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Open Access

Peer-reviewed

Research Article

Upregulation of angiotensin-(1–7) formation in human podocytes – enzyme activity assay upon fluid flow shear stress

  • Debora Kaiser-Graf ,

    Roles Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Software, Validation, Visualization, Writing – original draft, Writing – review & editing

    debora.kaiser@charite.de

    Affiliation Charité – Universitätsmedizin Berlin, corporate member of Freie Universität Berlin and Humboldt-Universität zu Berlin, Institute of Clinical Pharmacology and Toxicology, Berlin, Germany

    ⨯
  • Oliver Domenig,

    Roles Investigation, Methodology, Resources, Writing – review & editing

    Affiliation Attoquant Diagnostics GmbH, Vienna, Austria

    ⨯
  • Marko Poglitsch,

    Roles Methodology, Resources

    Affiliation Attoquant Diagnostics GmbH, Vienna, Austria

    ⨯
  • Reinhold Kreutz,

    Roles Conceptualization, Project administration, Resources, Supervision, Writing – review & editing

    Affiliation Charité – Universitätsmedizin Berlin, corporate member of Freie Universität Berlin and Humboldt-Universität zu Berlin, Institute of Clinical Pharmacology and Toxicology, Berlin, Germany

    ⨯
  • Juliane Bolbrinker

    Roles Conceptualization, Formal analysis, Methodology, Project administration, Supervision, Writing – review & editing

    Affiliation Charité – Universitätsmedizin Berlin, corporate member of Freie Universität Berlin and Humboldt-Universität zu Berlin, Institute of Clinical Pharmacology and Toxicology, Berlin, Germany

    ⨯

Abstract

The renin-angiotensin system is known for its role in renal physiology by regulating renal hemodynamics and natriuresis. Overactivation of this system exerts pathological effects in the kidney, primarily mediated by its main effector peptide angiotensin II. Deleterious angiotensin II-mediated effects are counter-regulated at least partly by the biologically active renin-angiotensin system component angiotensin-(1–7). The processing of angiotensin II to angiotensin-(1–7) by prolylcarboxypeptidase, angiotensin converting enzyme 2, and prolyl endopeptidase is cell- and species-specific, with limited knowledge regarding its conversion in human podocytes under conditions of glomerular hyperfiltration. Since hyperfiltration plays a critical mechanistic role in albuminuria progression and chronic kidney disease, understanding the mechanisms underlying podocyte damage due to glomerular hyperfiltration is essential. Therefore, we investigated the conversion of angiotensin II to angiotensin-(1–7) in cultured human podocytes exposed to fluid flow shear stress and subsequently incubated with spiked angiotensin II. Mass spectrometry of cell lysates and supernatants was performed to evaluate the formation of angiotensin-(1–7). Contribution of the respective enzymes to angiotensin-(1–7) formation was assessed using selective inhibitors of prolylcarboxypeptidase, angiotensin converting enzyme 2, and prolyl endopeptidase. We detected increased angiotensin-(1–7) formation upon fluid flow shear stress in podocyte lysates, which was mainly dependent on prolylcarboxypeptidase activity. Our study contributes to a deeper understanding of the intraglomerular processing of angiotensin II towards the alternative renin-angiotensin system and its modulation upon fluid flow shear stress.

Citation: Kaiser-Graf D, Domenig O, Poglitsch M, Kreutz R, Bolbrinker J (2026) Upregulation of angiotensin-(1–7) formation in human podocytes – enzyme activity assay upon fluid flow shear stress. PLoS One 21(1): e0339874. https://doi.org/10.1371/journal.pone.0339874

Editor: Junzheng Yang, Guangdong Nephrotic Drug Engineering Technology Research Center, Institute of Consun Co. for Chinese Medicine in Kidney Diseases, CHINA

Received: April 14, 2025; Accepted: December 12, 2025; Published: January 9, 2026

Copyright: © 2026 Kaiser-Graf et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Data Availability: All relevant data are within the manuscript and its Supporting Information files.

Funding: The author(s) received no specific funding for this work.

Competing interests: Oliver Domenig and Marko Poglitsch had a paid employment at Attoquant Diagnostics GmbH within the last 5 years. This commercial affiliation does not alter our adherence to PLOS ONE policies on sharing data and materials.

Introduction

The renin–angiotensin system (RAS) is a key system for regulation of renal hemodynamics and homeostasis [1,2]. Overactivation of RAS leads to various intrarenal hemodynamic and cellular alterations. These changes include efferent arteriolar vasoconstriction, elevated intraglomerular pressure, and glomerular hyperfiltration (GH), potentially leading to a decline in kidney function [3–7]. GH plays a crucial mechanistic role in the progression of albuminuria and the worsening of chronic kidney disease (CKD) in the context of hypertension and diabetes [8–11]. Several studies emphasize the critical role of angiotensin II (Ang II) in both physiological and pathological conditions in the kidney including vasoconstriction, growth and fibrosis, inflammatory response, and oxidative stress, thereby modulating renal hemodynamics [12–16]. Angiotensin II receptor type 1 (AGTR1) mediates the majority of the pathophysiological effects of Ang II in the kidney, while Angiotensin II receptor type 2 (AGTR2) antagonizes several of the actions mediated by AT1R [17]. Upregulation of renal Ang II-mediated effects are associated with increased blood pressure and play a central role in the development and maintenance of hypertension independent from changes in circulating RAS [18,19].

Adaptation of glomerular hemodynamics results in an elevated glomerular filtration rate (GFR) and concomitantly higher ultrafiltrate flow within Bowman’s space, leading to increased fluid flow shear stress (FFSS) at the podocyte surface and to podocyte damage [20–25]. Applying FFSS on podocytes using a shear stress device represents an in vitro approach to mimic the increased ultrafiltrate flow in Bowman’s space as observed upon GH, including cellular responses such as actin cytoskeleton rearrangement and morphological alterations that contribute to podocyte injury [23–29]. Podocytes are fully differentiated epithelial cells that form the third layer of the glomerular filtration barrier and are damaged in most proteinuric kidney diseases [10,30,31]. During disease progression, albuminuria often correlates with markers of podocyte injury including foot process effacement, podocyte hypertrophy, and apoptosis [32–34]. The crucial role of Ang II in podocyte injury and the progression of diabetic and non-diabetic kidney disease has been extensively investigated [31,35–40]. Podocytes are directly affected by Ang II–mediated injury through altered expression and distribution of podocyte proteins [31,41,42]. Furthermore, Ang II indirectly contributes to podocyte damage by enhancing calcium influx and reactive oxygen species production, which leads to actin cytoskeleton disruption, filtration barrier dysfunction as well as apoptosis [38,41,43].

Ang II is further metabolized to angiotensin-(1–7) [Ang-1–7] by prolyl endopeptidase (PREP; EC 3.4.21.26), prolylcarboxypeptidase (PRCP; EC 3.4.16.2), and angiotensin converting enzyme 2 (ACE2; 3.4.17.23)) [12,17,44–46]. Ang-(1–7) is also directly formed through hydrolysis from angiotensin I (Ang I) by PREP and membrane metalloendopeptidase (neutral-endopeptidase, nephrilysin) [12,44,47–49]. The Ang-(1–7) peptide is known to be a biologically active component of the RAS, mediating signaling through the MAS1 proto-oncogene receptor (MAS1), AGTR2, and MAS related GPR family member D receptor (MrgD) [13,50].

The ACE2-Ang-(1–7)-MAS1 axis is considered a counter-regulatory part of the ACE-Ang II-AGTR1 axis within the RAS as it modulates Ang II-mediated pathomechanisms [50]. In the kidney, Ang-(1–7) promotes vasodilation, inhibits growth, evokes anti-inflammatory responses and exerts antihypertensive and antifibrotic effects through the production of cAMP and nitric oxide (NO), activation of protein tyrosine phosphatases (PTPs), and inhibition of mitogen-activated protein kinases (MAPKs) and NADPH oxidases [6,12,13,46,51].

Reduced ACE2 activity, and consequently decreased Ang-(1–7) production, is associated with vasoconstriction, vascular remodeling, and oxidative injury in hypertension, diabetes, and kidney disease [47,52–55]. Therefore, activation of components of the ACE2-Ang-(1–7)-MAS1 axis are considered potential therapeutic targets [16,41,53].

Understanding the mechanisms and pathways underlying podocyte damage due to GH is essential for identifying potential new therapeutic targets. Thus, our objective was to clarify the regulation of the alternative and potentially renoprotective Ang-(1–7) formation in the context of elevated FFSS in human podocytes.

