human α-actinin 4 (1:300; OriGene, Rockville, MD), rabbit anti-mouse NCAM (1:1000, Millipore, Temecula, CA) and rat anti-mouse nestin (1:500; AbCam), followed by the specific FITC or Cy3-conjugated secondary antibodies (Jackson Immunoresearch Laboratories). Nuclei were stained with DAPI, and negative controls were obtained by omitting primary antibodies on adjacent sections. Fluorescence was examined by an inverted confocal laser scanning microscope (LS 510 Meta). For Snail quantification, at least 25 glomeruli were examined for each animal and Snail signal intensity was graded on a scale of 0–3 (0, no staining; 1, spotted; 2, weak; 3, strong diffusion). For the proliferation marker Ki-67, at least 25 glomeruli for each animal were examined and the percentage of positive glomeruli was evaluated. Quantification of nestin positive area was evaluated in 15–20 glomeruli randomly selected in each section. Positive staining of nestin was quantified by using ImageJ software and expressed as a percentage of the total glomerular area.
Estimation of Podocyte Number per Glomerulus and Glomerular Volume per Podocyte
Glomerular podocytes were identified using antibody anti-WT1, a podocyte-specific marker. The average number of podocytes per glomerulus was determined in 30 glomeruli for each animal by the stereologic method of particle density proposed by Weibel,35 on digital images acquired by confocal inverted laser microscope.36 The estimation of glomerular volume per podocyte was determined as the ratio between glomerular volume and number of podocytes per glomerulus.
Scanning Electron Microscopy
Mid coronal sections of kidneys were fixed in 2.5% glutaraldehyde, washed in cacodylate buffer, and post-fixed in 1% osmium tetroxide. Then, specimens were dehydrated, coated with atomic gold particles,37 and observed at scanning electron microscopy (Supra 55; Carl Zeiss).
Patient Enrollment
Renal tissues were obtained from biopsy specimens of eight patients with extracapillary GN who were admitted for diagnostic reasons to the Nephrology Unit of the “Azienda Ospedaliera Papa Giovanni XXIII” of Bergamo. The mean patient age was 49 years, and there was a slight male predominance. Demographic, clinical, and hematochemical variables at the time of renal biopsy were retrieved from the hospital database. Renal biopsy specimens from the uninvolved portion of the kidney, collected for tumor nephrectomy from six age- and sex-matched nonproteinuric patients, were used as normal controls. Written informed consent was obtained from all the patients enrolled in the study.
Statistical Analyses
Results are expressed as mean±SEM. Data were analyzed by ANOVA followed by Bonferroni multiple comparison test, or a t test for unpaired data, as appropriate. P<0.05 was considered to represent statistically significant differences.
Disclosures
None.
We thank Susanna Tomasoni for the helpful collaboration and discussion, Debora Conti for cell culture, Manuela Passera and Antonella Piccinelli for preparing the manuscript, and Giovanna Barcella and Cinzia Calvi for animal care.
P.R. is a recipient of a fellowship from Fondazione Aiuti per la Ricerca sulle Malattie Rare (ARMR), Bergamo, Italy.
Published online ahead of print. Publication date available at www.jasn.org.
See related editorial, “Arrestin(g) Podocyte Injury with Endothelin Antagonism,” on pages 423–425.
This article contains supplemental material online at http://jasn.asnjournals.org/lookup/suppl/doi:10.1681/ASN.2013040362/-/DCSupplemental.
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