Methods

Cell culture

Conditionally immortalized human podocytes (hPC) [56,57] were kindly provided in June 2019 by Moin A. Saleem (University of Bristol, UK) and were cultured as previously described [25]. Briefly, for proliferation cells were cultivated at 33 °C and 5% CO2 until 70–80% confluency in Roswell Park Memorial Institute (RPMI)-1640 medium (cat. no. BS.F1295, Bio&SELL, Feucht/ Nürnberg, Germany) supplemented with 1% Insulin-Transferrin-Selenium 100X (cat. no. 41400–045, Gibco, Grand Island, NY, US), 10% fetal bovine serum (FBS) (cat. no. F7524, Sigma, St. Louis, MO, US) and 1% ZellShield® (cat. no. 13–0150, Minerva Biolabs, Berlin, Germany). Cells were transferred to 38 °C and cultured until full confluency and proliferation arrest. For differentiation, cells were further cultivated at 38°C for another 14 days. Differentiated hPC phenotype was confirmed previously by detecting podocyte specific markers synaptopodin and nephrin on protein level [25].

RNA isolation and reverse transcription

Total RNA of hPC was isolated and DNase-treated using the RNeasy® Micro Kit (cat. no. 74004, Qiagen, Hilden, Germany) according to the manufacturer’s protocol. RNA quality was checked by 260/280 nm absorption ratio. cDNA synthesis was carried out on 2 µg of total RNA using the First Strand cDNA Synthesis Kit (cat. no. K1612, Thermo Fisher Scientific, Vilnius, Lithuania) following the manufacturer’s protocol. Expression analysis of RAS components in hPC was determined by polymerase chain reaction (PCR) (S1 File).

Immunofluorescence

Ang-(1–7) forming enzymes ACE2, PRCP and PREP were determined on protein level in hPC by immunofluorescence. Cells were seeded at 25 x 10³ cells per well in 8-chamber NUNC Lab-Tek II Chamber slides (cat. no. 154534, NUNC, Rochester, NY, US) and cultured for 48 h in RPMI-1640 medium with supplements for adherence. Medium was discarded and cells washed twice with phosphate buffered saline (PBS, cat. no. 14190094, Thermo Fisher Scientific, Waltham, MA, US). Cells were fixed with methanol (cat. no. 8045, JK.T. Baker, Deventer, Netherlands) for 5 min at −20 °C. Cells were then allowed to dry completely before being rehydrated and washed three times with PBS. Blocking solution consisting of PBS with 2% FBS was applied for 30 min at room temperature. Cells were incubated with primary antibody polyclonal rabbit anti-ACE2 (AB_2792286, cat.no. PA5–85139, Invitrogen, Rockford, IL, US) diluted 1:100, polyclonal rabbit anti-PRCP (AB_1855676, cat. no. HPA 017065, BIOZOL, Eching, Germany) diluted 1:1000, or polyclonal rabbit anti-PREP (AB_2900433, PA5–115798, Invitrogen, Rockford, IL, US) diluted 1:200 for 1 h at room temperature. After three washing steps, secondary antibody polyclonal goat anti-rabbit IgG Alexa Fluor™ Plus 488 (AB_2633280, cat. no. A32731, Invitrogen, Rockford, IL, US) diluted 1:2000 was added and incubated for 1 h at room temperature in darkness. Nuclei were stained with 4’,6‐diamidino‐2‐phenylindole (DAPI, cat. no. D1306, Life Technologies, Carlsbad, CA, US) diluted 1:5000 for 10 min in darkness. Cells were washed in PBS and covered with Vectashield® mounting medium (cat. no. H-1000, Vector Laboratories, Burlingame, Ca, US). Immunofluorescence was detected at 405 nm (DAPI), and 488 nm (goat anti-rabbit for ACE2, PRCP and PREP) on a Zeiss axiovert 200 microscope (Carl Zeiss Microscopy GmbH, Jena, Germany) using ZEN blue v.3.4.91.0 software (Carl Zeiss Microscopy GmbH, Jena, Germany). Additionally, ACE2 and PRCP were determined on protein level in hPC by western blot (S1 File).

Inhibition of Ang-(1–7) formation by different inhibitors

Ang-(1–7) formation from substrate Ang II by ACE2, PRCP, and PREP was investigated in the presence of five different inhibitors. Inhibition of enzyme activity was tested for the ACE2 inhibitor MLN-4760 (ACE2i) (cat. no. 305335-31-3, Sigma-Aldrich, St. Louis, MO, US), the PRCP inhibitor PRCPi (cat. no. 5.04044.0001, calbiochem, San Diego, CA, US), PREP inhibitor S17092 (PREPi1) (cat. no. SML0181, Sigma-Aldrich, Saint Louis, MO, US), PREP inhibitor KYP-2047 (PREPi2) (cat. no. 6272, Tocris Bioscience, Bristol, UK), and the dual PREP and PRCP inhibitor Z-Pro-Prolinal (dual PREPi + PRCPi) (ZPP, cat. no. 88795-32-8, Sigma Aldrich, St. Louis, MO, US). 10 ng/mL recombinant human PRCP (cat. no. 7164-SE, R&D Systems, Minneapolis, MN, US), 10 ng/mL recombinant human PREP (cat. no. 4308-SE, R&D Systems, Minneapolis, MN, US), and 10 ng/mL recombinant human ACE2 (cat. no. 933-ZN, R&D Systems, Minneapolis, MN, US) were incubated in presence of increasing ACE2i-, PRCPi-, PREPi1-, PREPi2-, and dual PREPi + PRCPi-concentrations [0.01 µmol/L – 100 µmol/L] for 1 h at 37 °C. Inhibitor solutions (pH 7.4) consisted of 10 µmol/L amastatin (cat. no. 4025800, Bachem, Bubendorf, Switzerland) and 10 µmol/L lisinopril (cat. no. L6292, Sigma-Aldrich, St. Louis, MO, US), as well as 5 mmol/L ZnCl2 (cat.no. 703516, Sigma-Aldrich, St. Louis, MO, US) and 1 µg/mL Ang II as substrate (A9525, Sigma-Aldrich, St. Louis, MO, US) (pH 7.4). Subsequently, adjustments in the experimental setting were made for the inhibitory effect of ACE2i on ACE2 by lowering the amount of recombinant ACE2–1 ng/mL and for the inhibitory effect of PRCPi on PRCP by lowering the PRCPi concentration to 0.01 nmol/L – 100 nmol/L. Inhibitor concentrations were log-transformed and data are plotted as Ang-(1–7) formation [ng/mL] over inhibitor concentrations. Log (IC50) values, IC50 values, and R-squared values (R2) were calculated using a non-linear regression (curve fit) of log(inhibitor) vs. response using GraphPad Prism 6.07 (GraphPad Software, San Diego, CA, US).

FFSS

For FFSS, cells were detached with Trypsin 0.25%/EDTA 0.02% solution (cat. no. P10-023100, PAN-Biotech, Aidenbach, Germany). 3.5 × 105 cells were seeded on collagen IV coated Culture Slips® (cat. no. CS-C/IV, Dunn Labortechnik GmbH, Asbach, Germany) and adhered for 48 hours. FFSS experiments were performed as previously described with minor changes [25]. Briefly, Culture Slips® with hPC were inserted in the Streamer® Shear Stress Device (cat. no. STR-400, Dunn Labortechnik GmbH, Asbach, Germany) and the system was placed in the incubator at 38 °C with 5% CO2. The Streamer® Shear Stress Device ensured a constant medium flow over the hPC surface the entire time. For the experiments, FFSS at 1 dyne/cm2 for 2 h was applied based on previous research [21,24,27]. Control cells were cultured in the incubator simultaneously without FFSS exposure. All experiments were performed in supplement-free RPMI-1640 medium with cell passages between 7 and 15.

For determination of Ang-(1–7) in cell lysates, cells were released from the device at the end of the experiments, washed twice with PBS and stored immediately at −80°C until further use. For the Ang-(1–7) formation analyses in hPC supernatants, cells were removed from the device, washed with 38 °C supplement-free RPMI-1640 medium and processed immediately. Experiments were conducted twice on different cell passages and different dates, each time with n = 6 replicates for cell lysates analyses and n = 4 replicates for analyses of supernatants, respectively.

Quantitative real-time PCR

Quantitative Real-Time PCR (qPCR) analyses were performed in CFX96™ Real-Time system (Bio-Rad, München, Germany, software version 3.1.1517.0823) using the comparative quantitative cycle method as reported [58,59]. Expression analysis of each sample was done in three technical replicates and only samples with an intra-triplicate standard deviation (SD) < 0.2 were used for further calculation. We assessed mRNA expression of ACE2 and PRCP. Primer sequences are listed in S1 Table. Normalization of expression data was done by the reference gene glucuronidase beta (GUSB). All experiments for qPCR analyses were performed in quadruples, each time with n = 4–6 replicates.

Determination of Ang-(1–7) formation in hPC upon FFSS

Cell lysates sample preparation.

Ang-(1−7) formation was determined in presence and absence of ACE2i, with and without concomitant PRCPi in hPC lysates after applying FFSS. Cells were scratched from slides and transferred to a vial containing PBS. Cell suspensions were sonicated for dissolution using Vibra-Cell™ (Sonics & Materials Inc., Newtown, CT; US) and centrifugated at 16260 rcf for 10 min at 4°C. Supernatants were collected and normalized to total protein concentration determined using BIO-RAD Protein Assay Dye with bovine serum albumin as protein standard (cat. no. 5000006, Hercules, CA, US). Samples normalized to 200 µg total protein/mL were diluted in 100 µL inhibitor solution containing Ang II as substrate (Methods 4, see above) +/- 10 µmol/L ACE2i and +/- 10 µmol/L PRCPi and incubated for 1 h at 37 °. Thereafter, samples were acidified for stabilization using formic acid (10%, cat. no. 64-18-6, Sigma-Aldrich, St. Louis, MO, US).

Cell supernatants sample preparation.

Ang-(1–7) formation rate on hPC surface was determined in presence and absence of ACE2i or PRCPi after applying FFSS. For this, hPC covered slides were inserted into 8-chamber casettes (Visium Casette Assembly, cat. no. 1000388, 10x Genomics, Pleasanton, CA; US) directly after FFSS experiments enabling segmentation of the slides. Inhibitor solutions containing substrate Ang II (Methods 4, see above) +/- 10 µmol/L ACE2i or 10 µmol/L PRCPi were added to supplement-free RPMI-1640 medium. 100 µL inhibitor solutions were applied each into one chamber of the cassette onto hPC and incubated for 1 h at 37 °C. Supernatants were collected and, following stabilization by acidification, Ang-(1–7) formation was quantified by LC-MS/MS (Methods 7.3).

LC-MS/MS.

Stable isotope-labeled internal standards for Ang II and Ang-(1–7) were added to acidified samples at 200 pg/mL. Following C18-based solid-phase extraction, samples were subjected to LC-MS analysis using a reversed-phase analytical column (Acquity UPLC C18; Waters, Milford, Massachusetts) operating in line with a XEVO TQ-S triple quadrupole mass spectrometer (Waters) in MRM mode. Internal standards were used to correct for matrix effects and peptide recovery of the sample preparation procedure for both angiotensin peptides in each individual sample. Angiotensin peptide concentrations were calculated considering the corresponding response factors determined in appropriate calibration curves, on condition that integrated signals exceeded a signal-to-noise ratio of ten. Ang II concentration was measured in a non-incubated control sample as reference and after incubation in all samples to confirm that the substrate is not exhausted following incubation. Assay controls using recombinant enzymes were included to ensure that the inhibitor concentration was functional and to exclude non-specific formation of Ang-(1–7). Ang-(1–7) formation rate for cell lysates is expressed as (fmol/ µg protein)/ h and for supernatants as (nmol/ L)/ h. In cell lysates, the sole effect of PRCP inhibition on the Ang-(1–7) formation rate was calculated by subtracting remaining Ang-(1–7) formation rate upon ACE2 inhibition in each sample from the respective total value. This value was then summed to the ACE2i + PRCPi value of the respective sample which finally represents the individual PRCPi value.

Statistics

Statistical analysis was performed using GraphPad Prism 6.07. Data are presented as mean ± SD, and p < 0.05 was considered as statistically significant. Normal distribution was tested with the Shapiro-Wilk test. Normally distributed data were compared by either unpaired, two-tailed Student`s t-test or one-way ANOVA with Tukey`s multiple comparison test. Data not normally distributed were analyzed by Mann-Whitney test as indicated. For identification of outliers, Grubbs’ outliers test (α = 0.05) was performed.

Results

Characterization of angiotensin system components in hPC

Characterization of angiotensin system components in differentiated hPC revealed no angiotensinogen (AGT) and no renin (REN) gene expression (S1 Fig, panel A and B), whereas ACE, ACE2, PRCP, and PREP as well as AGTR1 gene expression was observed (S1 Fig, panel C-G). AGTR2 was only weakly and inconsistently expressed (S1 Fig, panel H). Immunofluorescence revealed expression of ACE2, PRCP, and PREP on protein level in hPC (Fig 1). Additional western blot analyses confirmed ACE2 and PRCP expression on protein level in hPC (S4 Fig, panel A-C).

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Fig 1. Characterization of ACE2, PRCP, and PREP on protein level in hPC conducted by immunofluorescence.

Representative immunofluorescence images of (A) ACE2, (B) PRCP, and (C) PREP, demonstrating protein expression on hPC. (D) Control without primary antibody revealed no specific binding of secondary antibody goat anti-rabbit Alexa Fluor™ Plus 488. Nuclei were stained using DAPI (blue). Scale bar 50 µm.

https://doi.org/10.1371/journal.pone.0339874.g001

Enzymes involved in Ang-(1–7) formation from substrate Ang II

To determine the contribution of the three enzymes ACE2, PRCP, and PREP reported to generate Ang-(1–7) from Ang II on Ang-(1–7) formation in hPC, we first investigated the specificity of five different enzyme inhibitors. Enzyme activities of ACE2, PRCP, and PREP were determined by Ang II to Ang-(1–7) conversion with increasing inhibitor concentrations (Fig 2 and S2 Fig).

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Fig 2. Inhibitor effects on ACE2-, PRCP-, and PREP-mediated Ang II to Ang-(1-7) conversion.

(A) Inhibition of Ang-(1-7) formation [ng/mL] by ACE2 (10 ng/mL) was tested with increasing inhibitor concentrations of five different inhibitors. (B) For a complete inhibition of Ang-(1-7) formation by ACE2, 1 ng/mL ACE2 was incubated with increasing ACE2i concentrations [0.01 μM- 100 μM]. (C) Inhibition of Ang-(1-7) formation by PRCP (10 ng/mL) was tested for five different inhibitors [0.01 μM- 100 μM]. (D) For PRCPi, lower inhibitor concentrations [0.01 nM- 100 nM] were used to investigate inhibitory effects on Ang-(1-7) formation by PRCP.

https://doi.org/10.1371/journal.pone.0339874.g002

ACE2i was the only inhibitor blocking ACE2 activity with a remaining Ang-(1–7) formation detected at the highest ACE2i-concentration tested (Fig 2A). Using 1 ng/mL ACE2, complete inhibition of Ang-(1–7) formation was achieved (IC50 = 508.2 nM, R2 = 0.9994; Fig 2B).

PRCPi had the strongest inhibitory effect on Ang II to Ang-(1–7) conversion by PRCP as 10 nM PRCPi already completely blocked the PRCP activity (Fig 2C). Thus, lower PRCPi concentrations (0.01 nM – 100 nM) were tested. Ang-(1–7) formation by PRCP was effectively inhibited by PRCPi with IC50 = 78.36 pM (R2 = 0.9997; Fig 2D). The dual PREPi + PRCPi as well as PREPi2 also had inhibitory effects on PRCP-derived Ang-(1–7) formation with IC50 of 228.4 nM and 8336 nM respectively (R2 = 0.9994, R2 = 0.9779; S3 Fig, panel B).

We observed no PREP-mediated Ang-(1–7) conversion of 1 ng/mL (S2 Fig, panel A). Thus, a higher (1 µg/mL) PREP enzyme concentration was tested to ensure that PREP is not involved in the formation of Ang-(1–7) from Ang II. Even with this high enzyme concentration, only a very weak PREP-activity and no inhibitory effects were detected (S2 Fig, panel B; S3 Fig, panel D).

Overall, the total amount of Ang-(1–7) formation was 6-times higher with ACE2 (mean 519 ng/mL) compared to PRCP (mean 81 ng/mL) indicating ACE2 as the most effective enzyme in Ang-(1–7) formation. Inhibitor assays revealed that Ang-(1–7) formation from the substrate Ang II was mediated by ACE2 and PRCP, but not by PREP.

Ang II to Ang-(1–7) conversion in hPC upon FFSS

Upon FFSS, there was a significantly elevated Ang II to Ang-(1–7) conversion rate in cell lysates of hPC with 720 ± 111 (fmol/µg protein)/h compared to no flow controls 554 ± 87 (fmol/µg protein)/h, p = 0.0007 (Fig 3A). In contrast to elevated Ang-(1–7) formation in cell lysates, no effect was observed on Ang-(1–7) formation rate at the cell surface of hPC (Fig 3B). These results suggest a higher enzyme activity and Ang II to Ang-(1–7) conversion capacity upon FFSS in hPC.

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Fig 3. Ang-(1-7) formation in hPC lysates and supernatants upon FFSS.

(A) Ang-(1-7) formation rate in cell lysates was elevated upon FFSS (blue squares) compared to no flow controls (black circles). Experiments were performed twice on different cell passages and different dates, each consisting of n = 6 technical replicates, respectively. Statistics: ***, p < 0.001, assessed by two-tailed Student´s t-test. (B) Ang-(1-7) formation rate in supernatants was not significantly regulated upon FFSS (blue squares) compared to no flow controls (black circles). Experiments were performed twice on different cell passages and different dates, each consisting of n = 4 technical replicates, respectively. Statistics: assessed by Mann-Whitney test. Each data point represents one technical replicate. Data are plotted as mean ± SD (horizontal lines).

https://doi.org/10.1371/journal.pone.0339874.g003

Gene expression analysis revealed a slightly upregulated ACE2 expression upon FFSS exposure compared to controls (p < 0.05; Fig 4A). In contrast, PRCP expression levels were slightly reduced upon FFSS compared to controls (p < 0.05; Fig 4B). Overall the enzymes involved in Ang II to Ang-(1–7) conversion were only moderately regulated upon FFSS.

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Fig 4. Gene expression of ACE2 and PRCP in hPC upon FFSS.

(A) ACE2 gene expression increased in hPC upon FFSS (blue squares) compared to no flow control (black circles). (B) PRCP gene expression decreased in hPC upon FFSS (blue squares) compared to no flow control (black circles). qPCR results are presented as relative mRNA expression level normalized to GUSB and referred to unstimulated control group. Each data point represents one technical replicate of four separate experiments. Experiments were performed on different cell passages and different dates, each experiment consisting of n = 5-6 technical replicates, respectively. Data are plotted as mean ± SD (horizontal lines). Statistics: *, p < 0.05, assessed by two-tailed Student´s t-test.

https://doi.org/10.1371/journal.pone.0339874.g004

Contribution of ACE2 and PRCP for Ang-(1–7) formation in hPC upon FFSS

Cell lysate.

To determine the proportion of ACE2 and PRCP involved in Ang-(1–7) formation rate from Ang II, hPC lysates were incubated with and without ACE2i and PRCPi. After FFSS exposure as well as under no flow control conditions, inhibition of ACE2 did not affect the Ang-(1–7) formation rate in hPC lysates compared to the total Ang-(1–7) formation rate. Inhibition of PRCP resulted in a significantly reduced Ang-(1–7) formation rate by at least half regardless of FFSS exposure or no flow control (p < 0.0001; Fig 5). Despite ACE2 and PRCP inhibition, a residual Ang-(1–7) formation rate remained, which was even higher under FFSS conditions compared to controls. However, Ang-(1–7) formation in hPC whole cell lysates is mainly driven by PRCP. In summary, in cell lysates, Ang-(1–7) formation by ACE2 was low, and most of the conversion was mediated by PRCP. Increased Ang-(1–7) formation rate upon FFSS cannot be attributed to ACE2 and PRCP, but rather indicates that another enzyme is involved.

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Fig 5. Ang-(1-7) formation rate in hPC lysates upon FFSS.

(A) Ang-(1-7) formation rate under no flow control condition and (B) after FFSS exposure. Inhibition of ACE2 (blue squares) did not affect the Ang-(1-7) formation rate in hPC cell lysates compared to the total Ang-(1-7) formation rate (black circles). Inhibition of PRCP (green triangles) significantly reduced the Ang-(1-7) formation compared to total Ang-(1-7) formation (black circles) as well as ACE2 inhibition (blue squares). Experiments were performed twice on different cell passages and different dates, each experiment consisting of n = 6 technical replicates, respectively. Each data point represents one technical replicate. Data are plotted as mean ± SD (horizontal lines). Statistics: ****, p < 0.0001, assessed by one-way ANOVA with Tukey`s multiple comparison follow-up test.

https://doi.org/10.1371/journal.pone.0339874.g005

Cell supernatant.

To determine the contribution of ACE2 and PRCP in Ang-(1–7) formation in cell supernatants, hPC were incubated with and without ACE2i and PRCPi. Under no flow control conditions, ACE2 inhibition (mean 3.75 nmol/L/h) reduced the Ang-(1–7) formation rate significantly compared to total Ang-(1–7) formation rate (mean 8.02 nmol/L/h) and compared to PRCP inhibition (mean 7.70 nmol/L/h) (Fig 6A). Upon FFSS, Ang-(1–7) formation rate with PRCP inhibition (mean 5.55 nmol/L/h) was no longer significantly different from Ang-(1–7) formation rate with ACE2 inhibition (mean 3.54 nmol/L/h) (Fig 6B). This suggests, besides ACE2 being the main enzyme responsible for Ang-(1–7) formation in supernatants, PRCP-derived Ang-(1–7) formation was enhanced upon FFSS (Fig 6B). Taken together, Ang-(1–7) formation rate in supernatants is predominantly driven by ACE2 but contribution of PRCP increases upon FFSS suggesting increased enzyme transport to the cell surface through exocytosis.

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Fig 6. Ang-(1-7) formation rate from Ang II in hPC supernatants upon FFSS.

(A) Under no flow control condition, Ang-(1-7) formation with ACE2 inhibition (blue squares) was significantly lower than total Ang-(1-7) formation (black circles) and formation with PRCP inhibition (green triangles). (B) Upon FFSS exposure, ACE2 inhibition (blue squares) reduced the Ang-(1-7) formation significantly compared to total Ang-(1-7) formation without inhibitors (black circles). Ang-(1-7) formation with PRCP inhibition (green triangles) was no longer significantly different to Ang-(1-7) formation upon ACE2 inhibition. Experiments were performed twice on different cell passages and different dates, each experiment consisting of n = 4 technical replicates, respectively. Statistics: **, p < 0.01; ***, p < 0.001; n.s., not significant, assessed by one-way ANOVA with Tukey`s multiple comparison follow-up test. Each data point represents one technical replicate. Data are plotted as mean ± SD (horizontal lines).

https://doi.org/10.1371/journal.pone.0339874.g006

Discussion

The intraglomerular RAS is implicated in the pathogenesis of progressive glomerular diseases, with podocyte injury playing a central role in glomerular damage and proteinuria. Renal pathologies are often associated with RAS activation and elevated levels of its primary effector, the vasoconstrictor Ang II [60–62]. The effects of Ang II on podocyte injury have been extensively studied and are associated with pathomechanisms underlying the progression of renal disease and hypertension (reviewed in [41]). In hPC, Ang II induces the production of ROS and promotes cellular apoptosis via the Arf6-Erk1/2-Nox4 signaling pathway [38]. The molecular mechanisms involved in Ang II-induced podocyte apoptosis are mediated by AGT1R and include the activation of endoplasmic reticulum stress, protein kinase C delta, and the p38 MAPK pathway [36]. In immortalized mouse podocytes, mechanical strain triggered the activation of local tissue RAS, leading to elevated Ang II levels and AT1R-mediated podocyte apoptosis [31].

Our initial characterization of RAS components in differentiated hPC revealed gene expression of ACE, ACE2, PRCP, PREP, AGTR1, and weak expression of AGTR2. However, we did not detect expression of AGT and REN, despite previous reports indicating that podocytes possess an intrinsic and local RAS, which enables them to produce Ang II endogenously [31,63]. Notably, the expression of RAS components in podocytes has been investigated in several studies, with results varying significantly, suggesting that the experimental models and conditions used strongly influence the findings. For instance, in mice, podocytes exhibited ACE2 on protein level, while ACE was exclusively detected in glomerular endothelial cells [64]. In contrast, Velez et al. demonstrated functional ACE activity in cultured immortalized mouse podocytes and provided evidence for a functional intrinsic podocyte RAS [65].

However, the primary source of renal renin production remains localized to juxtaglomerular cells [66]. By combining tissue-specific KO models with a model of inducible podocyte injury, Matsusaka et al. showed that increased renal Ang II in response to podocyte injury was due to the increased filtration of systemic AGT, resulting from the loss of glomerular barrier function [67,68]. According to them, hepatic AGT is filtered through the glomerular filtration barrier and then further converted intrarenally.

In the present study, we characterized the enzymes involved in conversion of Ang II to Ang-(1–7) in hPC upon FFSS, using pharmacological inhibition of the potentially involved converting enzymes and LC/MS-MS analysis. FFSS parameters to model pathological hyperfiltration stress were used based on previous research. FFSS at the podocyte surface in healthy adult mice has been estimated to be approximately 0.3 dyne/cm² [24]. In models of unilateral nephrectomy reflecting a hyperfiltrating state, FFSS on mouse podocytes increases by 1.5–2.5-fold, reaching ~0.5–0.75 dyne/cm² [21]. As reported by others, applying 0.5 dyne/cm² on human primary podocytes promoted expression of podocyte-specific markers, supporting a physiological relevance of this stress level in hPC [27]. Based on these data, we selected 1 dyne/cm² for 2 hours to model pathological hyperfiltration, representing a mild but sustained increase over normal shear stress, sufficient to induce mechanosensitive responses in glomerular cells without causing acute cell detachment and loss.

Starting by testing the specificity of several inhibitors revealed MLN-4760 as a highly specific ACE2 inhibitor with an IC50 of 508 nM. The dual PRCP and PREP inhibitor ZPP showed a higher efficacy for inhibiting PRCP with IC50 values of 228 nM and 5.3 µM, respectively. PRCPi was highly selective for inhibiting PRCP (IC50 = 78 pM). The PREP inhibitor KYP-2047 also displayed an inhibitory effect on PRCP activity with an IC50 of 8.3 µM and thus should not be considered a selective PREP inhibitor, especially when high inhibitor concentrations are used in the experimental setting. Our observations are consistent with those of Jalkanen et al., who previously reported a 20% inhibition of PRCP activity by 10 µM KYP-2047 in an in vitro pharmacological profiling study [69].

In an in vitro assay using pure recombinant human enzymes, we investigated the conversion of Ang II to Ang-(1–7) and detected Ang-(1–7) formation by ACE2 and PRCP. Although some studies reported PREP to convert Ang II to Ang-(1–7) [12,45,46], PREP did not catalyze this conversion into Ang-(1–7) in our setting. This is in line with findings by others that PREP primarily converts Ang I, and not Ang II, directly to Ang-(1–7) [47–49,54]. We confirmed this by using Ang I as a substrate and detected PREP-catalyzed Ang-(1–7) formation, thereby verifying the activity of our recombinant PREP enzyme (S5 Fig). It is noteworthy that in some studies ZPP, a dual PREP and PRCP inhibitor, was chosen. This complicates the interpretation of the reported results as it remains unclear which of the enzymes contributes to the observed effects and to which extend.

ACE2 has been identified as the main enzyme responsible for the conversion of Ang II to Ang-(1–7) in human kidney homogenates [49] and is thus often regarded as the primary enzyme responsible for directing the intrarenal RAS towards the vasodilatory and renoprotective axis by increasing Ang-(1–7) levels. ACE2 deficiency exacerbates kidney injury, while several kidney disease models show reduced renal ACE2 levels, as previously summarized in [17].

In the present study, we assessed the metabolism of Ang II to Ang-(1–7) in hPC cell lysates and on the cell surface, and evaluated the respective contributions of ACE2 and PRCP. On the cell surface, ACE2 activity was found to play a more prominent role in Ang-(1–7) formation than PRCP, especially under control conditions. This is consistent with ACE2 being predominantly localized at the plasma membrane, where it cleaves Ang II in the ultrafiltrate within Bowman’s space [64,70–73]. In hPC lysates, Ang-(1–7) formation was predominantly driven by PRCP, reflecting both its higher expression levels compared to ACE2 and its mainly lysosomal localization, which enables it to cleave and degrade peptides intracellularly [74–76]. Upon FFSS, total Ang-(1–7) formation in supernatants was not altered but the contribution of PRCP in Ang-(1–7) formation was enhanced. This might result from increased enzyme transport to the cell surface through exocytosis or increased enzyme activity due to post-translational modification. Posttranslational modifications of PRCP were reported to include disulfide bonds, N-glycosylation sites, and both a signal sequence and a pro-peptide sequence, which may be cleaved upon maturation or activation [77,78].

In contrast to supernatants, our results reveal an increased conversion of Ang II to Ang-(1–7) in hPC lysates exposed to FFSS which was primarily driven by PRCP. The remaining Ang-(1–7) formation rate despite ACE2 and PRCP inhibition is suggestive of an additional enzyme involved in the conversion of Ang II in hPC lysates contributing to the increased Ang-(1–7) production observed upon FFSS. Additional enzymes involved in the degradation of angiotensin peptides that should be further investigated include Cathepsin A (CTSA; EC 3.4.16.5) and thimet oligopeptidase (THOP1; EC 3.4.24.15). Although CTSA is mainly known to hydrolyze Ang I [79], early studies suggest it may also cleave Ang II to Ang-(1–7) [80]. THOP1 is well established in cleaving Ang I to Ang-(1–7) [81,82], and further studies have indicated that it can also cleave Ang II, albeit inefficiently due to its low catalytic efficiency [83]. However, potential Ang II–derived cleavage products of THOP1 have not yet been identified. Future research might therefore specifically examine THOP1 and Cathepsin A under the experimental conditions in hPC to clarify their potential roles in local RAS regulation and defining more precisely the enzymatic mechanisms underlying Ang II processing in hPC.

In our study, PRCP is the predominant enzymatic contributor to Ang-(1–7) generation in podocyte lysates. PRCP cleaves carboxy-terminal residues from peptides with a penultimate proline, including Ang II and Ang III [78,84–86]. Its relevance for renal homeostasis is supported by findings that renal PRCP expression is reduced in hypertensive rats and restored by losartan [87], and that PRCP depletion in mice induced endothelial dysfunction, hypertension, alterations in renal and vascular eNOS, and increased ROS formation [88]. While FFSS did not directly activate PRCP, this enzymatic capacity provides a plausible mechanism to shift the local balance from Ang II toward Ang-(1–7). This is particularly relevant because Ang-(1–7) is well known to oppose Ang II–mediated injury by exerting vasodilatory, antioxidative, antiproliferative, antifibrotic, and anti-inflammatory actions [17].

Mechanical stress, including FFSS, is known to disrupt podocyte structure and survival by inducing cytoskeletal remodeling, detachment, and apoptosis, partly through mechanotransduction pathways involving Akt–GSK3β–β-catenin and MAPK/ERK signaling [24,89,90]. It has also been reported, that mechanical stress activates the local RAS, enhancing Ang II production and its downstream MAPK, TGF-β, and PI3K cascades, thereby reducing nephrin expression and promoting apoptosis and cytoskeletal destabilization [31,43,91,92]. Given these well-established Ang II-mediated injury mechanisms, a FFSS-associated shift toward Ang-(1–7) could be renoprotective and counterbalance FFSS-induced damage. Moreover, Ang-(1–7) has been demonstrated to directly signal protective in podocytes. It restored nephrin, podocin, and WT1 levels and reduced apoptosis under high-glucose conditions in mouse podocytes. These effects were blocked by the MAS1 receptor antagonist A-779 [93]. In human podocytes exposed to preeclampsia serum, Ang-(1–7) suppressed phosphorylation of p38, ERK1/2, and JNK in a MAS1 receptor-dependent manner, indicating inhibition of stress-activated MAPK pathways [94]. Ang-(1–7) also stimulated modest AT₂R-dependent nitric oxide production [95], a mediator of podocyte redox balance and cytoskeletal regulation [96]. The renoprotective effects of Ang-(1–7) extend to an in vivo mouse model, where cyclic Ang-(1–7) limited albuminuria progression, podocyte dysfunction, and glomerular fibrosis similarly to ACE inhibition [97].

The predominant contribution of PRCP to Ang-(1–7) formation supports the translational hypothesis that PRCP may shape how hPC engage Ang-(1–7)–dependent protective signaling under FFSS, thereby counteracting Ang II– and shear stress–induced injury.

It is well established that Ang-(1–7) signals through the MAS1 receptor, which we were able to detect in hPC via immunofluorescence (S6 Fig). Whether Ang-(1–7) acts in an autocrine manner on hPC or is directly degraded and inactivated remains to be determined. Another potential mechanism involves the internalization of the AT1R along with its ligand Ang II [13,17], followed by the intracellular conversion into Ang-(1–7). This process may exert additional protective effects through the reduction of Ang II levels. Further research is needed to determine if the proposed renoprotective role of PRCP can be functionally demonstrated, for instance by investigating whether PRCP inhibition exacerbates FFSS-induced damage, or whether PRCP overexpression or enhanced activity mitigates FFSS-induced injury.

As a limitation of our study, besides the in vitro setting of cell culture, we had to exogenously add Ang II, as the endogenous concentrations were too low to detect the cleavage pattern by the enzymes investigated. We deliberately chose a high Ang II concentration of 1 µg/mL to evaluate catalytic capacities under standardized conditions avoiding artificial limitations on enzyme activity. Although the exogenously applied Ang II concentration does not reflect physiological in vivo peptide levels, this approach enabled us to use spiked Ang II as a substrate, allowing clear identification of only the spiked product via mass spectrometry and the precise calculation of enzyme activity. Thus, this pharmacological approach contributes to a deeper understanding of Ang II to Ang-(1–7) conversion in human podocytes, particularly in the context of glomerular hyperfiltration. Yet, the molecular mechanisms and potential additional enzymes involved in upregulating Ang-(1–7) formation in response to FFSS need further investigations.

In conclusion, this study contributes to a better understanding of the intrarenal processing of Ang II towards the alternative RAS. Upon FFSS, Ang II to Ang-(1–7) conversion is elevated in hPC lysates which is mainly dependent on PRCP activity. Thus, PRCP may shape how hPC invoke Ang-(1–7)–dependent protective signaling under FFSS, thereby counteracting Ang II– and shear stress–induced injury.

Supporting information

S1 Table. Primer sequences for genes of interest.

https://doi.org/10.1371/journal.pone.0339874.s001

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S1 File. Supplement methods.

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S1 Fig. Characterization of angiotensin system components on RNA level in differentiated hPC.

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S2 Fig. Inhibitor effects on PREP-mediated Ang II to Ang-(1–7) conversion.

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S3 Fig. Inhibition of Ang-(1–7) formation from substrate Ang II by different inhibitors.

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S4 Fig. Western blot analyses of ACE2 and PRCP.

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S5 Fig. Results NEP assay.

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S6 Fig. Representative immunofluorescence image of MAS1.

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S1 Data. Excel-file containing primary LC-MS/MS data.

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S2 Data. Excel-file calculation of delta delta Ct values for gene expression analyses of ACE2 and PRCP.

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S3 Data. Raw images.

https://doi.org/10.1371/journal.pone.0339874.s011

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Acknowledgments

We thankfully acknowledge the contributions of Karen Böhme and Petra Karsten for excellent laboratory assistance. We furthermore thank Moin A. Saleem, University of Bristol, UK for providing hPC, and the Sonnenfeld-Stiftung, Berlin, Germany for providing the FFSS device.

References

  1. 1. Der Sarkissian S, Huentelman MJ, Stewart J, Katovich MJ, Raizada MK. ACE2: A novel therapeutic target for cardiovascular diseases. Prog Biophys Mol Biol. 2006;91(1–2):163–98. pmid:16009403
  2. 2. Inagami T. The renin-angiotensin system. Essays Biochem. 1994;28:147–64. pmid:7925317
  3. 3. Blazek O, Bakris GL. Slowing the Progression of Diabetic Kidney Disease. Cells. 2023;12(15):1975. pmid:37566054
  4. 4. Bhandari S, Mehta S, Khwaja A, Cleland JGF, Ives N, Brettell E, et al. Renin-Angiotensin System Inhibition in Advanced Chronic Kidney Disease. N Engl J Med. 2022;387(22):2021–32. pmid:36326117
  5. 5. DeFronzo RA, Reeves WB, Awad AS. Pathophysiology of diabetic kidney disease: impact of SGLT2 inhibitors. Nat Rev Nephrol. 2021;17(5):319–34. pmid:33547417
  6. 6. Warren AM, Knudsen ST, Cooper ME. Diabetic nephropathy: an insight into molecular mechanisms and emerging therapies. Expert Opin Ther Targets. 2019;23(7):579–91. pmid:31154867
  7. 7. Ruggenenti P, Cravedi P, Remuzzi G. The RAAS in the pathogenesis and treatment of diabetic nephropathy. Nat Rev Nephrol. 2010;6(6):319–30. pmid:20440277
  8. 8. Kourpa A, Schulz A, Mangelsen E, Kaiser-Graf D, Koppers N, Stoll M, et al. Studies in Zebrafish and Rat Models Support Dual Blockade of EP2 and EP4 (Prostaglandin E2 Receptors Type 2 and 4) for Renoprotection in Glomerular Hyperfiltration and Albuminuria. Hypertension. 2023;80(4):771–82. pmid:36715011
  9. 9. Perez-Hernandez J, Olivares MD, Solaz E, Martinez F, Martínez-Hervas S, Pichler G, et al. Urinary podocyte-associated molecules and albuminuria in hypertension. J Hypertens. 2018;36(8):1712–8. pmid:29677049
  10. 10. Benzing T, Salant D. Insights into Glomerular Filtration and Albuminuria. N Engl J Med. 2021;384(15):1437–46. pmid:33852781
  11. 11. Martinez F, Mansego ML, Chaves FJ, Redon J. Genetic bases of urinary albumin excretion and related traits in hypertension. J Hypertens. 2010;28(2):213–25. pmid:20095078
  12. 12. Savoia C, Burger D, Nishigaki N, Montezano A, Touyz RM. Angiotensin II and the vascular phenotype in hypertension. Expert Rev Mol Med. 2011;13:e11. pmid:21450123
  13. 13. Forrester SJ, Booz GW, Sigmund CD, Coffman TM, Kawai T, Rizzo V, et al. Angiotensin II Signal Transduction: An Update on Mechanisms of Physiology and Pathophysiology. Physiol Rev. 2018;98(3):1627–738. pmid:29873596
  14. 14. Zhou M-S, Schulman IH, Raij L. Nitric oxide, angiotensin II, and hypertension. Semin Nephrol. 2004;24(4):366–78. pmid:15252776
  15. 15. Hartmann C, Schulz I, Epe B, Schupp N. Angiotensin II-induced hypertension increases the mutant frequency in rat kidney. Arch Toxicol. 2019;93(7):2045–55. pmid:31098697
  16. 16. Giani JF, Janjulia T, Taylor B, Bernstein EA, Shah K, Shen XZ, et al. Renal generation of angiotensin II and the pathogenesis of hypertension. Curr Hypertens Rep. 2014;16(9):477. pmid:25097114
  17. 17. Lin H, Geurts F, Hassler L, Batlle D, Mirabito Colafella KM, Denton KM, et al. Kidney Angiotensin in Cardiovascular Disease: Formation and Drug Targeting. Pharmacol Rev. 2022;74(3):462–505. pmid:35710133
  18. 18. Crowley SD, Gurley SB, Herrera MJ, Ruiz P, Griffiths R, Kumar AP, et al. Angiotensin II causes hypertension and cardiac hypertrophy through its receptors in the kidney. Proc Natl Acad Sci U S A. 2006;103(47):17985–90. pmid:17090678
  19. 19. Redding KM, Chen BL, Singh A, Re RN, Navar LG, Seth DM, et al. Transgenic mice expressing an intracellular fluorescent fusion of angiotensin II demonstrate renal thrombotic microangiopathy and elevated blood pressure. Am J Physiol Heart Circ Physiol. 2010;298(6):H1807–18. pmid:20363893
  20. 20. Srivastava T, Hariharan S, Alon US, McCarthy ET, Sharma R, El-Meanawy A, et al. Hyperfiltration-mediated Injury in the Remaining Kidney of a Transplant Donor. Transplantation. 2018;102(10):1624–35. pmid:29847501
  21. 21. Srivastava T, Celsi GE, Sharma M, Dai H, McCarthy ET, Ruiz M, et al. Fluid flow shear stress over podocytes is increased in the solitary kidney. Nephrol Dial Transplant. 2014;29(1):65–72. pmid:24166460
  22. 22. Brenner BM. Nephron adaptation to renal injury or ablation. Am J Physiol. 1985;249(3 Pt 2):F324–37. pmid:3898871
  23. 23. Sharma M, Sharma R, McCarthy ET, Savin VJ, Srivastava T. Hyperfiltration-associated biomechanical forces in glomerular injury and response: Potential role for eicosanoids. Prostaglandins Other Lipid Mediat. 2017;132:59–68. pmid:28108282
  24. 24. Friedrich C, Endlich N, Kriz W, Endlich K. Podocytes are sensitive to fluid shear stress in vitro. Am J Physiol Renal Physiol. 2006;291(4):F856–65. pmid:16684926
  25. 25. Mangelsen E, Rothe M, Schulz A, Kourpa A, Panáková D, Kreutz R, et al. Concerted EP2 and EP4 Receptor Signaling Stimulates Autocrine Prostaglandin E2 Activation in Human Podocytes. Cells. 2020;9(5):1256. pmid:32438662
  26. 26. Srivastava T, McCarthy ET, Sharma R, Cudmore PA, Sharma M, Johnson ML, et al. Prostaglandin E(2) is crucial in the response of podocytes to fluid flow shear stress. J Cell Commun Signal. 2010;4(2):79–90. pmid:20531983
  27. 27. Yang SH, Choi JW, Huh D, Jo HA, Kim S, Lim CS, et al. Roles of fluid shear stress and retinoic acid in the differentiation of primary cultured human podocytes. Exp Cell Res. 2017;354(1):48–56. pmid:28320523
  28. 28. Srivastava T, Heruth DP, Duncan RS, Rezaiekhaligh MH, Garola RE, Priya L, et al. Transcription Factor β-Catenin Plays a Key Role in Fluid Flow Shear Stress-Mediated Glomerular Injury in Solitary Kidney. Cells. 2021;10(5):1253. pmid:34069476
  29. 29. Blaine J, Dylewski J. Regulation of the Actin Cytoskeleton in Podocytes. Cells. 2020;9(7):1700. pmid:32708597
  30. 30. Pavenstädt H, Kriz W, Kretzler M. Cell biology of the glomerular podocyte. Physiol Rev. 2003;83(1):253–307. pmid:12506131
  31. 31. Durvasula RV, Petermann AT, Hiromura K, Blonski M, Pippin J, Mundel P, et al. Activation of a local tissue angiotensin system in podocytes by mechanical strain. Kidney Int. 2004;65(1):30–9. pmid:14675034
  32. 32. Stitt-Cavanagh E, MacLeod L, Kennedy C. The podocyte in diabetic kidney disease. ScientificWorldJournal. 2009;9:1127–39. pmid:19838599
  33. 33. Jefferson JA, Shankland SJ, Pichler RH. Proteinuria in diabetic kidney disease: a mechanistic viewpoint. Kidney Int. 2008;74(1):22–36. pmid:18418356
  34. 34. Berg UB, Torbjörnsdotter TB, Jaremko G, Thalme B. Kidney morphological changes in relation to long-term renal function and metabolic control in adolescents with IDDM. Diabetologia. 1998;41(9):1047–56. pmid:9754823
  35. 35. Gao N, Wang H, Yin H, Yang Z. Angiotensin II induces calcium-mediated autophagy in podocytes through enhancing reactive oxygen species levels. Chem Biol Interact. 2017;277:110–8. pmid:28916337
  36. 36. Cardoso VG, Gonçalves GL, Costa-Pessoa JM, Thieme K, Lins BB, Casare FAM, et al. Angiotensin II-induced podocyte apoptosis is mediated by endoplasmic reticulum stress/PKC-δ/p38 MAPK pathway activation and trough increased Na+/H+ exchanger isoform 1 activity. BMC Nephrol. 2018;19(1):179. pmid:30005635
  37. 37. Zhao M, Bai M, Ding G, Zhang Y, Huang S, Jia Z, et al. Angiotensin II Stimulates the NLRP3 Inflammasome to Induce Podocyte Injury and Mitochondrial Dysfunction. Kidney Dis (Basel). 2018;4(2):83–94. pmid:29998123
  38. 38. Che G, Gao H, Hu Q, Xie H, Zhang Y. Angiotensin II promotes podocyte injury by activating Arf6-Erk1/2-Nox4 signaling pathway. PLoS One. 2020;15(3):e0229747. pmid:32119711
  39. 39. Anderson M, Roshanravan H, Khine J, Dryer SE. Angiotensin II activation of TRPC6 channels in rat podocytes requires generation of reactive oxygen species. J Cell Physiol. 2014;229(4):434–42. pmid:24037962
  40. 40. Hsu H-H, Hoffmann S, Endlich N, Velic A, Schwab A, Weide T, et al. Mechanisms of angiotensin II signaling on cytoskeleton of podocytes. J Mol Med (Berl). 2008;86(12):1379–94. pmid:18773185
  41. 41. Campbell KN, Raij L, Mundel P. Role of angiotensin II in the development of nephropathy and podocytopathy of diabetes. Curr Diabetes Rev. 2011;7(1):3–7. pmid:21067505
  42. 42. Márquez E, Riera M, Pascual J, Soler MJ. Renin-angiotensin system within the diabetic podocyte. Am J Physiol Renal Physiol. 2015;308(1):F1–10. pmid:25339703
  43. 43. Kang JS, Lee SJ, Lee J-H, Kim J-H, Son SS, Cha S-K, et al. Angiotensin II-mediated MYH9 downregulation causes structural and functional podocyte injury in diabetic kidney disease. Sci Rep. 2019;9(1):7679. pmid:31118506
  44. 44. Grobe N, Elased KM, Cool DR, Morris M. Mass spectrometry for the molecular imaging of angiotensin metabolism in kidney. Am J Physiol Endocrinol Metab. 2012;302(8):E1016–24. pmid:22318946
  45. 45. Serfozo P, Wysocki J, Gulua G, Schulze A, Ye M, Liu P, et al. Ang II (Angiotensin II) Conversion to Angiotensin-(1-7) in the Circulation Is POP (Prolyloligopeptidase)-Dependent and ACE2 (Angiotensin-Converting Enzyme 2)-Independent. Hypertension. 2020;75(1):173–82. pmid:31786979
  46. 46. Santos RAS, Ferreira AJ, Simões E Silva AC. Recent advances in the angiotensin-converting enzyme 2-angiotensin(1-7)-Mas axis. Exp Physiol. 2008;93(5):519–27. pmid:18310257
  47. 47. Kaltenecker CC, Domenig O, Kopecky C, Antlanger M, Poglitsch M, Berlakovich G, et al. Critical Role of Neprilysin in Kidney Angiotensin Metabolism. Circ Res. 2020;127(5):593–606. pmid:32418507
  48. 48. Kovarik JJ, Kaltenecker CC, Kopecky C, Domenig O, Antlanger M, Werzowa J, et al. Intrarenal Renin-Angiotensin-System Dysregulation after Kidney Transplantation. Sci Rep. 2019;9(1):9762. pmid:31278281
  49. 49. Domenig O, Manzel A, Grobe N, Königshausen E, Kaltenecker CC, Kovarik JJ, et al. Neprilysin is a Mediator of Alternative Renin-Angiotensin-System Activation in the Murine and Human Kidney. Sci Rep. 2016;6:33678. pmid:27649628
  50. 50. Karnik SS, Singh KD, Tirupula K, Unal H. Significance of angiotensin 1-7 coupling with MAS1 receptor and other GPCRs to the renin-angiotensin system: IUPHAR Review 22. Br J Pharmacol. 2017;174(9):737–53. pmid:28194766
  51. 51. Gava E, Samad-Zadeh A, Zimpelmann J, Bahramifarid N, Kitten GT, Santos RA, et al. Angiotensin-(1-7) activates a tyrosine phosphatase and inhibits glucose-induced signalling in proximal tubular cells. Nephrol Dial Transplant. 2009;24(6):1766–73. pmid:19144997
  52. 52. Dilauro M, Zimpelmann J, Robertson SJ, Genest D, Burns KD. Effect of ACE2 and angiotensin-(1-7) in a mouse model of early chronic kidney disease. Am J Physiol Renal Physiol. 2010;298(6):F1523–32. pmid:20357030
  53. 53. Calò LA, Schiavo S, Davis PA, Pagnin E, Mormino P, D’Angelo A, et al. ACE2 and angiotensin 1-7 are increased in a human model of cardiovascular hyporeactivity: pathophysiological implications. J Nephrol. 2010;23(4):472–7. pmid:20349406
  54. 54. Jiang F, Yang J, Zhang Y, Dong M, Wang S, Zhang Q, et al. Angiotensin-converting enzyme 2 and angiotensin 1-7: novel therapeutic targets. Nat Rev Cardiol. 2014;11(7):413–26. pmid:24776703
  55. 55. Montezano AC, Nguyen Dinh Cat A, Rios FJ, Touyz RM. Angiotensin II and vascular injury. Curr Hypertens Rep. 2014;16(6):431. pmid:24760441
  56. 56. Saleem MA, O’Hare MJ, Reiser J, Coward RJ, Inward CD, Farren T, et al. A conditionally immortalized human podocyte cell line demonstrating nephrin and podocin expression. J Am Soc Nephrol. 2002;13(3):630–8. pmid:11856766
  57. 57. Ni L, Saleem M, Mathieson PW. Podocyte culture: tricks of the trade. Nephrology (Carlton). 2012;17(6):525–31. pmid:22591222
  58. 58. Schulz A, Müller NV, van de Lest NA, Eisenreich A, Schmidbauer M, Barysenka A, et al. Analysis of the genomic architecture of a complex trait locus in hypertensive rat models links Tmem63c to kidney damage. Elife. 2019;8:e42068. pmid:30900988
  59. 59. Livak KJ, Schmittgen TD. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) Method. Methods. 2001;25(4):402–8. pmid:11846609
  60. 60. Burns KD. Angiotensin II and its receptors in the diabetic kidney. Am J Kidney Dis. 2000;36(3):449–67. pmid:10977776
  61. 61. Lee HS. Mechanisms and consequences of TGF-ß overexpression by podocytes in progressive podocyte disease. Cell Tissue Res. 2012;347(1):129–40. pmid:21541658
  62. 62. Del Prete D, Gambaro G, Lupo A, Anglani F, Brezzi B, Magistroni R, et al. Precocious activation of genes of the renin-angiotensin system and the fibrogenic cascade in IgA glomerulonephritis. Kidney Int. 2003;64(1):149–59. pmid:12787405
  63. 63. Liebau MC, Lang D, Böhm J, Endlich N, Bek MJ, Witherden I, et al. Functional expression of the renin-angiotensin system in human podocytes. Am J Physiol Renal Physiol. 2006;290(3):F710–9. pmid:16189286
  64. 64. Ye M, Wysocki J, William J, Soler MJ, Cokic I, Batlle D. Glomerular localization and expression of Angiotensin-converting enzyme 2 and Angiotensin-converting enzyme: implications for albuminuria in diabetes. J Am Soc Nephrol. 2006;17(11):3067–75. pmid:17021266
  65. 65. Velez JCQ, Bland AM, Arthur JM, Raymond JR, Janech MG. Characterization of renin-angiotensin system enzyme activities in cultured mouse podocytes. Am J Physiol Renal Physiol. 2007;293(1):F398–407. pmid:17429035
  66. 66. Castrop H, Höcherl K, Kurtz A, Schweda F, Todorov V, Wagner C. Physiology of kidney renin. Physiol Rev. 2010;90(2):607–73. pmid:20393195
  67. 67. Matsusaka T, Niimura F, Shimizu A, Pastan I, Saito A, Kobori H, et al. Liver angiotensinogen is the primary source of renal angiotensin II. J Am Soc Nephrol. 2012;23(7):1181–9. pmid:22518004
  68. 68. Matsusaka T, Niimura F, Pastan I, Shintani A, Nishiyama A, Ichikawa I. Podocyte injury enhances filtration of liver-derived angiotensinogen and renal angiotensin II generation. Kidney Int. 2014;85(5):1068–77. pmid:24284520
  69. 69. Jalkanen AJ, Piepponen TP, Hakkarainen JJ, De Meester I, Lambeir A-M, Forsberg MM. The effect of prolyl oligopeptidase inhibition on extracellular acetylcholine and dopamine levels in the rat striatum. Neurochem Int. 2012;60(3):301–9. pmid:22210165
  70. 70. Donoghue M, Hsieh F, Baronas E, Godbout K, Gosselin M, Stagliano N, et al. A novel angiotensin-converting enzyme-related carboxypeptidase (ACE2) converts angiotensin I to angiotensin 1-9. Circ Res. 2000;87(5):E1–9. pmid:10969042
  71. 71. Wysocki J, Batlle D. ACE2 in the Urine: Where Does It Come From? Kidney360. 2022;3(12):2001–4. pmid:36591363
  72. 72. Clarke NE, Turner AJ. Angiotensin-converting enzyme 2: the first decade. Int J Hypertens. 2012;2012:307315. pmid:22121476
  73. 73. Rouaud F, Méan I, Citi S. The ACE2 Receptor for Coronavirus Entry Is Localized at Apical Cell-Cell Junctions of Epithelial Cells. Cells. 2022;11(4):627. pmid:35203278
  74. 74. De Hert E, Bracke A, Lambeir A-M, Van der Veken P, De Meester I. The C-terminal cleavage of angiotensin II and III is mediated by prolyl carboxypeptidase in human umbilical vein and aortic endothelial cells. Biochem Pharmacol. 2021;192:114738. pmid:34418354
  75. 75. Kehoe K, Brouns R, Verkerk R, Engelborghs S, De Deyn PP, Hendriks D, et al. Prolyl carboxypeptidase activity decline correlates with severity and short-term outcome in acute ischemic stroke. Neurochem Res. 2015;40(1):81–8. pmid:25370794
  76. 76. Grobe N, Leiva O, Morris M, Elased KM. Loss of prolyl carboxypeptidase in two-kidney, one-clip goldblatt hypertensive mice. PLoS One. 2015;10(2):e0117899. pmid:25706121
  77. 77. Soisson SM, Patel SB, Abeywickrema PD, Byrne NJ, Diehl RE, Hall DL, et al. Structural definition and substrate specificity of the S28 protease family: the crystal structure of human prolylcarboxypeptidase. BMC Struct Biol. 2010;10:16. pmid:20540760
  78. 78. Mallela J, Yang J, Shariat-Madar Z. Prolylcarboxypeptidase: a cardioprotective enzyme. Int J Biochem Cell Biol. 2009;41(3):477–81. pmid:18396440
  79. 79. Jackman HL, Massad MG, Sekosan M, Tan F, Brovkovych V, Marcic BM, et al. Angiotensin 1-9 and 1-7 release in human heart: role of cathepsin A. Hypertension. 2002;39(5):976–81. pmid:12019279
  80. 80. Grynbaum A, Marks N. Characterization of a rat brain catheptic carboxypeptidase (cathepsin A) inactivating angiotensin-II. J Neurochem. 1976;26(2):313–8. pmid:1255194
  81. 81. Grobe N, Weir NM, Leiva O, Ong FS, Bernstein KE, Schmaier AH, et al. Identification of prolyl carboxypeptidase as an alternative enzyme for processing of renal angiotensin II using mass spectrometry. Am J Physiol Cell Physiol. 2013;304(10):C945–53. pmid:23392115
  82. 82. Rioli V, Kato A, Portaro FC, Cury GK, te Kaat K, Vincent B, et al. Neuropeptide specificity and inhibition of recombinant isoforms of the endopeptidase 3.4.24.16 family: comparison with the related recombinant endopeptidase 3.4.24.15. Biochem Biophys Res Commun. 1998;250(1):5–11. pmid:9735321
  83. 83. Rioli V, Gozzo FC, Heimann AS, Linardi A, Krieger JE, Shida CS, et al. Novel natural peptide substrates for endopeptidase 24.15, neurolysin, and angiotensin-converting enzyme. J Biol Chem. 2003;278(10):8547–55. pmid:12500972
  84. 84. Odya CE, Marinkovic DV, Hammon KJ, Stewart TA, Erdös EG. Purification and properties of prolylcarboxypeptidase (angiotensinase C) from human kidney. J Biol Chem. 1978;253(17):5927–31. pmid:28321
  85. 85. Shariat-Madar Z, Mahdi F, Schmaier AH. Recombinant prolylcarboxypeptidase activates plasma prekallikrein. Blood. 2004;103(12):4554–61. pmid:14996700
  86. 86. Wallingford N, Perroud B, Gao Q, Coppola A, Gyengesi E, Liu Z-W, et al. Prolylcarboxypeptidase regulates food intake by inactivating alpha-MSH in rodents. J Clin Invest. 2009;119(8):2291–303. pmid:19620781
  87. 87. Qin X-P, Zeng S-Y, Tian H-H, Deng S-X, Ren J-F, Zheng Y-B, et al. Involvement of prolylcarboxypeptidase in the effect of rutaecarpine on the regression of mesenteric artery hypertrophy in renovascular hypertensive rats. Clin Exp Pharmacol Physiol. 2009;36(3):319–24. pmid:19018804
  88. 88. Adams GN, LaRusch GA, Stavrou E, Zhou Y, Nieman MT, Jacobs GH, et al. Murine prolylcarboxypeptidase depletion induces vascular dysfunction with hypertension and faster arterial thrombosis. Blood. 2011;117(14):3929–37. pmid:21297000
  89. 89. Srivastava T, Dai H, Heruth DP, Alon US, Garola RE, Zhou J, et al. Mechanotransduction signaling in podocytes from fluid flow shear stress. Am J Physiol Renal Physiol. 2018;314(1):F22–34. pmid:28877882
  90. 90. Reynolds PA. The mechanobiology of kidney podocytes in health and disease. Clin Sci (Lond). 2020;134(11):1245–53. pmid:32501496
  91. 91. Durvasula RV, Shankland SJ. The renin-angiotensin system in glomerular podocytes: mediator of glomerulosclerosis and link to hypertensive nephropathy. Curr Hypertens Rep. 2006;8(2):132–8. pmid:16672146
  92. 92. Du S-J, Huang W, Hao Y, Zhang C, He F-F. Mechanical Stress and Protective Mechanisms in Podocytes: Insights into Hypertensive Nephropathy. Int J Mol Sci. 2025;26(19):9316. pmid:41096586
  93. 93. Lu J, Chen G, Shen G, Ouyang W. Ang-(1-7) attenuates podocyte injury induced by high glucose in vitro. Arch Endocrinol Metab. 2023;67(6):e000643. pmid:37364145
  94. 94. Tian J, Zhang L, Zhou Y, Xiao J, Li S, Chen Y, et al. Angiotensin-(1-7) attenuates damage to podocytes induced by preeclamptic serum through MAPK pathways. Int J Mol Med. 2014;34(4):1057–64. pmid:25092178
  95. 95. Semenikhina M, Bohovyk R, Fedoriuk M, Stefanenko M, Klemens CA, Oates JC, et al. Renin-angiotensin system-mediated nitric oxide signaling in podocytes. Am J Physiol Renal Physiol. 2024;327(3):F532–42. pmid:39024356
  96. 96. Semenikhina M, Stefanenko M, Spires DR, Ilatovskaya DV, Palygin O. Nitric-Oxide-Mediated Signaling in Podocyte Pathophysiology. Biomolecules. 2022;12(6):745. pmid:35740870
  97. 97. Cassis P, Locatelli M, Corna D, Villa S, Rottoli D, Cerullo D, et al. Addition of cyclic angiotensin-(1-7) to angiotensin-converting enzyme inhibitor therapy has a positive add-on effect in experimental diabetic nephropathy. Kidney Int. 2019;96(4):906–17. pmid:31307778